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4,832 | https://bio-protocol.org/en/bpdetail?id=4832&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Testing for Allele-specific Expression from Human Brain Samples
MD Maria E. Diaz-Ortiz
NJ Nimansha Jain
MG Michael D. Gallagher
MP Marijan Posavi
TU Travis L. Unger
AC Alice S. Chen-Plotkin
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4832 Views: 528
Reviewed by: Geoffrey C. Y. LauZhengrong YuanUte Angelika Hoffmann
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Original Research Article:
The authors used this protocol in Science Aug 2022
Abstract
Many single nucleotide polymorphisms (SNPs) identified by genome-wide association studies exert their effects on disease risk as expression quantitative trait loci (eQTL) via allele-specific expression (ASE). While databases for probing eQTLs in tissues from normal individuals exist, one may wish to ascertain eQTLs or ASE in specific tissues or disease-states not characterized in these databases. Here, we present a protocol to assess ASE of two possible target genes (GPNMB and KLHL7) of a known genome-wide association study (GWAS) Parkinson’s disease (PD) risk locus in postmortem human brain tissue from PD and neurologically normal individuals. This was done using a sequence of RNA isolation, cDNA library generation, enrichment for transcripts of interest using customizable cDNA capture probes, paired-end RNA sequencing, and subsequent analysis. This method provides increased sensitivity relative to traditional bulk RNAseq-based and a blueprint that can be extended to the study of other genes, tissues, and disease states.
Key features
• Analysis of GPNMB allele-specific expression (ASE) in brain lysates from cognitively normal controls (NC) and Parkinson’s disease (PD) individuals.
• Builds on the ASE protocol of Mayba et al. (2014) and extends application from cells to human tissue.
• Increased sensitivity by enrichment for desired transcript via RNA CaptureSeq (Mercer et al., 2014).
• Optimized for human brain lysates from cingulate gyrus, caudate nucleus, and cerebellum.
Graphical overview
Keywords: Allele-specific expression (ASE) RNA CaptureSeq Expression quantitative trait locus (eQTL) Human brain Neurodegeneration
Background
Most single nucleotide polymorphisms (SNPs) associated with disease traits by genome-wide association studies (GWAS) are in non-coding regions of the genome. One way in which these SNPs may influence disease risk is via effects on expression of nearby genes as expression quantitative trait loci (eQTL). Public resources such as the Genotype-Tissue Expression (GTEx) Project allow scientists to easily query large datasets from multiple tissues from humans without disease for eQTL relationships between SNP variants and specific RNA transcripts in various organ tissues from healthy individuals. However, in many cases, investigators may want to understand whether an eQTL relationship extends to specific tissues from specific individuals; for example, either in tissues not queried by these databases or in individuals with a disease condition. Because the mechanistic basis for many eQTL effects is allele-specific expression (ASE), we developed a protocol for assaying ASE in brain tissue from human subjects with Parkinson’s disease (PD) who are heterozygous for the eQTL SNP in question, affecting the target gene GPNMB. This protocol builds on the ASE method described by Mayba et al. (2014) by enriching for the desired transcripts via an RNA CaptureSeq (Mercer et al., 2014) step and by adapting the protocol to assay human brain tissue as opposed to cultured cells. Furthermore, our protocol can be adapted to multiple complex traits and tissues to advance our mechanistic understanding of many non-coding risk variants with eQTL effects.
Materials and reagents
Biological materials
Human brain samples (acquired from University of Pennsylvania’s CNDR Brain Bank)
Reagents
TriZol (Invitrogen, catalog number: 15596026)
Qiagen RNeasy kit (Qiagen, catalog number: 74104)
Agilent Bioanalyzer RNA 6000 Nano Kit (Agilent, catalog number: 5067-1511)
Agilent Bioanalyzer DNA 1000 Nano (Agilent, catalog number: 5067-1504)
Agencourt AMPure XP beads (Agencourt, catalog number: A63881)
KAPA RNA HyperPrep Kit (KAPA/Roche Sequencing solutions, catalog number: KK8540)
Roche SeqCap RNA probes (Roche, custom designed, see Procedure below)
Roche SeqCap EZ Accessory kit v2 (Roche, catalog number: 07145594001)
Roche SeqCap EZ Hybridization and Wash Kit (Roche, catalog number: 05634261001)
Ethanol (Decon Labs, Inc., catalog number: 2716)
Water for RNA applications (Fisher bioreagents, catalog number: BP5611)
Laboratory supplies
1.5 mL plastic tubes (Eppendorf, catalog number: 022431021)
0.2 mL PCR tubes (Denville, catalog number: C18098-4)
Equipment
Bioanalyzer (Agilent 2100 Bioanalyzer, catalog number: G2939BA)
Thermocycler (Applied Biosystems, Veriti 96-well Thermal Cycler, catalog number: 4375786)
Sequencer (Illumina HiSeq 2500, catalog number: SY-401-2501)
Dyna-mag2 magnet (Thermo Fisher, catalog number: 12321D)
Vortex mixer (Fisherbrand, Brand, catalog number: 02-215-365)
Microcentrifuge (Eppendorf Model 5424, catalog number: 022620401)
NanoDrop or similar spectrophotometer
Software and datasets
RStudio
FastQC
Trimmomatic (Version 032)
STAR
WASP
VariantAnnotation
TxDb.Hsapiens.UCSC.hg19.knownGene
Procedure
RNA isolation from human brain samples
Dissect human postmortem brain samples as previously described (Chen-Plotkin et al., 2008). In our publication (Diaz-Ortiz et al., 2022), we used neurologically normal controls (NC, n = 2) and PD (n = 4) individuals, and we dissected the caudate nucleus, cingulate gyrus, and cerebellum.
Isolate total RNA from postmortem brain samples using TRIzol and RNeasy Mini columns as previously described (Chen-Plotkin et al., 2008).
Assess RNA concentration and purity by spectrophotometric measurement of 260/280 nm OD ratios using a NanoDrop.
Note: We require 260/280 ratios between 1.90 and 2.10 (for ratios outside this range, see General notes and troubleshooting).
Assess RNA integrity by capillary electrophoresis on an Agilent 2100 Bioanalyzer following the RNA 6000 Nano Kit guide.
Note: While RNA Integrity Numbers (RINs) of > 7 are desired for most applications, this may not be possible in all cases; our RINs varied between 5.2 and 9.0 (average 6.5). In cases where RIN < 6.5, visual inspection of Bioanalyzer traces is encouraged to ascertain the etiology (see General notes and troubleshooting). Example Agilent 2100 Bioanalyzer traces with corresponding RINs are shown in Figure 1.
Figure 1. Example Bioanalyzer traces of RNA samples with various RNA integrity numbers (RINs), specifically RIN = 9.0 (A), RIN = 5.2 (B), and RIN = 3.2 (C). While both B and C have relatively low RIN numbers, the wide second peak from left to right in trace B is consistent with the presence of small RNAs, while the absence of four distinct peaks in trace C is consistent with true RNA degradation.
Aliquot RNA isolates and store at -80 °C until use.
Note: We prepared multiple 20 μL aliquots from each brain sample isolated, so that each aliquot can be single used to minimize freeze-thaw effects, which can affect RNA quality and downstream steps.
Detailed description of sections B–G are adapted from the Roche SeqCap RNA Enrichment System User Guide.
cDNA Library preparation
Prepare libraries with the KAPA RNA HyperPrep Kit per the Roche SeqCap RNA Enrichment System User Guide. An overview of the procedure is provided below:
Add spike in controls.
Spike-in ERCC controls into 100 ng of total RNA following manufacturer’s instructions.
Adjust the volume of the spiked RNA sample to a total volume of 10 μL in PCR-grade water.
Fragment RNA into 100–200 bp fragments.
Mix 10 μL of spiked sample with 10 μL of 2× Fragment, Prime, and Elute Buffer on ice by pipetting up and down 10 times.
Place samples from step B2a in a thermocycler and run for 8 min at 94 °C.
Quickly place samples on ice for next step.
Perform first-cDNA-strand synthesis.
Prepare First Strand Master Mix on ice by mixing 11 μL of 1st Strand Synthesis Buffer and 1 μL of KAPA Script per sample.
Mix 10 μL of the First Strand Master Mix with 20 μL of fragmented RNA sample on ice by gently pipetting up and down several times.
Place samples from step B3b in a thermocycler and run for 10 min at 25 °C, 15 min at 42 °C, and 15 min at 70 °C, followed by a 4 °C hold until ready for next step.
Perform second strand synthesis.
Prepare Second Strand Master Mix on ice by mixing 31 μL of 2nd Strand Marking Buffer with 2 μL of 2nd Strand Synthesis Enzyme Mix per sample (for a total volume of 33 μL, which will be 10% in excess of the 30 μL needed for the next step).
Mix 30 μL of 1st strand cDNA (samples from step B3c) with 30 μL of the 2nd Strand Master Mix on ice by gently pipetting up and down several times.
Place samples from step B4b in a thermocycler and run for 30–60 min at 16 °C followed by a 4 °C hold until ready for next step.
Clean up double-stranded cDNA.
Mix 60 μL of double-stranded cDNA samples from step B4c with 108 μL of Agencourt AMPure XP beads (prewarmed to room temperature for 30 min and vortexed to ensure a homogeneous suspension) by gently pipetting up and down several times.
Incubate the tube at room temperature for 15 min to allow the cDNA to bind to the beads.
Place the tube on a dyna-mag2 magnet to capture the beads and incubate until the liquid is clear.
Carefully remove and discard the supernatant using a P200 pipette.
Keeping the tube on the magnet, add 200 μL of freshly prepared 80% ethanol.
Incubate the tube at room temperature for ≥ 30 s.
Carefully remove and discard the ethanol.
Repeat steps B5e–B5g.
Allow the beads to dry at room temperature until all ethanol evaporates. Visually inspect for any remaining drops of ethanol in the tube. The sample is sufficiently dry when no drops are left. Caution: Over-drying the beads (for longer than 3–5 min) may result in dramatic yield loss.
Remove the tubes from the magnet.
Perform A-tailing.
Prepare A-tailing Master Mix by mixing 24 μL of PCR-grade water, 3 μL of 10× KAPA A-Tailing Buffer, and 3 μL of KAPA A-Tailing Enzyme.
For each sample, resuspend beads from step B5i in 30 μL of A-tailing master mix by pipetting up and down.
Place samples in a thermocycler and run for 30 min at 30 °C and 20–30 min at 60 °C followed by a 4 °C hold until ready for next step.
Ligate adapters.
Generate a 700 nM adapter working dilution for each adapter ligation reaction.
Note: Adapter concentration should be adjusted for input RNA amount other than 100 ng per user guide appendix F.
Prepare Adapter Ligation Mix by mixing 16 μL of PCR-grade water, 14 μL of 5× KAPA Ligation Buffer, and 5 μL of KAPA T4 DNA Ligase per reaction.
Mix each 30 μL of A-tailing reaction from step B6c with 35 μL of the Adapter Ligation Master Mix by pipetting up and down.
To each sample from step B7c, add 5 μL of the SeqCap Library Adapter working dilution (with the desired Index) and mix by gently pipetting up and down 10 times.
Note: Record the index used for each sample.
Incubate at 20 °C for 15 min.
First post ligation cleanup.
To each 70 μL of adapter ligation reaction, add 70 μL of thawed PEG/NaCl solution, resulting in a total volume of 140 μL, and mix by pipetting up and down several times.
Incubate the tube at room temperature for 15 min to allow the cDNA to bind to the beads.
Place the tube on a magnet to capture the beads. Incubate until the liquid is clear.
Carefully remove and discard 135 μL of supernatant.
Repeat steps B5e–B5j (wash twice with 200 μL of freshly prepared 80% EtOH, then allow to dry at room temperature, as described in step B5).
Thoroughly resuspend the beads in 50 μL of elution buffer (10 mM Tris-HCl, pH 8.0).
Incubate the tube at room temperature for 2 min to allow the cDNA to elute off the beads.
Second post ligation cleanup.
To each 50 μL of resuspended cDNA with beads, add 50 μL of thawed PEG/NaCl solution and mix thoroughly whilst avoiding bubbles by pipetting up and down multiple times.
Incubate the tube at room temperature for 15 min to allow the cDNA to bind to the beads.
Place the tube on a magnet to capture the beads. Incubate until the liquid is clear.
Carefully remove and discard 95 μL of supernatant.
Repeat steps B5e–B5j (wash twice with 200 μL of freshly prepared 80% EtOH, then allow to dry at room temperature, as described in step B5).
Thoroughly resuspend the beads in 22.5 μL of elution buffer (10 mM Tris-HCl, pH 8.0) by vortexing briefly.
Incubate the tube at room temperature for 2 min to allow the cDNA to elute off the beads.
Place the tube on dyna-mag2 magnet to capture the beads and incubate until the liquid is clear.
Transfer 20 μL of the clear supernatant to a new 0.2 mL PCR tube.
Note: Samples can be stored at 4 °C for up to one week or at -20 °C for up to one month.
Pre-capture library amplification by LM-PCR
Prepare the Pre-Capture LM-PCR Master Mix on ice by mixing 25 μL of 2× KAPA HiFi HotStart ReadyMix and 5 μL of 10× KAPA Library Amplification Primer Mix per sample, pipetting up and down 10 times.
Mix 20 μL of sample generated in section B with 30 μL of Pre-Capture LM-PCR Master Mix on ice by pipetting up and down several times.
Transfer samples to a thermocycler and run at the following settings: 45 s at 98 °C; 11 cycles of 15 s at 98 °C, 30 s at 60 °C, and 30 s at 72 °C; 5 min at 72 °C; and hold at 4 °C.
Purify the amplified samples with Agencourt AMPure XP beads by mixing 1:1 volumes of amplified samples to beads and then following steps outlined in section B, step 5.
Note: Alternatively, use Qiagen QIAquick PCR Purification Kit.
Assess DNA purity by spectrophotometric measurement of 260/280 nm OD ratios using a NanoDrop.
Note: 260/280 ratios between 1.7 and 2.0 are recommended (for ratios outside this range, see General notes and troubleshooting).
Analyze DNA library quality on a Bioanalyzer using Agilent DNA 1000 chip following the Agilent 2100 Kit user guide.
Note: The average fragment size should be between 150 and 500 bp.
SeqCap RNA ChoiceTM probe pool design
Design SeqCap RNA probe pools, using Roche Sequencing Solutions Custom Design (Roche, WI).
We designed probes of approximately 60 bp in length with no more than 20 close matches in the genome, as determined by the Sequence Search and Alignment by Hashing Algorithm (SSAHA). The goal was to provide sufficient coverage of transcripts of interest (GPNMB, KLHL7) while minimizing potential off-target effects. A close match is defined as any genomic sequence that differs from one of the probe sequences by five or fewer single-base insertions, deletions, or substitutions. Select exonic probes that cover the transcript(s) of interest (here, GPNMB and KLHL7). While single mismatches within a 60 bp probe are typically not thought to affect hybridization dynamics, as a conservative measure to minimize SNP-mediated capture bias, we ensured that probes do not overlap any SNPs in linkage disequilibrium (r2 > 0.2) with the sentinel SNP (here, rs199347).
Target cDNA enrichment
Perform target cDNA enrichment and sequencing as previously described (Mercer et al., 2014) using the SeqCap EZ Accessory kit v2, as summarized below:
Hybridize samples to SeqCap RNA probe pools.
Thaw 4.5 μL of SeqCap RNA probe pool aliquots (one per sample library, see user guide for instructions on how to generate these) on ice.
Spin down the lyophilized SeqCap HE Universal and SeqCap HE Index Oligos, then resuspend in PCR-grade water to 1 mM and vortex to mix. Combine the HE oligos such that the resulting HE oligo pool contains 50% SeqCap HE Universal Oligo and 50% of a mixture of the appropriate SeqCap HE Index oligos, for a combined amount of 2,000 pmol that is required for a single sequence capture experiment.
Combine equimolar amounts of cDNA libraries (already barcoded) from each brain sample for a total mass of 1 μg.
In a 1.5 mL tube, combine 5 μL of human COT DNA (control DNA included in SeqCap accessory kit v2), 1 μg of pooled cDNA from previous step, and 2,000 pmol of HE oligo pool.
Dry the samples from section E step 1d in a vacuum concentrator or at high heat at 60 °C and then resuspend in 7.5 μL of 2× Hybridization buffer with 3 μL of hybridization component A by vortexing and then spinning down for 10 s at maximum speed.
Place samples in a pre-warmed 95 °C heat block for 10 min to denature cDNA; then, centrifuge for 10 s at maximum speed.
Transfer 10.5 μL of sample to 4.5 μL aliquot of SeqCap RNA probe pool in a 0.2 mL PCR tube, vortex for 3 s, and spin down at maximum speed for 10 s.
Finally, hybridize the pooled cDNA library to SeqCap RNA probes by running in a thermocycler at 47 °C (lid temperature at 57 °C) for 16–20 h.
Wash and recover the captured multiplex cDNA sample.
Prepare the captured multiplex cDNA samples from section E step 1h by first ligating them to pre-washed capture beads (100 μL of beads per capture reaction in 0.2 mL tubes) in a thermocycler for 45 min at 47 °C with the lid at 57 °C.
Perform multiple washes, as specified in the Roche SeqCap RNA Enrichment System User’s Guide using the Dyna-mag2 magnet: once with 100 μL of 47 °C wash buffer I, twice with 200 μL of 47 °C stringent wash buffer, once with 200 μL of room temperature wash buffer I, once with 200 μL of room temperature wash buffer II, and once with 200 μL of room temperature wash buffer III).
Finally, resuspend the beads ligated to captured cDNA libraries in 50 μL of PCR-grade water.
Note: Samples can be stored at -20 °C until ready for post-ligation LM-PCR.
Post-capture library amplification by LM-PCR
Prepare the Post-Capture LM-PCR Master Mix on ice by mixing 25 μL of 2× KAPA HiFi HotStart ReadyMix and 5 μL of Post-LM-PCR Oligos 1 & 2 (5 μM) per reaction.
Mix 20 μL of bead-bound captured cDNA (vortexed to ensure homogeneous suspension) or 20 μL of PCR-grade water (for negative control) with 30 μL of Post-Capture LM-PCR Master Mix.
Transfer samples to a thermocycler and run at the following settings: 45 s at 98 °C; 14 cycles of 15 s at 98 °C, 30 s at 60 °C, and 30 s at 72 °C; 5 min at 72 °C; and hold at 4 °C.
Purify the amplified samples with Agencourt AMPure XP beads by adding 180 μL of prepared beads to 100 μL of amplified sample and then following steps outlined in section B step 5.
Note: Alternatively, use Qiagen QIAquick PCR Purification Kit.
Assess DNA purity and quality with NanoDrop and bioanalyzer as specified in the pre-capture library amplification section.
Note: The desired 260/280 ratio is 1.7–2.0, and the desired fragment size is 150–500 bp. If either of these conditions is not met, that is a red flag.
Sequencing
Pool post-capture PCR amplified libraries.
Perform paired-end sequencing of DNA sequencing of pooled libraries.
Note: We performed this sequencing on an Illumina HiSeq 2500.
Data analysis
Our approach to read mapping and testing for ASE has been previously described in Diaz-Ortiz et al. (2022) (in Materials and Methods, subsection Read mapping and ASE analyses), but is listed in more detail below, also including analysis scripts (see Supplementary Information 2 and 3).
Assess sequencing data, trim reads, and filter for poor-quality reads prior to mapping.
To assess RNA-seq reads quality we employed FastQC (Andrews, 2010), while for reads quality filtering and trimming we used Trimmomatic (Version 032) (Bolger et al., 2014). We ran Trimmomatic to remove low-quality fragments in a 4 base wide sliding window (average window quality below PHRED 20), and low quality leading and trailing bases (below PHRED 10). We also dropped all the reads with average PHRED quality below 25, as well as reads shorter than 75 bases. Depending on the sample, 65%–80% of reads passed this trimming and filtering step, resulting in 6.5–24 million read pairs per sample for mapping.
Map reads to the human genome.
To perform unbiased allele specific read mapping to the reference human genome (hg19), we applied a WASP–STAR pipeline (Figure 2, reproduced from Figure S7 of Diaz-Ortiz et al., 2022). First, we mapped reads with STAR (Dobin and Gingeras, 2016), applying 2-step alignment, and filtered them for mapping bias using WASP (van de Geijn et al., 2015). Before proceeding with variant calling, we removed duplicate reads using rmdup_pe.py script incorporated into WASP pipeline. To call and filter single nucleotide variants (SNV), we used GATK tools; HaplotypeCaller, SelectVariants, and VariantFiltration.
Figure 2. Allele-specific expression (ASE) workflow. (A) RNA-seq data were first QC filtered and then mapped to the human reference genome (GRChg37) with STAR aligner using a 2-step approach. To remove allelic bias, the aligned reads were split into reads that did and did not overlap with single nucleotide variants (SNVs) and into separate BAM files. (B) The SNV genotypes were flipped in all reads that overlapped a SNV (in each read the genotype was swapped with that of the other allele) and re-mapped to reference genome again. Reads that did not re-map to the original location were discarded. (C) Filtered BAM files were then merged with BAM files that did not overlap with SNVs. Duplicate reads were removed using rmdup_pe.py script incorporated into the WASP pipeline. (D) SNVs were called and filtered using GATK tools. The allele specific counts were obtained by ASEReadCounter GATK tools. MBASED algorithm was used to test for ASE at the gene level. This figure is reproduced from Figure S7 of Diaz-Ortiz et al. (2022).
Obtain ASE specific read counts.
We obtained allele-specific read counts by GATK - ASEReadCounter. In order to filter out intergenic variants, we functionally annotated SNVs using VariantAnnotation (Obenchain et al., 2014) and TxDb.Hsapiens.UCSC.hg19.knownGene (Carlson and Maintainer, 2015) R packages.
Test for ASE.
To test for ASE at the gene level, we first selected proxy SNPs that were highly linked (r2 > 0.6) with rs199347 and located within a coding region for the gene of interest in order to assign allele of origin. Where possible, it may be useful to select several proxy SNPs per locus, as the number of aligned reads with sequence data at each ASE proxy SNP may differ. For GPNMB, we ultimately assigned the allele of origin for each transcript read based on genotype at rs199355, but we initially performed ASE analyses for both rs199355 and rs5850. Because both SNPs yielded identical results, we presented the data for rs199355, which had greater read counts. For KLHL7, we assigned the allele of origin based on genotype at rs2072368, as there were few alternative ASE proxy SNPs available. We then tested for allelic imbalance with a beta-binomial model with overdispersion using the MBASED R package (Mayba et al., 2014). P-values were adjusted for false discovery rate using the Benjamini-Hochberg method (Benjamini and Hochberg, 1995).
Note: An example of the read count data for GPNMB in its raw form is provided in Table S1 to give the reader a sense of the variability in counts obtained in a typical experiment. Table S1 also provides statistical information from a number of alternative binomial testing and FDR correction methods.
Validation of protocol
Two aspects of our results (some of which are highlighted in Diaz-Ortiz et al., 2022) suggest the reproducibility of our protocol. First, GPNMB consistently showed ASE when assayed by proxy SNP rs199355, with a reproducible effect size and direction across individuals who were heterozygous at the sentinel locus (rs199347) regardless of brain region samples or disease-state. Secondly, when ASE for this locus was assayed with a probe targeting a second proxy exonic SNP (rs8580, data not shown), the results did not differ significantly.
Additionally, various aspects of our work highlight both internal and external validity of our work. The internal validity of our assay is supported by the results from two individual populations not shown in Diaz-Ortiz et al. (2022): 1) individuals who were homozygous at the proxy SNP (rs8580), and 2) individuals who were homozygous at the sentinel SNP (rs199347) but heterozygous at the proxy SNP (rs8580). While for individuals in category 1 one would expect monoallelic expression (i.e., 100% ASE), for individuals in category 2 one would expect near equal expression of both alleles (i.e., absence of ASE). As expected, our assay showed allele counts consistent with monoallelic expression for individuals in category 1 and non-significant ASE for individuals in category 2 (Table S2). Finally, the directionality of the effect is consistent with previously published eQTL for this locus in healthy individuals (GTEx Consortium, 2017), supporting the external validity of our assay.
General notes and troubleshooting
Troubleshooting
NanoDrop 260/280 ratio outside of desired range for RNA sample: if the 260/280 ratio is outside the ideal range, this could reflect some impurity contaminating the sample (for example, a ratio of less than 1.7 would be suggestive of either phenol or protein contamination). We would suggest doing an ethanol precipitation or using the Qiagen QIAquick PCR Purification Kit to increase the sample purity.
RIN < 7 for RNA sample: when dealing with difficult-to-obtain human tissue, RINs may be lower when compared to samples originating from cultured cells. This may be due to either true RNA degradation (Figure 1C) or the presence of small RNAs (Figure 1B). We recommend visually inspecting the traces to distinguish between these possible causes and, in the latter, attempting small RNA cleanup of the sample.
Spectrophotometric analysis of negative LM-PCR control on NanoDrop suggests DNA yield that is more than a negligible amount: this is suggestive of either A) unincorporated LM-PCR primers being carried over or B) contamination. We recommend visually inspecting the DNA 1000 bioanalyzer trace for the negative control. A sharp peak below the 150 bp range is consistent with primers carried over from the LM-PCR reaction. However, a signal in the 150–500 bp range suggests contamination.
Acknowledgments
This work was supported by the NIH (F31 NS113481 to MDO, RO1 NS115139 to A.C.P., U19 AG062418 to A.C.P.), a Biomarkers Across Neurodegenerative Diseases (BAND) grant from the Michael J. Fox Foundation/Alzheimer’s Association/Weston Institute. A.C.P. is additionally supported by the Parker Family Chair, the Chan Zuckerberg Initiative Neurodegeneration Challenge Network, and the AHA/Allen Brain Health Initiative.
Competing interests
M.D.O. and A.C.P. are the inventors of a provisional patent submitted to the University of Pennsylvania that relates to targeting GPNMB as a potential therapeutic in Parkinson’s Disease.
Ethical considerations
The study of brain samples from autopsy specimens is not considered human subjects’ research. However, informed consent to brain donation was obtained from all subjects prior to death, with consent verified at the time of death by the guardian/next of kin.
References
Andrews, S. (2010). FastQC: A Quality Control Tool for High Throughput Sequence Data [online]. Accessed: 12 July 2021.
Benjamini, Y. and Hochberg, Y. (1995). Controlling the False Discovery Rate: A Practical and Powerful Approach to Multiple Testing. J. R. Stat. Soc. B. 57(1): 289–300.
Bolger, A. M., Lohse, M. and Usadel, B. (2014). Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30(15): 2114–2120.
Carlson, M. and Maintainer, B. P. (2015). TxDb.Hsapiens.UCSC.hg19.knownGene: Annotation package for TxDb object(s). R package.
Chen-Plotkin, A. S., Geser, F., Plotkin, J. B., Clark, C. M., Kwong, L. K., Yuan, W., Grossman, M., Van Deerlin, V. M., Trojanowski, J. Q., Lee, V. Y., et al. (2008). Variations in the progranulin gene affect global gene expression in frontotemporal lobar degeneration. Hum. Mol. Genet. 17(10): 1349–1362.
Diaz-Ortiz, M. E., Seo, Y., Posavi, M., Carceles Cordon, M., Clark, E., Jain, N., Charan, R., Gallagher, M. D., Unger, T. L., Amari, N., et al. (2022). GPNMB confers risk for Parkinson’s disease through interaction with α-synuclein. Science 377(6608): eabk0637.
Dobin, A. and Gingeras, T. R. (2016). Optimizing RNA-Seq Mapping with STAR. In: Carugo, O. and Eisenhaber, F. (Eds.). Data Mining Techniques for the Life Sciences (pp. 245–262). Methods in Molecular Biology. Humana Press, New York.
van de Geijn, B., McVicker, G., Gilad, Y. and Pritchard, J. K. (2015). WASP: allele-specific software for robust molecular quantitative trait locus discovery. Nat. Methods 12(11): 1061–1063.
GTEx Consortium (2017). Genetic effects on gene expression across human tissues. Nature 550(7675): 204–213.
Mayba, O., Gilbert, H. N., Liu, J., Haverty, P. M., Jhunjhunwala, S., Jiang, Z., Watanabe, C. and Zhang, Z. (2014). MBASED: allele-specific expression detection in cancer tissues and cell lines. Genome Biol. 15(8): e1186/s13059-014-0405-3.
Mercer, T. R., Clark, M. B., Crawford, J., Brunck, M. E., Gerhardt, D. J., Taft, R. J., Nielsen, L. K., Dinger, M. E. and Mattick, J. S. (2014). Targeted sequencing for gene discovery and quantification using RNA CaptureSeq. Nat. Protoc. 9(5): 989–1009.
Obenchain, V., Lawrence, M., Carey, V., Gogarten, S., Shannon, P. and Morgan, M. (2014). VariantAnnotation: a Bioconductor package for exploration and annotation of genetic variants. Bioinformatics 30(14): 2076–2078.
Supplementary information
The following supporting information can be downloaded here:
Allele specific expression workflow.sh
MBASED beta-binomial with overdispersion GPNMB unphased with simulations.R
Scripts inWORDformat 6 28 23
Table S1. ASEProtocol
Table S2. rs199355 Allele Counts in special cases
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Neuroscience > Nervous system disorders > Parkinson's disease
Molecular Biology > RNA > RNA sequencing
Systems Biology > Genomics > Sequencing
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Application of Electrical Stimulation to Enhance Axon Regeneration Following Peripheral Nerve Injury
SW Supriya S. Wariyar
PW Patricia J. Ward
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4833 Views: 359
Reviewed by: Alessandro DidonnaAnand Ramesh PatwardhanZheng Zachory Wei
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Original Research Article:
The authors used this protocol in Developmental Neuroscience May 2021
Abstract
Enhancing axon regeneration is a major focus of peripheral nerve injury research. Although peripheral axons possess a limited ability to regenerate, their functional recovery is very poor. Various activity-based therapies like exercise, optical stimulation, and electrical stimulation as well as pharmacologic treatments can enhance spontaneous axon regeneration. In this protocol, we use a custom-built cuff to electrically stimulate the whole sciatic nerve for an hour prior to transection and repair. We used a Thy-1-YFP-H mouse to visualize regenerating axon profiles. We compared the regeneration of axons from nerves that were electrically stimulated to nerves that were not stimulated (untreated). Electrically stimulated nerves had longer axon growth than the untreated nerves. We detail how variations of this method can be used to measure acute axon growth.
Keywords: Peripheral nerve injury Sciatic nerve Electrical stimulation Cuff Fibrin glue Axon regeneration
Background
Peripheral nerve injury is an important clinical problem; generally, the functional recovery is very poor (Lee and Wolfe, 2000; Höke and Brushart, 2010; Enax-Krumova et al., 2017; Bulut et al., 2018; Zheng et al., 2018; Wang et al., 2019) if the injury is more proximal to the spinal cord, involves a gap of ≥ 5 mm, or involves mixed sensory and motor nerves. The main reason for this poor recovery is that the injured neurons are required to regenerate their axons over long distances, sometimes many centimeters, to reconnect to their peripheral target. The regenerating axons have a very slow rate of growth of ∼1 mm/day (Sulaiman and Gordon, 2000 and 2013; Sulaiman and Kline, 2006; Sulaiman et al., 2011). Our lab and many others have demonstrated that activity-based therapies (like exercise in the form of treadmill running, optical stimulation, or electrical stimulation) after peripheral nerve injury significantly enhances axon regeneration (Al-Majed et al., 2000; Brushart et al., 2002; Wood et al., 2012; Thompson et al., 2014; Gordon and English, 2016; Ward and English, 2019). Many studies have shown that electrical stimulation for 1 h at 20 Hz enhances axon regeneration after nerve crush or transection models in rats, mice (Al-Majed et al., 2000; Brushart et al., 2002 and 2005; English et al., 2007; Geremia et al., 2007; Elzinga et al., 2015; Shapira et al., 2019), and human subjects (Gordon et al., 2010; Wong et al., 2015; Barber et al., 2018). More recently, conditioning electrical stimulation of the intact nerve, prior to nerve transection and repair, has been shown to enhance nerve regeneration and functional recovery (Senger et al., 2018, 2019, 2020a and 2020b; Gordon, 2020). Here, we demonstrate that a single session of electrical stimulation for an hour at a controlled frequency applied to the nerve using a cuff prior to transection and repair significantly enhances axon regeneration compared to no treatment. We used Thy-1-YFP-H mice (Jackson Laboratory, stock no. 003782) to visualize regenerating axons. Thy-1-YFP-H are transgenic mice that express yellow fluorescent protein (YFP) at high levels in a subset of motor and sensory neurons, as well as subsets of central neurons. The axons are brilliantly fluorescent all the way to the terminals, which provides a useful visual outcome that can be quantified to compare regeneration. The H strain is nearly identical to Thy-1-YFP-16 (Jackson laboratory, stock no. 003709), except for the percentage of neurons expressing YFP. Thy-1-YFP-H express YFP in 10%–30% of dorsal root ganglia sensory neurons, whereas expression of Thy-1-YFP-16 is much greater. The advantage of sparse labeling is that individual axons can be accurately and precisely (length and branching) traced into the distal stump. The advantage of using an electrical cuff to stimulate (in contrast to needle electrodes or wires) is that the target nerve can be selectively stimulated without unwanted stimulation of muscle or neighboring nerves. The sciatic nerve is commonly used to study peripheral axon regeneration, and it consists of three terminal branches (Figure 1) that are held together by epineurium to form the whole sciatic nerve. The epineurium can be delicately surgically removed to separate the branches proximally, and the cuff can be placed on specific branches of the sciatic nerve for various experimental designs.
Figure 1. White star shows the three terminal branches of the sciatic nerve. 1) Tibial nerve, 2) common fibular or common peroneal nerve, and 3) sural nerve. The branches were separated by removing a portion of the epineurium using fine forceps. Blue star indicates a small proximal branch of the sciatic nerve that innervates a portion of the hamstring muscle. This image was taken on an AmScope ZM-4TW3-FOR-9M Digital Professional Trinocular Stereo Zoom Microscope at 4.5× zoom.
Materials and reagents
Silastic sheeting (Dow Corning Corporation medical products, catalog number: 501-1)
0.5 mL tubes (Corning, catalog number: 3750)
Thrombin (MP Biomedical, catalog number: 76461-568)
Fibrinogen (Sigma, catalog number: F3879-1G)
Fibrin glue (Akhter et al., 2019; Ward and English, 2019)
Isoflurane (Piramal Healthcare, catalog number: RXISO-250)
Gauze pads (Fisher Brand, catalog number: 22-362-186)
Betadine solution (Purdue Frederick Company, catalog number: 10224)
Ethanol (Thermo Fisher, catalog number: T038181000CS)
10 μL pipette tips (Eppendorf, catalog number: 022492004)
0.9% sodium chloride (NaCl) (Fisher, catalog number: S271-1)
4% PFA (paraformaldehyde prepared in 0.1 M PBS) (Sigma, catalog number: P6148-500G)
1× phosphate buffered saline (PBS) prepared from PBS tablets (Sigma, catalog number: P4417-100TAB)
Euthanasia solution (10 mg/mL made in sterile saline) (Med Vet International, catalog number: RXEUTHASOL)
Sylgard 184 silicone elastomer base and curing agent (Krayden DOW, catalog DC4019862)
Equipment
Cuff electrode
Dumont Forceps #5 (Fine Science Tools, catalog number: 11251-10)
Surgical scissors (Fine Science Tools, catalog number: 14084-09)
Spring scissors (Fine Science Tools, catalog number: 15006-09)
Silastic tubing (Laboratory Tubing, catalog number: 508-006)
Insulated wire (Cooner Wire, catalog number: AS631)
Flexible silicone (Dow Corning, catalog number: Sylgard® 184)
Sewing needle size #9 (Singer, catalog number: 01125)
Table clamp (to hold the silicone tubing in place)
Wooden applicator stick (Electron microscopy sciences, catalog number: 07230)
Silk braided thread size 0 (J.A Daknatel & Son Inc., catalog number: 3-766-900)
Microscope (Nikon, model number: SMZ800)
LED Tester, set at 15 V for checking current leakage from cuff electrode (Hewlett Packard Palo Alto Hp California 630)
For electrical stimulation
Grass Stimulator and Amplifier Box (Ward and English, 2019)
Custom Lab View Program (Ward and English, 2019)
Surgery
Thy-1 YFP-H adult mice (2–4 months old; male and/or female weighing 18–30 g) (Jax laboratory, stock number: 003782)
Surgical drapes (Med-Vet International, catalog number: SKU DR1826)
Needle holder with suture cutter (Fine Science Tools, catalog number: 12502-14)
Hair clipper (Wahl combo kit #9990-1201)
Deltaphase Isothermal Pad (Braintree scientific, model: 39DP)
Warm water circulator (Gaymar TP500)
Surgical stereoscope (AmScope SM-4BZ-80S)
Glass bead sterilizer (Braintree Scientific, catalog number: GER 5287-120V)
Surgical board (Fisher Brand, catalog number: 0900224C)
Surgical tape (3M Micropore tape, 1532-1)
Cotton tipped applicator (Dukal corporation, 9006)
Eye ointment (Refresh P.M.)
Triple antibiotic ointment (Mckesson Medical Surgical Inc, catalog number: 955410)
Meloxicam oral suspension (Med-Vet International, catalog number: RXMELOXIDYL10)
Absorbable: coated Visorb undyed braided polyglycolic acid suture NSF-2 19 mm 3/8 30" (75 cm) (CP medical, catalog number: 421A)
Non-absorbable Monofilament polyamide suture (Redilon 5-0) 1 metric 18" (45 cm) (CP medical, catalog number: 661B)
Software
FIJI (NIH) to process and analyze the images
Microsoft Excel to organize the axon length data
Procedure
Cuff building (Rios et al., 2019)
Take approximately 1 inch of silicon tubing. Insert the tubing onto a wooden applicator that is held by a holder. Under a stereoscope, cut a slit halfway through the tubing (Figures 2 and 3A).
Figure 2. Cuff building schematics. A) Schematic of silicon tubing. B) Silicon tubing is placed on a wooden applicator for ease of handling while building. A 1/2 inch slit is made through the tubing.
Figure 3. Cuff building procedure. A) Cuff tubing of approximately 1 inch in length. The tube is cut to create a slit. B) A total of six holes are made in the tube: two on the left of the slit and four on the right of the slit. C) Cooner wire is de-insulated approximately 4–5 cm in length. D) The de-insulated wire is tied with a silk thread. E) The silk thread along with the Cooner wire is passed along the first hole through the superior hole, back into the inferior hole, and then back from the first hole. F) The de-insulated wire is wrapped around the insulated wire to secure it, so that the wire does not detach from the cuff tubing. G) Same process (from C–F) is repeated to make a complete cuff consisting of two de-insulated wires lying inside the cuff tubing. The outside exposed wire is covered with flexible silicone to prevent current leakage. The cuff is trimmed to a length of ∼1 cm. H) Setup for testing the cuff (bubble test) consisting of a Petri dish filled with 0.9% NaCl and LED tester maintained at 15 V. I) The cuff shows bubbles escaping only from inside the cuff tubing where the Cooner wire is de-insulated. J) Full view of the custom-made cuff electrode; extra length allows for a more flexible setup on the surgical table (surgical supplies, stimulator, and experimental animal) during electrical stimulation, e.g., a very short cuff requires the stimulus isolation unit to be placed a short distance away from the animal.
Using a 30 G needle, make a hole (1) approximately 4–5 mm from the slit on the left side of the tubing. On the right side of the tubing, make a superior hole (2) at 2–3 mm from the slit. Similarly, make an inferior hole (3) at 2–3 mm below the superior hole 2 (Figure 4).
Note: Align hole 1 with 2. Align 3 such that the 1 on the left side of the tubing is in the center of 2 and 3.
Figure 4. Schematic of the location of holes to be made in the tubing with a 30 G needle. These holes provide passage for the Cooner wire to be threaded through, so that the de-insulated wire rests completely within the tubing.
Similarly, make another hole (4) leaving 2–3 cm distance from 1 on the left. On the right view, make a superior hole (5), leaving 2–3 cm distance from 2, and an inferior hole (6) (below 5) leaving 2–3 mm from the superior hole 5. A total of six holes are made in the tubing with the 30 G needle (Figure 4).
Note: Align hole 4 with 5 and 6, such that 4 is in the center of 5 and 6. Holes 4, 5, and 6 are equidistant from 1, 2, and 3.
Take approximately 30 cm of Cooner wire. De-insulate approximately 4–5 cm in length of the tip of the wire (Figures 5 and 3C). Thread 20 cm of silk thread into the eye of a #9 sewing needle. Tie a knot on the de-insulated section of the fine wire with the silk thread. Secure the knot so that it does not detach from the fine wire (Figure 3D).
Note: Cooner wire consists of 13 fine braided metallic wires within insulation. As the insulation is removed at the ends of the wire, the metallic wires tend to fray. Care should be taken to coil it around itself to minimize the fray.
Figure 5. Schematic of the inside of the tubing, showing the placement of insulated and de-insulated portions of the Cooner wire
Pass the silk thread through hole 1 (left) and then through hole 2 (right), such that a portion of the de-insulated wire lies inside the tubing (Figures 5 and 3E–3F).
Turn the needle 180° and pass it through hole 3 on the right and then out through hole 1 on the left of the tubing again, making sure a portion of the de-insulated wire lies inside the tubing (Figures 5 and 3E–3F). Repeat the process with another 30 cm of Cooner wire, and thread it through hole 4 on the left through to the corresponding two holes 5 and 6 on the right.
The remaining de-insulated wire that comes out of the left hole is tightly coiled 4–5 times around the insulated wire, to secure it from detaching from the cuff (Figure 3F).
Note: Care should be taken to coil the de-insulated wire so that none of the 13 small metallic wires become frayed. Cut off any excess de-insulated wire to minimize current leakage.
Mix 0.8 g of Sylgard 184 silicone elastomer base with 0.1 g of Sylgard 184 silicone elastomer curing agent. Use this mixture to seal any non-insulated metallic fine wire that is outside the cuff tubing to prevent current leakage. The flexible silicon is cured in an incubator maintained at 100 °C for 5 min.
Note: The flexible silicone is repeatedly applied to and cured in the incubator until all the exposed (uninsulated) wire outside the cuff tubing is covered.
Trim the cuff to ∼1 cm in length (Figure 3G). Test the cuff (bubble test) using a LED tester that is maintained at 15 V. Fill a Petri dish with 0.9% NaCl (saline). Submerge the cuff in the saline solution. De-insulate the distal ends (1–2 cm) of the Cooner wire and connect to the LED tester. Perform a bubble test, wherein a properly built cuff should have bubbles escaping only from inside the cuff where the wire is de-insulated. Any bubbles escaping from outside the cuff should be sealed again using the flexible silicone (Figure 3I). This step is critical to prevent off-target stimulation of nearby nerves/muscle.
Surgical preparation
Sterilize all surgical instruments in a glass bead sterilizer maintained at 250 °C for 30 s. Cool the surgical instruments at room temperature for at least 5 min (or until cool to the touch).
Place heating pad or circulator under a surgical board to maintain body heat.
Place the nose cone on the surgical board and secure it with surgical tape.
Thaw out aliquots of thrombin (thrombin is made by adding 25 μL of thrombin stock solution to 975 μL of 45 mM CaCl2) at room temperature. Fibrinogen is prepared by mixing 10 mg of fibrinogen with 100 μL of distilled water until completely dissolved. Fibrinogen tends to coagulate when left at room temperature and should be used within 5–6 h of preparation. Coagulated fibrinogen should not be used because the nerve stumps will not be secured leading to a large gap, which greatly reduces the number of axons able to regenerate.
Weigh the animal and place it in the induction chamber to induce anesthesia (5% isoflurane in 1 L/min oxygen).
Place the animal on the surgical board with its nose fitted to a nose cone. The anesthesia is then maintained at 2% isoflurane. Clip hair using electric hair clipper starting from hip to ankle of the hindlimb.
Remove all the hair and the first drape from under the animal. Wipe the hindlimb of the animal with alcohol and betadine in succession for three times each.
Perform a toe pinch to check any reaction to ensure the animal is within the surgical plane. Titrate isoflurane as needed.
Nerve surgery should be performed with aide of surgical microscope.
Drape the animal with sterile drape such that only the site of surgery is exposed.
Identify the femur by palpating the skin.
Make a 4 cm incision along the length of the femur. Blunt dissect the fascia between the biceps femoris and hamstring muscles.
Using #5 forceps, carefully isolate the sciatic nerve from the underlying connective tissues (Figure 6A).
Figure 6. Sciatic nerve stimulation with cuff. A) Sciatic nerve is exposed by making a ~4 cm incision on the skin. B) A custom-made cuff is wrapped around the whole sciatic nerve (or a chosen branch), such that the nerve rests on top of the de-insulated wire inside the cuff. The de-insulated distal ends of the cuff are connected to a grass stimulator. Nerve stimulation can occur prior to or after nerve transection and repair, which has been previously detailed (Akhter et al., 2019).
Wrap the cuff around the sciatic nerve such that the nerve is in contact with the de-insulated wire inside the cuff (Figure 6B). Connect the de-insulated wires at the distal tip of the cuff to the amplifier box. Using a grass stimulator, stimulate the nerve for 1 h at 20 Hz (Gordon 2020). Stimulating the nerve immediately prior to transection allows for visual confirmation of stimulation (visible muscle twitches).
After stimulating the nerve for 1 h, place a small 4 mm square of silastic sheeting under the nerve. Apply fibrin glue consisting of thrombin (2 μL) and fibrinogen (1 μL) (2:1 ratio) to the sciatic nerve, so that the nerve is secured to the silastic sheet (1–2 min for the fibrin glue to coagulate).
Then, completely transect the nerve with sharp micro scissors. Care should be taken so that the cut nerve stumps are in close proximity of each other. Apply fibrin glue again for 1–2 min.
Without disturbing the transected nerve, suture the muscles using absorbable suture and the skin using non-absorbable nylon suture.
Apply triple antibiotic cream on the surgical site using a sterile cotton-tipped applicator.
Post-surgery
Remove the animal from the nose cone and place it into a clean recovery cage (without bedding) placed on a warm water circulating heating pad (Gaymar TP500). Once the animal is ambulatory, administer oral analgesic [meloxicam (5 mg/kg)]. Two weeks after nerve transection, there should be no observable scar formation in the skin or muscle.
Two weeks after nerve transection, inject the animal with 0.3–0.4 mL of euthanasia solution (10 mg/mL). Following euthanasia, dissect the sciatic nerve from the animal (4 mm proximal to the repair site to the distal end). Care should be taken to carefully collect the whole length of the distal nerve without damaging the repair site.
Fix the nerve in 1 mL of 4% PFA for 1 h at room temperature and then transfer to 1× PBS for 5–10 min.
Place the nerve on a clean slide and add a few drops of Vectashield with DAPI (hard mount) to the nerve.
Place a glass coverslip on the nerve. Add weights (3–5 g) on top of the coverslip so as to flatten the nerve for 24 h.
Image the nerve using Nikon AR125 HD confocal microscope using GFP (488) laser with a 10× objective.
Use the image tile function so that the whole length of the nerve on the slide is visible on the computer screen. Set up Z stack with a step size of 4.75 microns. Apply a line average of 4 to reduce background noise. Image the nerve on resonant mode for faster imaging. Do not apply zoom for imaging. The image is automatically stitched by the Nikon software to reconstruct the proximal and distal end of the nerve in three dimensions.
Data analysis
Load the nerve image into FIJI software (NIH).
Using the straight-line tool with a diameter of 10 mm, place a region of interest (ROI) at the injury site. When the image is zoomed in, the injury site is apparent by the disrupted and unorganized axons. That point is where the transection of the nerve was performed: the injury site.
Using the freehand tool, measure the length of each axon from the repair site to the distal tip following the axon throughout the Z stack. When measuring the length of each axon, always start measuring the shortest axons first, as they will be difficult to visualize later when longer axon profiles are overlaid on the image (Figure 7). Compare the length of axon growth between electrically stimulated nerves and untreated nerves. Ensure the scale bar and image calibration is correct (J > Tools > Scale bar).
Figure 7. Image of a nerve from a Thy-1-YFP-H mouse. The regenerating axons can be visualized by the YFP reporter. A) Beginning with the shortest regenerating axons, individual axon profile lengths are measured by tracing through the Z stack. B) Zoomed image of the nerve showing the length of growing axons from the injury site.
Sort the length of each regenerating axons into bins of increasing lengths in Microsoft Excel.
Calculate the frequency by selecting the entire column of frequency and then use the formula {= frequency (values in the length column, all values in the bin column)}.
Calculate the proportion using the formula = [(each frequency value × 100)/total number of axons].
Then, calculate the cumulative percentage of total number of regenerating axons.
Electrically stimulated nerves exhibit longer axons compared to non-stimulated nerves, i.e., Untreated (Figure 8).
Figure 8. Stimulated vs. unstimulated nerve growth following nerve transection. A) Image of a nerve following transection and repair with fibrin glue. The injury site and direction of axon growth are indicated. Note that many regenerating axon profiles are close to the repair site. B) Image of a nerve following electrical stimulation and transection and repair with fibrin glue. The injury site and direction of axon growth are indicated. Note the longer length of many regenerating axons compared to the untreated nerve. C) The cumulative frequency distributions of each group. The cumulative distribution of axon lengths in the electrically stimulated fibrin glue–repaired group is significantly shifted to the right, which indicates longer axons.
Notes
As with all experimental models, this has its own advantages and disadvantages. In this protocol, we utilized a reporter mouse (Thy-1-YFP-H). The reporter is advantageous in that nerves do not require sectioning and immunostaining. Alternatively, peripheral regeneration assays can use nerves from wildtype mice C57BL/6J (Jackson laboratory, stock no. 000664) by sectioning and immunostaining the nerves with neurofilament heavy chain (NF200) (Dun and Parkinson, 2015). SCG10 is another commonly used antibody, but it is preferentially expressed in regenerating sensory and sympathetic axons rather than motor (Shin et al., 2014; Lee et al., 2022). Because of their abundance, precise analysis of individual axons is difficult when using NF200 or SCG10.
Care should be taken not to add excess flexible silicon on the cuff to cover the de-insulated wires outside the cuff, because it makes the cuff bigger and bulkier. Bulkier cuffs are more difficult to apply to the nerve and also require a larger surgical site. Furthermore, if the cuff is too bulky, the nerve might lose contact with the de-insulated wire inside the cuff.
When properly cared for, cuffs can be re-used for multiple stimulations. The cuff is cleaned with soap and water and wiped gently with a delicate task wipe soaked in 70% ethanol. Eventually, the cuff will become leaky. It is good practice to bubble test each cuff before use. If the bubble test shows the bubbles escaping from inside and outside of the cuff, then a new cuff must be built.
Fibrinogen tends to coagulate; hence, it should always be made fresh on the day of the surgery. When doing multiple surgeries, fresh pipette tips are used to extract the solution each time to avoid mixing with thrombin, which will cause it to coagulate.
To avoid mushrooming of the cut nerve stumps, sharp spring scissors should be used to transect the nerve.
In our studies, we routinely use fibrin glue to repair transected nerves and found that fibrin glue is an adjuvant to activity-based therapy, such as exercise (Wariyar et al., 2022). If fibrin glue is not used, another means of securing the cut nerve stumps together must be used (fine sutures).
English and colleagues reported a sex difference in axon regeneration when using a form of activity-based treatment (treadmill running) (Wood et al., 2012). At the transcriptional level, sexually dimorphic programs are acutely activated by nerve injury (Chernov and Shubayev, 2022), which may influence regenerative ability and response to treatments. Potential sex differences must be considered in experimental design.
Ethical considerations
All procedures involving animals were performed according to and approved by the Institutional Animal Care and Use Committee of Emory University.
Acknowledgments
This work was supported in part by the NIH National Institute of Neurological Disorders and Stroke under award number K01NS124912 as well as a developmental grant from the NIH-funded Emory Specialized Center of Research Excellence in Sex Differences U54AG062334.
Competing interests
The authors declare that no competing interests exist.
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4,834 | https://bio-protocol.org/en/bpdetail?id=4834&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Protein Level Quantification Across Fluorescence-based Platforms
HR Hector Romero
AS Annika Schmidt
MC M. Cristina Cardoso
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4834 Views: 855
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Original Research Article:
The authors used this protocol in Nucleus Feb 2022
Abstract
Biological processes are dependent on protein concentration and there is an inherent variability among cells even in environment-controlled conditions. Determining the amount of protein of interest in a cell is relevant to quantitatively relate it with the cells (patho)physiology. Previous studies used either western blot to determine the average amount of protein per cell in a population or fluorescence intensity to provide a relative amount of protein. This method combines both techniques. First, the protein of interest is purified, and its concentration determined. Next, cells containing the protein of interest with a fluorescent tag are sorted into different levels of intensity using fluorescence-activated cell sorting, and the amount of protein for each intensity category is calculated using the purified protein as calibration. Lastly, a calibration curve allows the direct relation of the amount of protein to the intensity levels determined with any instrument able to measure intensity levels. Once a fluorescence-based instrument is calibrated, it is possible to determine protein concentrations based on intensity.
Key features
• This method allows the evaluation and comparison of protein concentration in cells based on fluorescence intensity.
• Requires protein purification and fluorescence-activated cell sorting.
• Once calibrated for one protein, it allows determination of the levels of this protein using any fluorescence-based instrument.
• Allows to determine subcellular local protein concentration based on combining volumetric and intensity measurements.
Graphical overview
Protein level quantification across fluorescence-based platforms
Keywords: Quantification Single-cell protein levels FACS Fluorescence Microscopy Western blotting
Background
The use of fluorescently tagged proteins is a common tool in cell biology to study a variety of molecular processes. Although fluorescence is directly related to concentration of the fluorophore, the values of intensities do not translate directly into concentration of molecules. This is important to reproduce (patho)physiological concentrations of molecules including proteins. Western blots can be used to determine the average amount of proteins within a cell population. We performed fluorescence-activated cell sorting (FACS) prior to analysis by western blot to quantify protein levels within a cell population with defined fluorescence intensities and cell amounts. In addition, we developed a method that allows the direct relation of fluorescence intensities to get molecular concentration categories at the single-cell level. Furthermore, we expanded the method to allow concentration comparison between samples detected in different systems. Lastly, we extended the method to calculate average subcellular concentration variations. Other existing methods to determine protein concentrations are based on single-molecule imaging to calculate the number of molecules in beads (Chiu et al., 2001; Sugiyama et al., 2005) or, alternatively, use lipid or polymer layers in which the density of the fluorophore is known (Dustin, 1997; Zwier et al., 2004; Galush et al., 2008). These methods, although maybe more accurate in the estimation of concentration than ours, require calibration for each experiment and/or sophisticated equipment, whereas our method relies on equipment available in most molecular cell biology laboratories.
In the publication in which we utilized the method (Zhang et al., 2022), we used it to quantify ectopic protein concentrations in single cells and subcellular compartments. This allowed us to reproduce the same concentrations in cells and in in vitro experiments and to relate it to physiological tissue concentrations. Variations of the method can be used to quantify endogenous levels of tagged proteins (i.e., in cells with genomically engineered loci) or proteins labeled with antibodies.
Materials and reagents
Biological samples
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A8412); see General note 1
C2C12, mouse (Mus musculus) myoblasts (Yaffe and Saxel, 1977); see General note 1
Plasmid pc1208 (pEG-MeCP2) (Kudo et al., 2003); see General note 1
Rat IgG anti-MeCP2 4H7 antibody. Self-made, not commercial (Jost et al., 2011); see General note 1
Donkey anti-rat IgG-Cy3 antibody (Jackson ImmunoResearch, catalog number: 111560); see General notes 1 and 2
Reagents
1,4-Diazabicyclo-[2.2.2]octane (DABCO) (Sigma-Aldrich, catalog number: D2522)
2-propanol (AppliChem GmbH, catalog number: 131090.1212)
4′,6′-diamidine-2-phenylindole dihydrochloride (DAPI) (Carl Roth, catalog number: 6335.1)
Aluminum sulfate 14–18 hydrate (Carl Roth, catalog number: 3731.1)
Ammonium persulfate (Carl Roth, catalog number: 9592.3)
Bromophenol blue (Bio-Rad Laboratories, catalog number: 161-0404)
Coomassie, Brilliant Blue R (Sigma-Aldrich, catalog number: 1.12553)
Dimethylsulfoxide (DMSO) (Sigma-Aldrich, catalog number: D4540)
Di-sodium hydrogen phosphate 7-hydrate (Na2HPO4·7H2O) (Carl Roth, catalog number: X987.2)
Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D9779)
Ethanol absolute pure, pharma grade (AppliChem GmbH, catalog number: A4230)
Ethylenedinitrilotetraacetic acid (EDTA) (AppliChem GmbH, catalog number: 131026.1211)
Glucose (Sigma-Aldrich, catalog number: G5400)
Glycerol (Sigma-Aldrich, catalog number: G9422)
Hybond ECL membrane (nitrocellulose membrane) (VWR, catalog number: RPN3032D)
Low fat milk pulver (Sucofin)
Methanol for analysis EMPARTA® ACS (Sigma Aldrich, catalog number: 1070182511)
Mowiol® 4-88 (Sigma-Aldrich, catalog number: 81381)
NonidetTM P-40 substitute (NP-40) (Roche, catalog number: 11332473001)
Pepstatin A (Sigma-Aldrich, catalog number: P5318)
Phenylmethylsulfonyl fluoride (PMSF) (Carl Roth, catalog number: 6367.1)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9541)
Potassium dihydrogen phosphate (KH2PO4) (Carl Roth, catalog number: 3904.1)
Tris (Sigma-Aldrich, catalog number: 93362)
Sodium chloride (NaCl) (Carl Roth, catalog number: 3957.1)
Sodium dodecyl sulfate (SDS) (Sigma-Aldrich, catalog number: 11667289001)
Trans-epoxysuccinyl-L-leucylamido(4-guanidino)butane (E64) (Sigma-Aldrich, catalog number: E3132)
Tween 20 (Carl Roth, catalog number: 131026.1211)
Whatman 3MM CHR (Whatman paper) (Cytiva, catalog number: 3030-672)
Solutions
Acrylamide/bis-acrylamide 30% (Polyacrylamide) (Sigma-Aldrich, catalog number: A3699)
Dulbecco’s modified Eagle’s medium, high glucose (Sigma-Aldrich, catalog number: D7777)
Fetal bovine serum advanced (Capricorn Scientific, catalog number: FBS-11A)
Formaldehyde solution, 36.5%–38% in H2O (Sigma-Aldrich, catalog number: F8775)
Orthophosphoric acid (Carl Roth, catalog number: 6366.1)
PierceTM 660 nm Protein Assay Reagent (Thermo Fisher, catalog number: 22660)
PierceTM 10× Western Blot Transfer buffer, methanol-free (transfer buffer) (Thermo Fisher, catalog number: 35040)
Tetramethylethylenediamine (TEMED) (Sigma-Aldrich, catalog number: T9281)
Trypsin (Sigma-Aldrich, catalog number: T4049)
Ammonium persulfate 10% (see Recipes)
Coomassie destaining solution (see Recipes)
Coomassie staining solution (see Recipes)
DAPI solution (see Recipes)
E64 solution (see Recipes)
Loading buffer (see Recipes)
Low-fat milk 3% in PBS (see Recipes)
Low-fat milk 5% in PBS (see Recipes)
Lysis buffer (see Recipes)
Mounting media (see Recipes)
Pepstatin A solution (see Recipes)
Phosphate buffered saline (PBS) (see Recipes)
PBS-EDTA 0.02% (w/v) (see Recipes)
PMSF solution (see Recipes)
Polyacrylamide gel 5% (stacking) for 10 mL gel (see Recipes)
Polyacrylamide gel 8% (separating) for 10 mL gel (see Recipes)
Running buffer 10× (see Recipes)
Sodium dodecyl sulfate (SDS) 10% (see Recipes)
Tris 1 M pH 6.8 (see Recipes)
Tris 1 M pH 8.5 (see Recipes)
Tris 1.5 M pH 8.8 (see Recipes)
Recipes
Ammonium persulfate 10%
Reagent Final concentration Quantity
Ammonium persulfate 10% (w/v) 1 g
H2O n/a to 10 mL
Total n/a 10 mL
Coomassie destaining solution
Reagent Final concentration Quantity
Ethanol 10% (v/v) 40 mL
Orthophosphoric acid 2% (v/v) 2 mL
H2O n/a 58 mL
Total n/a 100 mL
Coomassie staining solution
Reagent Final concentration Quantity
Coomassie 0.2% (w/v) 0.2 g
Orthophosphoric acid 2% (v/v) 2 mL
Ethanol 10% (v/v) 10 mL
H2O n/a to 100 mL
Total n/a 100 mL
DAPI solution
Reagent Final concentration Quantity
DAPI 1 mg/mL 10 mg
H2O n/a to 10 mL
Total n/a 10 mL
E64 solution
Reagent Final concentration Quantity
E64 1 mM 360 μg
Ethanol 50% (v/v) 500 μL
H2O n/a 500 μL
Total n/a 1 mL
Loading buffer
Reagent Final concentration Quantity
Tris 50 mM 0.3 g
SDS 2% (v/v) 2 mL
Glycerol 10% (v/v) 10 mL
Bromophenol blue 0.01% (v/v) 100 μL
DTT 100 mM 1.5 g
H2O n/a to 100 mL
Total n/a 100 mL
Low-fat milk 3% in PBS
Reagent Final concentration Quantity
Low-fat milk pulver 3% (w/v) 0.3 g
PBS n/a to 10 mL
Total n/a 10 mL
Low-fat milk 5% in PBS
Reagent Final concentration Quantity
Low-fat milk pulver 3% (w/v) 0.5 g
PBS n/a to 10 mL
Total n/a 10 mL
Lysis buffer
Reagent Final concentration Quantity
Tris 25 mM 0.03 g
NaCl 1 M 0.6 g
Glucose 50 mM 0.09 g
EDTA 10 mM 0.03 g
Tween 20 0.2% (v/v) 20 μL
NonidetTM P-40 substitute 0.2% (v/v) 20 μL
PMSF solution 1 mM 1 μL
E64 solution 10 μM 1 μL
Pepstatin A solution 29 μM 1 μL
H2O n/a 10 mL
Total n/a 10 mL
Mounting media
Reagent Final concentration Quantity
Mowiol 4-88 13% (w/v) 8 g
Tris-HCl 1 M pH 8.5 133 mM 8 mL
H2O n/a 32 mL
Glycerol 33% (v/v) 20 mL
DABCO 2% (w/v) 1.2 g
Total n/a 60 mL
Add mowiol to Tris and H2O and heat to 50–60 °C while stirring.
Cool down to room temperature.
Add glycerol and stir again.
Add DABCO and dissolve it by stirring.
Spin the solution at 5,000× g for 15 min.
Aliquot the supernatant and store at -20 °C.
Pepstatin A solution
Reagent Final concentration Quantity
Pepstatin A 2.9 mM 2 mg
DMSO n/a 1 mL
Total n/a 1 mL
Phosphate buffered saline (PBS)
Reagent Final concentration Quantity
NaCl 1.37 M 80 g
KCl 27 mM 2 g
Na2HPO4·7H2O 10 mM 21.7 g
KH2PO4 10 mM 2.4 g
H2O n/a to 1 L
Total n/a 1 L
PBS-EDTA 0.02% (w/v)
Reagent Final concentration Quantity
EDTA 0.02 % (w/v) 2 mg
PBS n/a to 100 mL
Total n/a 100 mL
PMSF solution
Reagent Final concentration Quantity
PMSF 100 mM 17.42 mg
2-propanol n/a 1 mL
Total n/a 1 mL
Polyacrylamide gel 5% (stacking) for 10 mL gel
Reagent Final concentration Quantity
Polyacrylamide 5% (v/v) 0.33 mL
Tris 1 M pH 6.8 125 mM 0.25 mL
SDS 10% 0.1% (v/v) 0.02 mL
Ammonium persulfate 10% 0.1% (v/v) 0.02 mL
TEMED 0.001% (v/v) 0.002 mL
H2O n/a 1.4 mL
Total n/a 2 mL
Polyacrylamide gel 8% (separating) for 10 mL gel (see General note 1)
Reagent Final concentration Quantity
Polyacrylamide 8% (v/v) 2.7 mL
Tris 1.5 M pH 6.8 375 mM 2.5 mL
SDS 10% 0.1% (v/v) 0.1 mL
Ammonium persulfate 10% 0.1% (v/v) 0.1 mL
TEMED 0.0006% (v/v) 0.006 mL
H2O n/a 4.6 mL
Total n/a 10 mL
Running buffer 10×
Reagent Final concentration Quantity
Tris base 25 mM 30 g
Glycin 1.92 M 144 g
SDS 1% (v/v) 100 mL
H2O n/a to 1 L
Total n/a 1 L
SDS 10%
Reagent Final concentration Quantity
SDS 10% (v/v) 10 mL
H2O n/a 90 mL
Total n/a 100 mL
Tris 1 M pH 6.8
Reagent Final concentration Quantity
Tris 1 M 12.1 g
H2O n/a to 100 mL
Total n/a 100 mL
Tris 1 M pH 8.5
Reagent Final concentration Quantity
Tris 1 M 12.1 g
H2O n/a to 100 mL
Total n/a 100 mL
Tris 1.5 M pH 8.8
Reagent Final concentration Quantity
Tris 1.5 M 18.15 g
H2O n/a to 100 mL
Total n/a 100 mL
Equipment
AMAXA nucleofector (Lonza) or equivalent. Required for transfection of cells. See General note 1
Confocal microscope Leica TCS SPE-II (Leica) or equivalent. Required for imaging z-stacks to calculate volumes. See General note 1
Imager AI600 (Amersham) or equivalent. Required for imaging of SDS-PAGE gels (Epi-white light of 470–635 nm, any appropriate filter to see Coomassie stained proteins) and western blots (fluorescence epi light: 460, 520, or 630 nm with corresponding emission filters Cy2-525BP20, Cy3-605BP40, or Cy5-705BP40, see General note 2)
Mini-PROTEAN Tetra Vertical Electrophoresis Cell (Bio-Rad Laboratories) or equivalent
PowerPac Basic Power Supply (Bio-Rad Laboratories) or equivalent
S3e Cell Sorter (Bio-Rad Laboratories) or equivalent. It requires an illumination source and filters fitting to the fluorescent tag fused to the protein of interest: for EGFP, a 488 nm laser with emission filter 525/30 nm can be used
Single-molecule setup on a Nikon Eclipse Ti (Nikon). Used as example of fluorescence platforms. In the example given, EGFP images were taken using an OBIS 488 nm (100 mW) laser and a Quadbandpass (432/25 515/25 595/25 730/70 nm) filter from Nikon. See General note 1
Trans-Blot® SD Semi-Dry Transfer Cell (Bio-Rad Laboratories) or equivalent
Wide field microscope Axiovert 200 (Zeiss). Used as example of fluorescence platforms. In the example given, EGFP images were taken using a HBO100 bulb and a hard coated EGFP filter (ex: 482/18; bs: 495LP; em: 520/28). See General note 1
12 mm round coverslips (thickness 0.13–0.16 mm) (Diagonal, catalog number 41001112) or equivalent according to your microscope sample
#1.5H 24 mm round coverslips (thickness 175 ± 5 μm) (Thor laboratories, catalog number: CG15XH1). Used for live-cell microscopy in single-molecule setup. See General note 1
Widefield microscope Nikon Eclipse TiE2 (Nikon). Used for validation of the gate sorting across platforms. Spectra X LED 395 ± 25 nm (295 mW) and 370 ± 24 nm were used as illumination for DAPI and GFP respectively and a Quadbandpasss (432 ± 25, 515 ± 25, 595 ± 25, 730 ± 70 nm) as emission filter. Images were acquired using a 20× SPlan Fluor LWD DIC objective with numerical aperture of 0.7
Software and datasets
S3 ProSortTM Software (Bio-Rad Laboratories) or equivalent. Used to analyze and sort the cells in the cell sorter
ImageJ (FIJI) (Schindelin et al., 2012). In the examples given, FIJI version 1.52q was used and the plugin BioFormats was required to open the images generated in the widefield and confocal microscopes
Volocity 6.3 (Perkin-Elmer). See General note 1. ImageJ plugin 3D suite can be used instead to calculate volumes and intensity in z-stacks
Procedure
Protein purification and concentration validation by SDS-polyacrylamide gel electrophoresis (PAGE)
Purify the protein. You will need a purified version of the protein of interest (POI) that you want to study. It can be either untagged or tagged (see General note 3). Due to the many possibilities in protein purification, we will not include a description of the protein purification that we used, which can be found in the manuscript (Zhang et al., 2022).
Analyze the purified protein using SDS-PAGE.
Prepare dilutions with known quantities (250, 500, 750, 1,000, 1,500, 2,000 ng) of a reference protein (i.e., BSA) for a final volume of 15 μL in H2O.
Estimate the concentration of the purified protein by a colorimetric assay (e.g., PierceTM 660 nm Protein Assay Reagent).
Prepare specific quantities of the purified protein using the estimations (400, 600, 800, 1,000 ng) to a final volume of 15 μL in H2O.
Mix the dilutions of reference protein with 5 μL of loading buffer 4× to a final concentration 1×, boil them in 95 °C for 5 min, and then keep on ice.
Mix the different quantities of purified protein with 5 μL of loading buffer 4× to a final concentration 1×, boil them at 95 °C for 5 min, and then keep on ice.
Place a polyacrylamide gel in the electrophoresis chamber and fill it with 1× running buffer.
Analyze the samples by polyacrylamide gel electrophoresis with constant 100 V.
Incubate the gel in Coomassie staining solution overnight (~16 h).
Incubate the gel in destaining solution (2× 10 min) and equilibrate in H2O.
Image the gel in the imager (Figure 1A).
Figure 1. SDS-PAGE stained with Coomassie and analysis to validate the concentration of the purified protein. (A) Example of gel for validation. Known quantities of bovine serum albumin (BSA) are loaded together with different quantities of purified protein of interest (POI) estimated by colorimetric assay. Modified from Zhang et al. (2022). The white dashed line represents the selection used for quantification in B. (B) Intensity profile obtained in ImageJ using the gel analysis tool. Selections of the width of the band were done for individual bands and the area of the band of interest was calculated as the area under the peak. (C) Calibration curve based on the intensity measures obtained in B. The equation of the regression curve is described within the graph. (D) Calculations to obtain the corrected concentration (conc) of the POI, based on the intensity measurements of the POI bands using the equation of the regression curve from B, allow the determination of the corrected amount of POI from the calculated intensity of the band and the concentration in ng/μL as the average from the values obtained, 1624.6 ± 80.3 ng/μL. The values displayed are the calculation from left (POI 400 ng) to right (POI 1000) in the gel shown in A. The calculation of the error is described in the section data analysis. (E) Measurements and standard deviations for three technical replicates, each of them taking four band intensities to calculate the average as shown in D (which corresponds to left replicate).
Validate the concentration of purified protein by image analysis of the gel using ImageJ.
Generate a rectangular selection that covers a complete band.
Create a region of interest using the function Analyze → Gels → Select First Lane (for the first lane) and Analyze → Gels → Select Next Lane (for the others) (Figure 1A, white dashed line).
Get the intensity measurements using the function Analyze → Gels → Plot Lanes (Figure 1B).
Quantify the intensity of the bands as the area under the peak (Figure 1B). To do so: i) draw a horizontal lane in the base of the peak using the tool straight line so the peak is closed; ii) use the tool wand to select the peak; iii) automatically, it will display the area of the region selected.
Plot the intensity (area) against the known concentration of the reference protein and generate a regression curve (Figure 1C).
The equation of the regression curve is Y = a × X + b, being X the quantity of protein (in ng) and Y the intensity. Therefore, considering X = (Y - b)/a, you can calculate the protein amount based on intensity (Figure 1C). The parameter b corresponds to the background of the specific gel.
Use the intensity values obtained from the area under the peaks for each estimated amount of POI to get the actual values (Figure 1D).
You can get the concentration of the POI by dividing the amount of POI by the volume from the original solution added in each well (Figure 1D). The concentration of POI is the average of these values.
Calculation of protein amounts in cell fractions using fluorescence activated cell sorting and western blot
Sample preparation: grow the cells in the specific conditions required for its culture. You will need 106–107 cells in a 10 cm cell culture plate from wildtype (untransfected cells) and transfected cells. This step should be adapted to the specific cell line used. In the case of C2C12 using AMAXA electroporation:
Seed 50 cells/cm2 in a 10 cm cell culture plate.
Grow overnight in growth media at 37 °C with 5% CO2.
Remove media.
Wash cells with PBS-EDTA.
Add 2 mL of trypsin.
Incubate for 5 min at 37 °C.
Stop the trypsin with 4 mL of growth media.
Seed 0.5 mL of cells into a new 10 cm cell culture plate with 9 mL of growth media (untransfected sample).
Centrifuge 1 mL of cells at 0.3× g for 5 min and discard the supernatant.
Mix 2–5 μg of plasmid in 100 μL of room-temperature AMAXA M1 solution.
Resuspend the pellet in the AMAXA M1 solution containing the plasmid and collect in a cuvette.
Place the cuvette in the AMAXA nucleofector and select the program B-032.
Collect the cells and seed them in a new 10 cm cell culture plate with 10 mL of growth media (transfected sample).
Incubate the untransfected and transfected samples overnight at 37 °C with 5% CO2.
Sample processing; the individual plates are processed individually:
Wash cells with PBS.
Add 2 mL of trypsin.
Incubate for 5 min at 37 °C.
Collect the cells in a 15 mL tube.
Centrifuge at 0.3× g for 5 min and discard the supernatant.
Resuspend the pellet in 2 mL of PBS.
Centrifuge at 0.3× g for 5 min and discard the supernatant.
Resuspend the pellet in 3 mL of PBS.
Fluorescence-activated cell sorting (FACS)
Analyze the untransfected cells in analysis mode to determine the GFP intensity vs. the area and save the minimum and maximum intensity values. We recommend a count of approximately 106 cells for reproducibility. If the software of the FACS sorter does not provide the values, these can be inferred from the resulting graphs.
Analyze the transfected cells in analysis mode with the same settings as the untransfected cells and save the maximum intensity value. We recommend a count of approximately 106 cells for reproducibility. If the software of the FACS sorter does not provide the values, these can be inferred from the resulting graphs.
Use the acquired values for calculation of the gates (Table 1). See General note 4.
Table 1. Parameters for gate calculation based on intensities. *The use of logarithm is for presentation purposes and does not affect the following calculations.
Parameter Description Formula
MIN Minimum value of intensity in the untransfected sample log(min int untransfected)*
NEG_MAX Maximum value of intensity in the untransfected sample log(max int untransfected)*
POS_MAX Maximum value of intensity in the transfected sample log(max int transfected)*
BIN Reference interval (see General note 4) (POS_MAX-NEG_MAX)/40
POS_TH Threshold to define a positive count MIN + 11 * BIN
LOW_MIN Minimum threshold for the category “low” MIN + 13 * BIN
LOW_MAX Maximum threshold for the category “high” MIN + 22 * BIN
HIGH_MIN Minimum threshold for the category “low” MIN + 24 * BIN
HIGH_MAX Maximum threshold for the category “high” MIN + 33 * BIN
Sort the transfected cells into “low” and “high” based on the intensity values defined by the gates (Figure 2). Collect the cells and save the cell number count.
Figure 2. Example of western blot analysis to quantify the amount of protein of interest (POI) in FACS cell fractions. (A) Example of a gel for validation. Known quantities of purified POI are loaded together with lysates of known quantities of cells. Modified from Zhang et al. (2022). (B) Quantification of the bands of the gel using the gel tool analysis from ImageJ to determine the intensity of the purified POI and generate a calibration curve. (C) Combine the intensity measurements of the bands of lysates with the validation curve to obtain corrected amounts of total protein and, knowing the number of cells, the amount of protein per cell.
Extraction of total proteins from cells for western blot.
Centrifuge the cells from the sorted fractions at 0.3× g for 5 min and discard the supernatant.
Resuspend the pellet in 1 mL of lysis buffer with vigorous pipetting to disrupt the membranes.
Quantification of the amount of protein in sorted cells by western blot.
Prepare known quantities of purified POI (30, 60, 90, 120, 150, 180, 210, and 240 ng) according to the concentration calculated in A. See General note 5.
Mix the standard POI and the samples with a 4× loading buffer to a final concentration of 1× and boil them in 95 °C for 5 min. Clear the cell lysates by centrifugation at 10,000× g for 10 min and keep the supernatant on ice.
Analyze the fractions by SDS-PAGE as described before in steps A2e–A2f.
Prepare transfer buffer 1× by diluting 3 mL of transfer buffer 10× in 27 mL of H2O.
Cut Whatman papers and nitrocellulose membrane in the size of the gel and soak them in transfer buffer 1× for 15 min.
Wash the gel in H2O for 10 min.
Equilibrate the gel in transfer buffer 1× for 10 min.
Assemble the transfer unit into the transfer chamber (from down to up): 2× Whatman, nitrocellulose membrane, gel, 2× Whatman.
Remove the bubbles.
Transfer the proteins to a nitrocellulose membrane with constant 25 V for 20–45 min.
Incubate the membrane in 5% low-fat milk in PBS for 30–60 min at room temperature in a rotator.
Incubate with antibodies against your POI in the appropriate dilution overnight at 4 °C in a rotator.
Incubate with secondary antibodies linked to a fluorophore in the appropriate dilution in 3% low-fat milk in PBS for 1 h at room temperature in a rotator.
Detect the fluorescent signals using the imager (Figure 2A).
Measure the intensity in the bands using ImageJ as described before in steps A3a–A3d (Figure 1B).
Plot the intensity against the known concentrations to generate a regression curve.
Using the equation from the regression curve, calculate the protein amount for the “low” and “high” fractions of sorted cells. The intensities of the POI should be within the (linear) range of the calibration curve.
Divide the protein amount by the number of cells analyzed on the gel to obtain the number of molecules per cell.
Calculation of concentration (and use across imaging platforms)
Sample preparation. Grow the cells in the condition required in substrates suitable for imaging (i.e., chamber slides or coverslips). Follow the procedure described in step B1. See General note 6. The method is suitable for live or fixed cells. In case of fixed cells:
Remove media.
Wash the coverslips with PBS twice.
Incubate coverslips with formaldehyde solution 1:10 in PBS at room temperature for 10 min.
Wash the coverslips with PBS three times.
Add a 15 μL droplet of DAPI solution on parafilm and place the coverslips upside down. Incubate for 15 min at room temperature in darkness.
Turn the coverslips upside up.
Wash the coverslips with PBS three times.
Wash the coverslips once with H2O.
Add a 10 μL droplet of mounting media on a glass slide and place the coverslips upside down on it.
Let it dry in darkness.
Generation of calibration curve. See General note 6.
Acquire images of the cells in both negative and transfected cells with the same settings for the channel of the POI. Additionally, one can acquire additional channels to facilitate segmentations. In the examples given, we used a second channel with either DAPI (in fixed cells) or a protein that shows nuclear distribution (in live-cell imaging).
Segment the cells (or nuclei, in our case) using ImageJ (Figure 3A). In the example given, cells were segmented using the following ImageJ protocol: i) Process → Filters → Gaussian blur…, applying 2 pixels sigma (radius); ii) Process → Subtract background, applying 15 pixels of Rolling ball radius, and all other options not selected; iii) Image → Adjust → Threshold, using “Li” as threshold method. Images were then revised to find nucleus that were too close together and recognized as one to either discard them (when they overlapped) or separate into two regions of interest (when possible).
Calculate the total intensity in the channel of the POI for all cells. This value corresponds to the column “IntDen” in ImageJ result table and can be selected in Analyze → Set measurements…→ Integrated density.
Calculate the intensity values of the gates as described in Table 1 (Figure 3B).
Plot the frequencies of cells in range of the logarithm of intensity (Figure 3C).
Figure 3. Generation of a calibration curve for different instruments. (A) Scheme of the minimal analysis required to generate a calibration curve in a new imaging system. (B) Calculation of the gates based on the values of intensity obtained. Bins are calculated as 1/40 of the difference between the minimal (min) and maximal (max) values observed. (C) Examples of curves for the same POI in different imaging systems: FACS (left panel), widefield (middle-left panel), confocal (right-left panel), and single-molecule microscope (right panel). Increasing the number of cells imaged leads to higher resemblance to the curve used for quantifying the protein of interest (POI) by western blot (FACS). FACS, widefield, and single molecule are modified from Zhang et al. (2022). (D) Depiction of the effects of replicates in the generation of the gates. The error bar corresponding to the three biological replicates (with n > 500 each) used to generate the widefield graph in C, including the standard deviation of the calculation of the gates using the single datasets.
Check the quality of the calibration curve: all negative cells should be categorized as negative, and cells categorized as high should have a clear signal in the POI channel. See troubleshooting.
Calculation of concentration.
Acquire Z-stacks that contain the whole cellular structure where the POI is located, following the Nyquist sampling (i.e., in a confocal microscope), including the channel of the POI.
Segment the structure (additional channels can be used) and determine the intensity in the POI channel for the volume. See General note 8. In the example given, 3D segmentation was done in Volocity using the DAPI channel using the following protocol: I) Measure → Finding → Find objects; II) Measure → Processing → Dilate (iteration: 3); III) Measure → Processing → Fill holes in objects; IV) Measure → Processing → Erode (iteration: 3); V) Measure → Filtering → Exclude objects by size. The volume can be calculated from the Voxel count while the intensity is described in Sum, in the table that describes the ROIs.
Using the values for the gates of the calibration curve, classify the cells into low and high. See General note 7.
Calculate the average volume of the structure in the different categories.
The concentration of the POI for each category corresponds to the amount of protein per cell divided by the average volume of the structure for this category.
Data analysis
Note that, in the examples given, the protein concentration calculated for individual cells is classified in a broad category due to the selection of gates. However, the method can be as precise as it is possible to reduce the gate size and get enough cells to perform a measurable western blot. Even in this situation, there will be an error with three components: i) calculating the protein amount by western blot; ii) variability in cellular (or nuclear) volume; iii) error in the intensity measurements. Taking into account all these considerations, this method allows to assign single cells based on the POI fluorescence intensity to corresponding ranges of protein concentrations. To properly consider the mentioned errors in the final concentration number, error propagation should be taken into account at every step. In error propagation, given a function: f = a × b/y, in which the errors of the variables a, b, and y are ∂a, ∂b, and ∂c, respectively, the error of f (∂f) can be calculated as follows:
Here, you can find the sources of error and how to obtain error values:
Protein concentration for the purified protein in ng/μL (Figure 1) and in the fractions in μg/cell (Figure 2).
Intensity measurements: can be estimated as the average standard deviation of the background signal. To calculate the background signal, generate several ROIs in the same size between lines and obtain the intensities. This value corresponds to the column “IntDen” in ImageJ result table and can be selected in Analyze → Set measurements…→ Integrated density.
In replicate error: the standard deviation between the different lanes calculated (in the case of Figure 1).
Replicate error: standard deviation between technical and/or biological replicates plus the propagated error of the calculations.
Concentration of the protein in μM.
Protein concentration calculation as described above.
Volume calculation: standard deviation between cells.
In the example given in Figure 1, the background standard deviation, calculated as described in step 1a, is 15.793. Therefore, the concentrations in ng/μL for each line are 1,714.4 ± 6.8, 158.3 ± 5.5, 1,633.8 ± 4.8, and 1,601.8 ± 4.2. The final concentration number is then the average of the values, 1624.6, and the error corresponds to the standard deviation of the replicates (69.5), to which is added the propagation error of the calculations (10.8), giving the final number of 1,624.6 ± 80.3 ng/μL.
Validation of protocol
This method was validated in Zhang et al. (2022), corresponding to figures 3, supplementary figure 3, and supplementary tables S9 and S10.
For quantification, three replicates of transfected cells were sorted and analyzed with western blot to obtain the number of molecules. At least 50 cells from two biological replicates were used to calculate the volumes for the concentration for each condition.
Different calibrating curves were used in widefield microscopy (three biological replicates with approximately 500 cells per experiment) with comparable results of intensity values (Figure 3D).
To validate the use across platforms, we prepared samples as described in step B1. One-third of the cells (transfected or untransfected) were grown on coverslips and two-thirds were grown on a tissue culture dish. After 24 h, coverslips were collected, fixed with ice-cold methanol for 6 min, counterstained with DAPI, and mounted as described in steps C1d to C1i. All coverslips were imaged using a widefield microscope (Figure 4A). The DAPI channel was used for segmentation by processing in ImageJ (Gaussian blur, sigma 5 pixels; Subtract background, rolling, 20 pixels). Only nuclei with areas between 90 and 140 μm2 were considered based on our prior knowledge with C2C12 myoblasts. The calibration curve was done using the GFP intensities (Figure 4B, histogram) within the segmented nuclei. The cells grown on dishes were collected and sorted as described in steps B2 and B3, and the resulting fractions low and high were fixed with ice-cold methanol for 6 min, dried on to the coverslips by incubation at 37 °C overnight, and followed by counterstaining with DAPI and mounting as described before. These coverslips were imaged in the same conditions as the calibration (Figure 4A). The GFP intensity of the sorted cells was overlaid in the histogram (Figure 4B), resulting in segregation in the correct gates in most cases.
Figure 4. Validation of the reproducibility of the quantification across platforms. (A) Scheme of the validation process: cells (untransfected and transfected) were seeded in coverslips and, in parallel, a fraction of the transfected cells was sorted using fluorescence-activated cell sorting (FACS) in the categories low and high. The resulting fractions were fixed on coverslips by dehydration. All coverslips were counterstained with DAPI and imaged using a widefield microscope. DAPI was used for segmenting cell nuclei and quantifying total intensity of the GFP channel in the nucleus. (B) Non-sorted cells were used for determining the calibration curve, based on the frequencies (n untransfected: 63, n transfected: 1,539). Orange dots and magenta diamonds represent individual intensity values for the sorted low and high fraction, respectively.
General notes and troubleshooting
General notes
This material is not specifically needed for the protocol but has been used in the examples given.
Fluorescent or radioactive coupled secondary antibodies should be used for western blot to ensure linearity of the signal intensities. Enzymatic reactions (i.e., antibodies coupled to horseradish peroxidase) are not recommended.
Use tagged purified protein in cases where the protein of interest does not have suitable antibodies for western blot.
The selection of 40 bins as reference interval is meant to be versatile to generate different gate outcomes (number of categories) without changing the reference interval but can be adapted to the concrete experimental settings. Please note that the number of bins determines the range of variability that is accepted when setting gates. This step is the most susceptible to optimization when using other fluorophores and/or proteins.
The amount of protein of interest analyzed on the gel should be similar to the amount expected for the number of cells loaded. When the band corresponding to the cells is not within the calibration curve, you can modify the amounts of purified protein loaded for calibration and/or use a different number of cells.
You will need untransfected wildtype cells without a tag as negative control and transfected cells in the same conditions as described in section B for generating the calibration curve for each platform. The system is suitable for both live and fixed-cell microscopy, including immunostaining. The only requirement is that any further fluorescence, antibody, or other component should not interfere with the measurement of the intensity of the POI (i.e., overlapping of emission spectra with detection).
Each imaging instrument will require its own calibration curve. The accuracy of the measurements is highly dependent on the number of cells used for the calibration curve (Figure 3C), being ~200 cells the minimum to get reproducible results. The calibration curve can be used in the same platform, as long as the acquisition conditions (i.e., laser power, exposure time) remain constant between experiments. Please note that calibration curves should be different if single plane and volumes are used. This method should not be affected by any additional processing method (i.e., different deconvolution algorithm). If additional processing is used to determine intensities during the analysis, the images used for calibration must be treated the same way.
This section can be used to determine subcellular concentration for the POI by segmenting the subcellular structure instead of the whole cell.
Troubleshoot in calibration curves
Problem 1: Negative cells are included in the low gate.
Possible causes: (a) Dead cells with high autofluorescence have been segmented; (b) two or more cells have been segmented as one. This can be detected by plotting intensities vs. cell area; (c) bins are not accurate enough due to insufficient number of cells.
Solutions: (a) Revise the images and eliminate these cells; (b) revise the segmentation process; or (c) acquire more images from the same sample in the same conditions or merge several experiments imaged in the same conditions.
Problem 2: Transfected cells are included in the negative category.
Possible cause: Inappropriate imaging conditions (i.e., laser power or acquisition time) lead to saturation of high expressed cells.
Solution: Discard the dataset. If the dataset is extremely valuable, you can estimate the percentage of saturated cells compared to other datasets and displace the gates accordingly (note that this will add an imperfectible error!).
Acknowledgments
This work was financed by grants CA 198/10-1 project number 326470517 and CA 198/16-1 project number 425470807 to M.C.C. We thank Hui Zhang for his advice during the development of the method.
Competing interests
The authors declare no competing interests.
References
Chiu, C. S., Kartalov, E., Unger, M., Quake, S. and Lester, H. A. (2001). Single-molecule measurements calibrate green fluorescent protein surface densities on transparent beads for use with ‘knock-in’ animals and other expression systems. J. Neurosci. Methods 105(1): 55–63.
Dustin, M. L. (1997). Adhesive Bond Dynamics in Contacts between T Lymphocytes and Glass-supported Planar Bilayers Reconstituted with the Immunoglobulin-related Adhesion Molecule CD58. J. Biol. Chem. 272(25): 15782–15788.
Galush, W. J., Nye, J. A. and Groves, J. T. (2008). Quantitative Fluorescence Microscopy Using Supported Lipid Bilayer Standards. Biophys. J. 95(5): 2512–2519.
Jost, K. L., Rottach, A., Milden, M., Bertulat, B., Becker, A., Wolf, P., Sandoval, J., Petazzi, P., Huertas, D., Esteller, M., et al. (2011). Generation and Characterization of Rat and Mouse Monoclonal Antibodies Specific for MeCP2 and Their Use in X-Inactivation Studies. PLoS One 6(11): e26499.
Kudo, S., Nomura, Y., Segawa, M., Fujita, N., Nakao, M., Schanen, C. and Tamura, M. (2003). Heterogeneity in residual function of MeCP2 carrying missense mutations in the methyl CpG binding domain. J. Med. Genet. 40(7): 487–493.
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.
Sugiyama, Y., Kawabata, I., Sobue, K. and Okabe, S. (2005). Determination of absolute protein numbers in single synapses by a GFP-based calibration technique. Nat. Methods 2(9): 677–684.
Yaffe, D. and Saxel, O. (1977). Serial passaging and differentiation of myogenic cells isolated from dystrophic mouse muscle. Nature 270(5639): 725–727.
Zwier, J. M., Van Rooij, G. J., Hofstraat, J. W. and Brakenhoff, G. J. (2004). Image calibration in fluorescence microscopy. J. Microsc. 216(1): 15–24.
Zhang, H., Romero, H., Schmidt, A., Gagova, K., Qin, W., Bertulat, B., Lehmkuhl, A., Milden, M., Eck, M., Meckel, T., et al. (2022). MeCP2-induced heterochromatin organization is driven by oligomerization-based liquid–liquid phase separation and restricted by DNA methylation. Nucleus 13(1): 1–34.
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4,835 | https://bio-protocol.org/en/bpdetail?id=4835&type=0 | # Bio-Protocol Content
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Isolation and Analysis of B-cell Progenitors from Bone Marrow by Flow Cytometry
HZ Hongchang Zhao
RS Roger Sciammas
JB Jacqueline H. Barlow
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4835 Views: 969
Reviewed by: Chiara AmbrogioCatherine Hurd Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Dec 2022
Abstract
B cells play a critical role in host defense, producing antibodies in response to microbial infection. An inability to produce an effective antibody response leaves affected individuals prone to serious infection; therefore, proper B-cell development is essential to human health. B-cell development begins in the bone marrow and progresses through various stages until maturation occurs in the spleen. This process involves several sequential, complex events, starting with pre- and pro-B cells, which rearrange the heavy and light chain genes responsible for producing clonally diverse immunoglobulin (Ig) molecules. These cells then differentiate into immature B cells, followed by mature B cells. The bone marrow is a complex ecological niche of supporting stromal cells, extracellular matrix components, macrophages, and hematopoietic precursor cells influencing B-cell development, maturation, and differentiation. Once fully mature, B cells circulate in peripheral lymphoid organs and can respond to antigenic stimuli. As specific cell surface markers are expressed during each stage of B-cell development, researchers use flow cytometry as a powerful tool to evaluate developmental progression. In this protocol, we provide a step-by-step method for bone marrow isolation, cell staining, and data analysis. This tool will help researchers gain a deeper understanding of the progression of B-cell development and provide a pertinent flow gating strategy.
Keywords: B-cell development Lymphocyte staining Bone marrow isolation Adaptive immunity Flow cytometry
Background
Hematopoietic stem cells (HSCs) produce all cellular components of the blood through a series of unequal cellular divisions in the bone marrow. HSCs give rise to common myeloid progenitor and common lymphoid progenitor (CLP) cells that give rise to both T and B lymphocyte lineages. B cell–specific development starts when CLPs give rise to progenitor B cells. Progenitor B cells first differentiate into pro-B cells, which then undergo a series of cell division and differentiation events to create immature B cells (reviewed in [1,2]). Distinct surface receptors are expressed at each stage of B-cell development, allowing investigators to mark successive stages of development: B220 and CD43 are expressed on pre-pro-B cells, the earliest stage of B-cell commitment; CD19 expression becomes visible on early pro-B cells and is retained throughout development; finally, CD43 expression is lost on pre-B cells [3–7] (Figure 1).
Figure 1. Diagram of B-cell development. Major stages of adult B-cell development in the bone marrow focusing on the cell surface markers relevant to this protocol.
These stages are also defined by successive VDJ and VJ recombination events of the immunoglobulin heavy (IgH) and immunoglobulin light (IgL) chains gene loci, respectively, leading to IgM expression [8–10]. Upon successful assembly of the B-cell receptor (BCR), B cells migrate to secondary lymphoid organs and await their chance to encounter antigen. Antigen stimulation of the BCR results in a receptor signaling cascade that instructs cell activation pathways including changes in gene expression, G0–G1 progression, and bioenergetic metabolism in preparation for clonal expansion. Critically, the BCR simultaneously delivers antigenic cargo to endocytic processing compartments where proteolytic fragments are loaded onto major histocompatibility antigen class II (MHCII) and recycled to the cell surface as peptide:MHC complexes. T-cell recognition of such peptide:MHCII results in release of cell surface CD40L and secretion of the IL21 on activated T cells. Stimulation of CD40 and IL21R signaling on B cells by activated T cells licenses clonal expansion. Clonal expansion and coordinated changes in gene expression promote B-cell differentiation and B-cell fate choices into plasma cells, germinal center B cells, or memory B cells [11]. The distinct activity of these cell fates comprises the antibody response. Importantly, some clones will also undergo DNA class switch recombination, whereby the exons of the constant region segment (not the antigen combining site VDJ/VJ exons) are exchanged for a different one, e.g., μ to γ, ϵ, or α while retaining original antigen specificity. Antibodies of different isotypes exhibit different functionality, including fixation of complement, opsonization, or antibody-dependent cellular cytotoxicity to elaborate the general immune response. Because antibody responses are fundamental to host protection from pathogenic infections, the field has intensively studied the generation of B cells and their subsequent activation and differentiation. In this protocol, we focus on B-cell development and provide a time-tested flow cytometry–based assay for evaluating successive stages of B-cell development in the bone marrow [12–15]. Thus, this protocol will be useful for the evaluation of B-cell development in different mouse models as well as for the interrogation of discrete events in each stage [16]. This protocol describes the details of bone marrow isolation, cell surface antibody labeling to differentiate distinct developmental stages, and the flow cytometry gating strategy to quantify each population.
Materials and reagents
Scissors (VWR, catalog number: 82027-578)
Tweezers (VWR, catalog number: 82027-398)
Dissection pins (VWR, catalog number: 76549-022)
70% ethanol
Kimwipe task wipers (Kimtech brand, Amazon)
Pipette tips (non-filter)
18 G syringe needle (Fisher Scientific, catalog number: 14-826)
0.22 μm syringe filter (Fisher Scientific, catalog number: SLHP033RS)
Biohazard bag(s)
CO2 tank
0.5 mL microcentrifuge tubes (VWR, catalog number: 0011-830)
1.5 mL microcentrifuge tubes (Genesee Scientific, catalog number: 24-281)
70 μm mesh net cut into ~1.5 cm squares to cover tube opening (Component Supply Company Inc, U-CMN-70); 70 μm filters can be used instead (Fisher Scientific, catalog number: 07201431)
Bovine serum albumin (BSA) (Sigma, catalog number: BP1600)
NaCl (Sigma, catalog number: BP358)
KCl (Sigma, catalog number: P9541)
Na2HPO4 (Sigma, catalog number: S374500)
KH2PO4 (Sigma, catalog number: P9791)
Sodium azide (Sigma, catalog number: BP922I)
NH4Cl (Sigma, catalog number: A661)
KHCO3 (Sigma, catalog number: P235)
EDTA-Na2 (Fisher BioReagents, catalog number: BP120-1)
Counting beads (Spherotech, catalog number: ACBP-50-10)
Mice (C57BL/6, C57BL/6 × 129/Sv)
Anti-mouse CD16/CD32 (Fc shield) antibody (2.4G2) [Tonbo (now Cytek), catalog number: 70-0161-U100]
Rat anti-mouse monoclonal, APC CD19 (BD Pharmingen, catalog number: 561738)
Rat anti-mouse monoclonal, PE-Cy7-CD43 (BD Pharmingen, catalog number: 562866)
Rat anti-mouse monoclonal, PE-IgM (BD Pharmingen, catalog number: 562033)
Rat anti-mouse monoclonal, BV421-CD11b (BD Pharmingen, catalog number: 562605)
Rat anti-mouse monoclonal, PerCP-CD45R/B220 (BD Pharmingen, catalog number: 553093)
Wash buffer (see Recipes)
1× PBS buffer (see Recipes)
ACK (ammonium-chloride-potassium) lysis buffer (see Recipes)
Recipes
Wash buffer
Sterile PBS (1×)
1% BSA
0.05% sodium azide
Dissolve 1 g of BSA powder and 0.05 g of sodium azide in 100 mL of PBS (1×) to make the wash solution. Filter the solution through a 0.22 μm syringe filter and store it at 4 °C for up to one week.
1× PBS buffer
NaCl 137 mM
KCl 2.7 mM
Na2HPO4 10 mM
KH2PO4 1.8 mM
To prepare 1 L of 1× PBS, dissolve the reagents listed above in 800 mL of H2O. Adjust the pH to 7.4 with HCl, then add H2O to 1 L. Sterilize by autoclaving for 20 min and store at room temperature (RT) for up to one year.
ACK buffer
8.29 g of NH4Cl (0.15 M final concentration)
1 g of KHCO3 (1 mM final concentration)
37.2 mg of Na2EDTA (0.1 mM final concentration)
Add 800 mL of H2O and adjust the pH to 7.2–7.4 with 1 N HCl (sterilize by filtration using a 0.2 μm filter)
Store at RT for up to one year.
Equipment
Vortex (VWR, model: 88880017)
Centrifuge (Eppendorf, model: 5424R)
Pipettes (Eppendorf)
Tissue culture light microscope equipped with brightfield and 20× objective (Olympus CK-40)
Flow cytometer (LSR II, BD)
Software
FlowJo software (FlowJo, LLC.)
Prism 6 software (GraphPad Software, Inc.)
Procedure
Euthanize the mice and isolate the femur
Euthanize the mice with CO2 under standard protocol [17] or to the specifications of the users’ mouse facility.
Pin the mouse onto a foam board with needles in a supine position (Figure 2).
Figure 2. Mouse dissection diagram focusing on the location and isolation of intact femur
Spray the mouse with 70% ethanol.
Make an incision on the hind leg and extend it from foot to groin.
Pull the skin to the back and cut it to expose the muscle.
Carefully remove the muscle with scissors from knee joint to groin joint.
Cut the knee joint and groin joint with scissors.
Isolate the femur containing the kneecap and femoral head.
Note: Keep the integrity of the kneecap and femoral head (rounded with less opaque cartilage) to make sure the bone marrow is not released from femur.
Remove extra muscle tissue with Kimwipes so there is no lint.
Repeat with second hind leg to isolate two femurs in total.
Discard mouse carcass according to UC Davis Institutional Animal Care and Use Committee (IACUC protocol #21828).
Bone marrow isolation
Puncture the bottom of a 0.5 mL microcentrifuge tube by hand from the inside of the tube with an 18 G needle and place it into a 1.5 mL microcentrifuge tube.
Note: Take care and wear thick protective gloves.
Cut both ends of the femur with scissors and place them in the 0.5 mL tube.
Centrifuge the tube at 2,000× g for 10 s at RT.
Discard the 0.5 mL centrifuge tube.
Resuspend the bone marrow cells with ACK lysis buffer for 2 min at RT.
Centrifuge the cells at 500× g for 5 min at RT.
Throw away the supernatant, resuspend the cells with 1 mL of wash buffer, and filter the cells through the mesh net.
Count the cells. Potential stop point: Isolated bone marrow cells can be stored at 4 °C for a few hours.
Note: We recommend preparing 4 million cells per staining. Less than 1 million cells can be used per staining; however, this may cause quantification problems after flow cytometry.
B-cell staining
Prepare antibody staining solutions: dilute CD11B, B220, IgM, and CD19 antibodies 1:99 in wash buffer; note that the CD43 antibody is not diluted. The fluorochrome combination used below was optimized for use on cytometers (BD Fortessa and Sony SH800S) with at least three lasers: blue (488 nm), red (640 nm), and violet (405 nm). Fluorochrome combinations should be chosen based on the instrument used and its laser availability; consultation with flow core personnel to minimize emission overlap is strongly recommended.
Pipette the cell suspension obtained after isolation into individual sample tubes; we recommend 4 × 106 cells for each experimental sample. For the full five antibody staining, prepare 11 additional control samples following the table below [no stain, single antibody staining, multi-antibody staining, and fluorescence minus one (FMO) staining controls].
Adjust the volume with wash buffer to 100 μL/sample.
Add 10 μL (1:99 diluted) of anti-FC blocking antibody to each staining tube.
Add 10 μL of prepared antibody solution for CD11B, B220, IgM, and CD19 to all experimental samples and the relevant control tubes. Add 1 μL of undiluted CD43 antibody to all experimental samples and the relevant control tubes.
Incubate in the dark for at least 1 h.
Centrifuge the sample at 1,000× g for 2 min at RT.
Wash twice with 1 mL of wash buffer and resuspend with 490 μL of wash buffer.
Add 10 μL of counting beads (invert several times before pipetting the beads) before flow cytometry.
Table 1. Representative sample panel for five color staining including all control samples necessary for compensation. No stain (no fluorescent antibody, blocking antibody only), S (single antibody staining), FMO (fluorescence minus one staining). *Volume of antibody used relative to the original concentration from the manufacturer.
Staining combination
Antibody Volume* No stain S1 S2 S3 S4 S5 Testing sample FMO1 FMO2 FMO3 FMO4 FMO5
BV421-CD11B 0.1 μL - + - - - - + - + + + +
FITC-B220 0.1 μL - - + - - - + + - + + +
PE-IgM 0.1 μL - - - + - - + + + - + +
APC-Cy7-CD43 1 μL - - - - + - + + + + - +
APC-CD19 0.1 μL - - - - - + + + + + + -
Flow cytometry operation
The no stain sample is used to set laser voltages, the single-color staining samples are used to compensate fluorochrome emission overlap, and the FMO samples are used to provide empirical evidence that the instrument was well compensated. Fluorochrome emission is often broad and the broad emission spectra overlap potentially creating false positive signals. Instrument compensation, which superficially is a mathematical-based subtraction of fluorescence signal in a distinct channel, is essential for proper interpretation of the results. In the preliminary run, perform flow cytometry with an experienced user or flow core manager to help with compensation so samples are not over- or under-compensated.
When running each single-color-stained sample, check that other desired channels display minimal emission, i.e., if a sample is stained with an APC-Cy7-conjugated antibody alone, there should not be any signal in the FITC channel. If there is, then instrument compensation is in order and entails increasing compensation such that detection in the FITC channel is eliminated; in this case, the mean fluorescence intensity on the FITC axis of the APC-Cy7-stained population is equivalent to the non-APC-Cy7-stained population. Repeat with each single-color-stained sample against all other desired channels (Figure 3).
Figure 3. Representative flow cytometry graphs post-compensation using the APC-Cy7-CD43 staining. Isolated bone marrow B cells were stained with APC-Cy7-CD43 antibody following the protocol, to determine if it is detected by other channels being used (FITC, PE, BV-421, and PE); every channel is checked separately. This confirms the APC-Cy7-CD43 positive (69.5%) population is being identified in the appropriate channel, while compensation in other single channels eliminates/strongly reduces the bleed-through signal from overlap in spectral emission.
Run and record all samples, including all controls, after compensation adjustment.
Note: Current analysis software can use digital compensation to adjust compensation post-sample acquisition; however, given that laser voltage is intimately correlated with emission, i.e., higher voltages result in greater emission and thus more overlap, performing pre-acquisition compensation is an important practice for result quality. Recording all controls maintains a record of the staining quality and allows for digital re-adjustment of compensation later.
Data analysis using FlowJo software
Gate the beads and the cells according to Figure 4. If beads are included, one can calculate the absolute number of cells of a given population that is designated by a given gate. This is helpful in understanding whether frequencies of developmental progenitors are unchanged, but progeny output is diminished. Specifically, the absolute number of B cells at different stages of development is calculated by: (number of events for the test samples/number of events for the counting beads) × (number of beads used/volume of test sample initially used).
Figure 4. Representative flow cytometry plots for each stage of the gating pipeline for data analysis. Sample is from a wildtype mouse.
Gating pipeline (see Figure 4):
Step 1. Gate on live large cells using the forward and side scatter channels (FSC and SSC, respectively).
Avoid fragmented and dead cells.
*Optional: if quantitation is desired, gate beads to calculate absolute cell numbers.
Step 2. Distinction between myeloid vs. lymphoid cells is defined here as CD11B+/CD11B-.
CD11B+ cells: myeloid—should be low for B220 signal.
Lymphoid cells: CD11B-negative—these are further gated for B-cell developmental stages.
Step 3. Distinction between pre-pro B cells and later stages is defined here by the density of B220 and CD19 expression within the CD11B- population.
B220+ CD19- are pre-pro B cells.
B220+ CD19+ comprise a mixture of more differentiated B cells (steps 4 and 5 below).
Step 4. Pro-, small pre-, large pre-, and mature B cells are distinguished based on IgM expression within the B220+ CD19+ population.
IgM- cells comprise pro-, small pre-, and large pre-B cells based on CD43 and FSC parameters (steps 5 and 6).
IgM+ comprise immature and recirculating mature B cells based on their B220 density (steps 7 and 8, respectively).
Step 5. Pro-, small pre-, and large pre-B cells are distinguished between levels of CD43 expression.
Pro-B cells display the greatest CD43 expression.
Small pre- and large pre-B cells comprise the CD43- population and can be distinguished by size (FSC); see step 6.
Step 6. Small and large pre-B cells are distinguished based on the FSC spread.
Large pre-B cells exhibit the greatest FSC; these cells express a complex of a rearranged heavy chain paired with surrogate light chain, also known as the pre-BCR.
Small pre-B cells display the lowest FSC and follow large pre-B cells and are in the process of rearranging the VJ segments of the light chain.
Step 7. Mature recirculating B cells have downregulated CD43 expression among cells defined by the gate in step 4b; B220Hi CD19+ IgM+ cells.
Step 8. Immature B cells that express a functional surface BCR and are migrating to secondary lymphoid organs have downregulated CD43 expression among cells defined by the gate in step 4b; B220Low CD19+ IgM+ cells.
Validation of protocol
The protocol presented here used data acquired from the bone marrow of a wildtype mouse (Figures 3 and 4) and was used in [16].
Acknowledgments
This work was supported by research funding from the National Cancer Institute K22CA188106, University of California Cancer Research Coordinating Committee (UC-CRCC) seed grant CRR-20-635379, and National Institute for General Medical Studies grants R01 GM134537 (JHB), and R21 AI151610 (RS). This study utilized the University of California Davis Cancer Center Flow Cytometry core partially supported by National Institute of Health grant S100D018223. Thanks to Bridget McLaughlin and Jonathan Van Dyke of the cancer Center Flow Cytometry core for their assistance with data acquisition. Use of the BD Fortessa was supported by the Flow Cytometry Core of the California National Primate Research Center under NIH award number P51OD011107.
This protocol was derived from the original work [16].
Competing interests
The authors report no competing interests.
References
LeBien, T. W. and Tedder, T. F. B lymphocytes: how they develop and function. (2008). Blood 112(5): 1570–1580. doi:10.1182/blood-2008-02-078071
Pieper, K., Grimbacher, B. and Eibel, H. B-cell biology and development. (2013). J. Allergy Clin. Immunol. 131(4): 959–971. doi:10.1016/j.jaci.2013.01.046
Wells, S. M., Kantor, A. B. and Stall, A. M. (1994). CD43 (S7) expression identifies peripheral B cell subsets. J. Immunol. 153(12): 5503–5515. doi: 10.4049/jimmunol.153.12.5503
Perez-Andres, M., Grosserichter-Wagener, C., Teodosio, C., van Dongen, J. J., Orfao, A. and van Zelm, M. C. (2011). The nature of circulating CD27+CD43+ B cells. J. Exp. Med. 208(13): 2565–2566. doi: 10.1084/jem.20112203
Hardy, R. R., Carmack, C. E., Shinton, S. A., Kemp, J. D. and Hayakawa, K. (1991). Resolution and characterization of pro-B and pre-pro-B cell stages in normal mouse bone marrow. J. Exp. Med. 173(5): 1213–1225. doi: 10.1084/jem.173.5.1213
Wang, K., Wei, G. and Liu, D. (2012). CD19: a biomarker for B cell development, lymphoma diagnosis and therapy. Exp. Hematol. Oncol. 1(1): e1186/2162-3619-1-36. doi: 10.1186/2162-3619-1-36
Otero, D. C. and Rickert, R. C. (2003). CD19 Function in Early and Late B Cell Development. II. CD19 Facilitates the Pro-B/Pre-B Transition. J. Immunol. 171(11): 5921–5930. doi: 10.4049/jimmunol.171.11.5921
Melchers, F. (2015). Checkpoints that control B cell development. J. Clin. Invest. 125(6): 2203–2210. doi: 10.1172/jci78083
Weill, J. C. and Reynaud, C. A. (2020). IgM memory B cells: specific effectors of innate-like and adaptive responses. Curr. Opin. Immunol. 63: 1–6. doi: 10.1016/j.coi.2019.09.003
Capolunghi, F., Rosado, M. M., Sinibaldi, M., Aranburu, A. and Carsetti, R. (2013). Why do we need IgM memory B cells? Immunol. Lett. 152(2): 114–120. doi: 10.1016/j.imlet.2013.04.007
Akkaya, M., Kwak, K. and Pierce, S. K. (2020). B cell memory: building two walls of protection against pathogens. Nat. Rev. Immunol. 20(4): 229–238. doi: 10.1038/s41577-019-0244-2
Hardy, R. R. and Hayakawa, K. (2001). B cell development pathways. Annu. Rev. Immunol. 19, 595–621. doi:10.1146/annurev.immunol.19.1.595
Loffert, D., Schaal, S., Ehlich, A., Hardy, R. R., Zou, Y. R., Muller, W. and Rajewsky, K. (1994). Early B-Cell Development in the Mouse: Insights from Mutations Introduced by Gene Targeting. Immunol. Rev. 137(1): 135–153. doi: 10.1111/j.1600-065x.1994.tb00662.x
Melchers, F., Karasuyama, H., Haasner, D., Bauer, S., Kudo, A., Sakaguchi, N., Jameson, B. and Rolink, A. (1993). The surrogate light chain in B-cell development. Immunol. Today 14(2): 60–68. doi: 10.1016/0167-5699(93)90060-x
Lu, L. and Osmond, D. G. (2000). Apoptosis and its modulation during B lymphopoiesis in mouse bone marrow. Immunol. Rev. 175(1): 158–174. doi: 10.1111/j.1600-065x.2000.imr017506.x
Zhao, H., Hartono, S. R., de Vera, K. M. F., Yu, Z., Satchi, K., Zhao, T., Sciammas, R., Sanz, L., Chédin, F., Barlow, J., et al. (2022). Senataxin and RNase H2 act redundantly to suppress genome instability during class switch recombination. eLife 11: e78917. doi: 10.7554/elife.78917
Raj, A. B., Leach, M. C. and Morton, D. B. (2004). Carbon dioxide for euthanasia of laboratory animals. Comp. Med. 54(5): 470–471. https://pubmed.ncbi.nlm.nih.gov/15575359/
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Cell Biology > Cell staining > Whole cell
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4,836 | https://bio-protocol.org/en/bpdetail?id=4836&type=0 | # Bio-Protocol Content
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A Rapid and Simple Procedure for the Isolation of Embryonic Cells from Fish Eggs
VG Vasily Golotin
AL Anatoly Lyutikov
TF Tatiana Filatova
VS Vladimir Sharoyko
OA Olga Apalikova
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4836 Views: 383
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Abstract
Fertilized teleost fish eggs are a complex formation, in which dividing cells arelocated in a small point in the entire volume of eggs. Isolating embryonic cellscan be considered a necessary step in the research of developmentalpeculiarities of fish cells at the earliest stages of embryogenesis beforeembryo formation. The main advantages of the offered protocol are rapidisolation, no enzymes, and overall low cost compared to other protocols. Theprotocol is suitable for the isolation of embryonic cells from medium-sized eggsat the stages of blastula or gastrula, for studies in a variety of applications(e.g., microscopy, flow cytometry, and other methods). Fertilized nelma eggs(Stenodus leucichthys nelma) are used in the protocol as a model.
Key features
• Fast and cheap isolation of cells from fish eggs at early stages (blastula orgastrula).
• Applicable for most of the methods for cell study (any staining, microscopy, flowcytometry, etc.).
• Can be applied to other teleost fish eggs with medium egg diameter of 3–4mm.
Graphical overview
Keywords: Aquaculture Stenodus leucichthys nelma Blastodisc Cell isolation Flow cytometry Propidium iodide Microscopy
Background
To date, many methods for isolating living cells from various tissues are used fordifferent purposes. Mainly, living cells from tissues are disaggregated bytrypsinization. This method is very common and involves the extraction of an embryoor tissue with further trypsinization (Durkin et al.,2013). Some other protocols are based on mechanical crushing of tissues orformed embryos to obtain cells (Fetherman et al., 2015).
When rapid testing of samples is required as soon as possible after eggfertilization, it is necessary to isolate cells directly from the blastodisc at theearliest stages of teleost fish embryogenesis before formation of embryo, larva, orfry. Cell isolation from fertilized eggs by the methods described above does notlead to the desired result, since the eggs contain huge amounts of varioussubstances and a small number of cells relative to the weight of the egg. Inaddition, the cells in the blastodisc are not tightly connected with each other.That is why the use of trypsin (and other enzymes) is not ideal, with undesirablechemical effects on the cells that can lead to the destruction of cell membranes.
To date, we have only found one other protocol describing a similar procedure (Rieger, 2019). That protocol requires thepresence of pronase from Streptomyces griseus for thedechorionization of embryos. The pronase only softens the chondrion, requiringadditional washing to remove it from the embryo. Besides, the isolation procedure iscomplicated by particular features of enzymes, as most biological catalysts havenarrow operating limits as well as a short shelf life. We aimed to ensure maximumsimplicity and low cost of the isolation method with minimal requirements forlaboratory equipment.
Materials and reagents
Biological materials
Fertilized fish eggs in the stages from blastodisc to the late gastrula.
Solutions
PBS soluble tablets (Sigma-Aldrich, catalog number: P4417), for cell isolation
70% ethanol solution (Sigma-Aldrich, catalog number: 65348-85), for cell preservation
o-safranin ready-to-use solution 0.1% (Scientific Laboratory Supplies, catalog number: TMS-009-C), for protocol verification
Propidium iodide ready-to-use solution 1 mg/mL (Thermo Fisher Scientific, catalog number: P3566), for protocol verification
RNAse A ready-to-use solution 10 mg/mL (Thermo Fisher Scientific, catalog number: EN0531)
Phosphate-saline buffer solution (see Recipes)
70% ethanol (see Recipes)
Recipes
Phosphate-saline buffer solution
Reagent Final concentration Quantity
PBS
150 mM total of all salts:
137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, 2 mM KH2PO4
1 tablet
H2O n/a Up to 100 mL
Total n/a 100 mL
70% ethanol
Reagent Final concentration Quantity
Ethanol (95%) 70% 74 mL
H2O n/a 26 mL
Total n/a 100 mL
Laboratory supplies
Microcentrifuge tube pestles (Sigma-Aldrich, catalog number: BAF199230001-100EA)
Conical bottom microcentrifuge tubes (Sigma-Aldrich, catalog number: HS4325-1000EA)
Centrifuge tubes (Sigma-Aldrich, catalog number: CLS430790)
Equipment
Microcentrifuge (Thermo Fisher Scientific, catalog number: 75004081)
Microscope with 8–20× zoom lens for data validation (Thermo Fisher Scientific, catalog number: 4479672)
Vortex mixer (VELP Scientifica, catalog number: F202A0173)
Dissecting needles (Thermo Fisher Scientific, catalog number: 13-820-024)
Flow cytometer for data validation (Beckman Coulter, catalog number: CO9752)
Software and datasets
CytExpert software for flow cytometry (used for data validation)
Procedure
This protocol has been tested on fertilized nelma eggs of medium size (3–4 mm in diameter) among teleost fish (including zebrafish). Before the procedure, rinse the eggs with PBS: place the eggs in a centrifuge tube (15 mL) and pour 10–20 mL of cool PBS solution into it. Drain the liquid and repeat this step to prevent any contamination that could affect further analysis. This step can also be done in any way available in the laboratory: using a tea strainer, beakers, gauze, etc.
Using a pestle, pop the eggs (or single egg) in the tube. It is better to add eggs and process them gradually. Do not homogenize nor strongly push the pestle to avoid damage to the cells: it is enough to break the shell. The sample volume from the processed eggs should not exceed 2/3 of the tube. A 1.5 mL centrifuge tube can be fitted with a different number of eggs, depending on their size. Thus, approximately 30 eggs of 3–4 mm diameter can be sequentially placed in the tube during the process.
Fill the tube with eggs with cold PBS and shake it on a vortex for 5 s or mechanically using a pestle or dissecting needle to release the contents of all eggs outside the visible shell.
Mechanically remove all shells from the tube using a dissecting needle. It is very important to ensure that there are no residual shells in the resulting translucent medium to avoid cell loss.
Centrifuge at 300–500× g for 5 min at 4 °C.
Discard the supernatant without affecting the precipitate. Add 1–1.5 mL of cold PBS and centrifuge again.
(Optional) Repeat step 6 1–2 times to wash the cells out of small debris and resuspend the pellet in 100 μL of cold PBS.
Following these steps, we obtain a pellet containing the cells. The volume of PBS can be changed as desired (50–500 μL). This suspension contains a certain amount of small suspended non-cellular particles. After the procedure, the total proportion of cells among other floating non-cellular particles in the suspension is at least 10%.
The cell suspension is ready to use for any analysis. It can also be fixed for long-term cell storage using standard protocols such as using chilled ethanol (Ciancio et al., 1988).
Data analysis
Flow cytometry and microscopy were used for protocol verification. The CytExpert software was used for all cytometry data.
Validation of protocol
Maturating nelma eggs (Stenodus leucichthys nelma) were used to validate the protocol (Figure 1A).
Figure 1. Maturating Stenodus leucichthys nelma egg. (A) White arrow shows the position of the blastodisc. (B) Isolated cells from the blastodisc stained with O-safranin.
Thirty fish eggs were processed by the protocol described above. Then, the cell-containing pellet was resuspended in 300 μL of PBS and fixated in 70% ethanol (Ciancio et al., 1988) to ensure long-term preservation prior to analysis. A small 10 μL aliquot of the obtained suspension was stained with safranin (Swain and De, 1990). The suspension was air-dried at room temperature on a glass slide causing the cells to adhere to it. Then, the slide was immersed in a solution of safranin for 5 min and washed one time with PBS. The result was checked using a light microscope (Figure 1B ).
In order to verify the results and additionally show the real presence of isolated cells, an alcohol suspension of the obtained cells (300 μL) was stained with propidium iodide (Riccardi and Nicoletti, 2006). The alcohol was washed off the cells twice with PBS by centrifugation at 300× g for 5 min; cells were then resuspended in 100 μL of PBS (+0.1 μg/mL RNase A and 0.01 μg/mL propidium iodide). After 30 min of incubation in the dark at 4 °C, stained cells were analyzed using a flow cytometer (channel PE) (Figure 2). The flow cytometer analyzed 500,000 events. Using a software interface, side scatter, and forward scatter data, only single cells were selected for analysis. The histogram along the PE channel clearly showed peaks corresponding to the cell cycle of isolated cells (Figure 2A). The vertical line on the dot plot (Figure 2B) showed the position corresponding to the G1 peak of the cell cycle in the histogram (Figure 2A). On the dot plot diagram (Figure 2B) for the same channel, there is size cell dispersion (y-axis) that corresponds to the stage of embryogenesis (small cell blastula) where cells acquire different sizes (Figure 2B). According to our data, approximately 100,000 cells were isolated (Figure 2B) from 30 eggs (Figure 1A), which was quite commensurate with other known methods.
Figure 2. Results of flow cytofluorometry (PE channel). Cell cycle histogram (A) and dot plot (B) showing the heterogeneity of the cell analyzed from small cell blastula.
Thus, the protocol was tested in two independent ways: visual (microscopic) and photometric. The procedure efficiency for cell isolation from fish egg blastodiscs was proved. The protocol can be applied to most teleost fish species with middle-sized eggs at early maturating stages (blastula, gastrula); this protocol may be suitable for other representatives of teleost fish with different diameters of eggs [large (salmon) or small (zebrafish)].
General notes and troubleshooting
General notes
Before you start: to isolate cells from the blastodisc, it is enough to have developing eggs at the stage of large cell blastula. Based on specific goals, it is important to mind the number of cells needed for analysis. If it is necessary to analyze each egg individually, it is important to know the total number of required cells in the developing egg. For examining a specific number of eggs (if general statistics on the target analysis is needed), the stage of development of the eggs is not so important.
Troubleshooting
No cells after procedure:
Take more eggs for processing and repeat the procedure.
Ensure that you have taken the developing eggs. An embryo should be visible under a light microscope.
Remember that in the first step of the isolation protocol, after rinsing the eggs, it is sufficient to gently pop the eggs. Do not completely homogenize.
Acknowledgments
This study was supported by the RSF grant 23-26-00257.
Competing interests
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
References
Ciancio, G., Pollack, A., Taupier, M. A., Block, N. L. and Irvin, G. L. (1988). Measurement of cell-cycle phase-specific cell death using Hoechst 33342 and propidium iodide: preservation by ethanol fixation. J. Histochem. Cytochem. 36(9): 1147–1152.
Durkin, M., Qian, X., Popescu, N. and Lowy, D. (2013). Isolation of Mouse Embryo Fibroblasts. Bio Protoc 3(18): e908.
Fetherman, E. R., Lepak, J. M., Brown, B. L. and Harris, D. J. (2015). Optimizing Time of Initiation for Triploid Walleye Production Using Pressure Shock Treatment. North Am. J. Aquacult. 77(4): 471–477.
Riccardi, C. and Nicoletti, I. (2006). Analysis of apoptosis by propidium iodide staining and flow cytometry. Nat. Protoc. 1(3): 1458–1461.
Rieger, S. (2019). Dechorionation of zebrafish embryos with Pronase for metronidazole-mediated ß-cell ablation V.2. DiaComp Protocols.
Swain, D. and De, D. N. (1990). Differential Staining of the Cell Cycle of Plant Cells Using Safranin and Indigo-Picrocarmine. Stain Technol. 65(4): 197–204.
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Developmental Biology
Cell Biology > Cell isolation and culture > Cell isolation
Biological Sciences > Biological techniques
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4,837 | https://bio-protocol.org/en/bpdetail?id=4837&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Co-culture Wood Block Decay Test with Bacteria and Wood Rotting Fungi to Analyse Synergism/Antagonism during Wood Degradation
JE Julia Embacher *
SZ Susanne Zeilinger
SN Sigrid Neuhauser
MK Martin Kirchmair *
(*contributed equally to this work)
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4837 Views: 417
Reviewed by: Xiaofei LiangYufang LuKristin L. Shingler
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Abstract
Mixed communities of fungi and bacteria have been shown to be more efficient in degrading wood than fungi alone. Some standardised protocols for quantification of the wood decay ability of fungi have been developed (e.g., DIN V ENV 12038:2002 as the legal standard to test for the resistance of wood against wood-destroying basidiomycetes in Germany). Here, we describe a step-by-step protocol developed from the official standard DIN V ENV12038 to test combinations of bacteria and fungi for their combined wood degradation ability. Equally sized wood blocks are inoculated with wood decay fungi and bacterial strains. Axenic controls allow the analysis of varying degradation rates via comparison of the wood dry weights at the end of the experiments. This protocol provides new opportunities in exploration of inter- and intra-kingdom interactions in the wood-related environment and forms the basis for microcosm experiments.
Key features
• Quantification of wood decay ability of mixed cultures.
• Allows testing if fungi are more efficient in degrading wood when bacteria are present.
Keywords: ENV 12038 standard Wood decay Bacterial–fungal interactions Synergism/Antagonism Serpula lacrymans
Background
In nature, basidiomycetous fungi are associated with other microbes including prokaryotes and eukaryotes. This results in complex competitive and antagonistic interactions as well as commensal and mutualistic behaviour (Boer et al., 2005; Kobayashi and Crouch, 2009). Fungi are the most efficient wood decomposers, as their multicellular appearance and hyphal growth harbour a mobility advantage in comparison to prokaryotes. Nonetheless, bacteria are known to have direct influence on the decay process as well, by e.g., degrading complex wood components like cellulose, lignin, and hemicellulose (McGuire and Treseder, 2010) or by altering wood permeability and structure, thus improving accessibility of the wooden microfibrils (Clausen, 1996), aiding other organisms in wood decay. Synergistically acting species benefit from each other’s enzymatic abilities when they are cultivated on wood (Cortes-Tolalpa et al., 2017; Sugano et al., 2021). These results highlight the importance of inter- and intra-kingdom interactions, as they show that the combined action of different species enhances the decay ability of the whole community.
To date, several protocols and studies exist for the assessment of wood resistance against wood-destroying basidiomycetes [e.g., DIN V ENV 12038:2002 (German Institute for Standardisation, 2002), wood block test of Bravery (1978), or work of Lohwag (1965) and Hegarty et al. (1987)]. However, as these protocols evolved in large part to study timber preservation and were not designed to study the influence of other microbes, we adapted a protocol based on DIN V ENV 12038:2002 to the here described co-culture wood block decay test. This step-by-step protocol provides an easily feasible standard procedure for the assessment of wood decay properties of mixed cultures in comparison to axenic decay rates by evaluating the dry weight of wood blocks that were exposed to microbial deterioration. The purpose of this wood decay assay is to determine the influence of bacteria on the decay properties of wood rotting fungi like Serpula lacrymans [see e.g., Embacher et al. (2022), article in prep.].
Materials and reagents
Fungal and bacterial strains
Fungal strain to test [Serpula lacrymans, origin: Innsbruck, no. 1SLIBK2018 (Embacher et al., 2021)]
Bacterial strains to test [e.g., Microbacterium spp. (Embacher et al., 2021 and 2022)]
Materials
Autoclaved wood blocks (50 mm × 25 mm × 15 mm) (e.g., Picea abies; all blocks for one experiment should originate from the same batch timber)
Vessels (88 mm height, Ø 75 mm, volume 300 mL, e.g., ROTILABO®, catalog number: EP28.1) filled with 25 mL malt extract agar
Petri dishes 94 mm × 16 mm, without vents (Greiner Bio-One, catalog number: 632180)
Pre-cultured bacterial isolates to be tested
Pre-cultured fungal strain(s) to be tested (e.g., the wood rotting fungus S. lacrymans)
FisherbrandTM Easy ReaderTM conic centrifuge tubes 15 mL, PP (Fisher Scientific, catalog number: 11819650)
Sterile toothpicks
1.5 mL microcentrifuge tubes (Greiner Bio-One, catalog number: 616201)
Reagents
Casein peptone (Roth, catalog number: 8952.2)
Soy peptone (Roth, catalog number: 2365.2)
NaCl (Roth, catalog number: 0601.1)
Agar (Roth, catalog number: 5210.2)
Malt extract (Roth, catalog number: X976.2)
dH2O
D(+)-Glucose monohydrate (Roth, catalog number: 6780.1)
(NH4)2HPO4 (Roth, catalog number: P736.1)
KH2PO4 (Roth, catalog number: 3904.1)
MgSO4·7H2O (Roth, catalog number: T888.1)
CaCl2·2H2O (Roth, catalog number: T885.1)
FeCl3 (Roth, catalog number: 5192.1)
Thiamine-HCl (Roth, catalog number: T911.1)
EDTA disodium salt (Roth, catalog number: 8043.1)
ZnSO4·7H2O (Roth, catalog number: K301.2)
H3BO3 (Roth, catalog number: 5935.1)
MnCl2·4H2O (Roth, catalog number: 0276.1)
CoCl2·6H2O (Roth, catalog number: 7095.1)
CuSO4·5H2O (Roth, catalog number: 8175.6)
(NH4)6Mo7O24·4H2O (Roth, catalog number: 7311.1)
FeSO4·7H2O (Roth, catalog number: P015.1)
KOH (Roth, catalog number: 6751.1)
NaOH (9356.1)
PCR reagents:
Red Taq 2× DNA Polymerase Master Mix (VWR, Radnor, USA)
Primers 27F (5′-AGA GTT TGA TCA TGG CTC A-3′) and 1492R (5′-TAC GGT TAC CTT GTT ACG ACT T-3′) (both 10 μM)
Distilled water
Bovine serum albumin (BSA) 2% [(Roth, catalog number: 3854.2); see as well (Embacher et al., 2021)]
1% agarose gel for gel electrophoresis system [e.g., agarose powder (Sigma, catalog number: A9539)]
Polyethylene glycol 6000 (PEG) 20% (Roth, catalog number: 0158.1)
80% EtOH (-20 °C)
Nuclease-free water
Tryptone soya agar (TSA) plates (see Recipes)
TSA soft agar (freshly prepared on harvesting day, store at ~50 °C until usage) (see Recipes)
Liquid Tryptone soya (TS) medium (see Recipes)
Malt extract agar (MEA) (see Recipes)
Modified Melin-Norkrans (MMN) (see Recipes)
Hutner’s Trace metals (see Recipes)
0.85% NaCl solution (see Recipes)
Sodium borate buffer (20× Stock solution, pH 8) (for agarose gel electrophoresis, see Recipes)
Recipes
Tryptone soya agar (TSA) plates
Casein peptone 1.5% (w/v), 15 g
Soy peptone 0.5% (w/v), 5 g
NaCl 0.5% (w/v), 5 g
Agar 1.8% (w/v), 18 g
dH2O, add up to 1 L
(pH = 7.3 ± 0.2)
Prepare TSA medium, autoclave (120 °C), let it cool down, pour into Petri dishes, and let cool to room temperature (RT). Store at 4 °C.
TSA soft agar
Casein peptone 1.5% (w/v), 15 g
Soy peptone 0.5% (w/v), 5 g
NaCl 0.5% (w/v), 5 g
Agar 0.7% (w/v), 7 g
dH2O, add up to 1 L
(pH = 7.3 ± 0.2)
Prepare TSA medium and autoclave (120 °C). Store at ~50 °C until usage for plate casting after Koch.
Liquid Tryptone soya TS medium
Casein peptone 1.5% (w/v), 15 g
Soy peptone 0.5% (w/v), 5 g
NaCl 0.5% (w/v), 5 g
dH2O, add up to 1 L
(pH = 7.3 ± 0.2)
Prepare TS liquid medium, autoclave (120 °C), and let it cool down. Store at 4 °C.
Malt extract agar (MEA)
Malt extract 3% (w/v), 30 g
Soy peptone 0.3% (w/v), 3 g
Agar 1.8% (w/v), 18 g
dH2O, add up to 1 L
(pH = 3.5 ± 0.2)
Prepare MEA medium, autoclave (120 °C), and cool it down to 50 °C. Pour into Petri dishes and let cool to RT. Store at 4 °C.
Modified Melin-Norkrans (MMN)[after Tauber et al. (2016)]
Glucose, 5 g
(NH4)2HPO4, 0.25 g
KH2PO4, 0.5 g
MgSO4·7H2O, 0.15 g
CaCl2·2H2O, 0.067 g
NaCl, 0.025 g
FeCl3 (1%), 1.2 mL
Thiamine-HCl (10%), 1 μL
Hutner’s trace metals (0.01×), 100 μL
Agar, 20 g
dH2O, add up to 1 L
(pH = 5.6)
Prepare FeCl3, Thiamine-HCl (10%), and Hutner’s trace metals (0.01×) (Hutner et al., 1950) separately. Add them to the media while mixing on the rotary shaker. Measure the pH (and adjust if necessary) and adjust the media to 1,000 mL. Autoclave at 120 °C, let it cool down, pour into Petri dishes, and let cool to RT. Store at 4 °C.
Hutner’s trace metals
EDTA disodium salt, 50 g in 250 mL dH2O
ZnSO4·7H2O, 22 g in 100 mL
H3BO3, 11.4 g in 200 mL
MnCl2·4H2O, 5.06 g in 50 mL
CoCl2·6H2O, 1.61 g in 50 mL
CuSO4·5H2O, 1.57 g in 50 mL
(NH4)6Mo7O24·4H2O, 1.10 g in 50 mL
FeSO4·7H2O, 4.99 g in 50 mL
20% KOH solution (w/v)
For 1 L final mix, dissolve each component in the volume of water indicated. The EDTA should be dissolved in boiling water, and the FeSO4·7H2O should be prepared last to avoid oxidation.
Mix all solutions except EDTA. Bring to boil, then add the EDTA solution. The colour of the mixture turns to green. When everything is dissolved, let cool to 70 °C. While keeping the temperature at 70 °C, add 85 mL of hot KOH (20%). Cool to RT and fill up to 1 L final volume. Close the flask with a cotton plug (allows air exchange) and swirl it once a day while incubating for 1–2 weeks. Usually, the solution will initially be clear green but turns dark red or purple over the next few days, leaving a rust-brown precipitate. If no precipitate forms or the solution remains green, check the pH (should be at approximately 6.7; if there is a big deviation, try adding either KOH or HCl to adjust it).
Filter through two layers of Whatman #1 filter paper and repeat, if necessary, until the solution is clear. Store refrigerated or frozen in convenient aliquots.
0.85% NaCl Solution
Sodium chloride 0.85% (w/v), 8.5 g
dH2O, add up to 1 L
Prepare solution, autoclave (120 °C), and let cool down. Store at RT.
Sodium borate buffer (20× stock solution, pH = 8)
Boric acid, 48 g (concentration 1 M)
NaOH pellets, 8 g
dH2O, add up to 1 L
Prepare 900 mL of dH2O in a suitable container and add the boric acid and the sodium hydroxide to the solution. Stir until all solids are dissolved. Adjust pH to 8 by adding boric acid or NaOH. Fill to 1 L with dH2O.
Dilute the 20× stock solution to 1× before usage for gel electrophoresis (can be used for preparation for agarose gels and in the gel tank).
Equipment
Eppendorf centrifuge 5810 equipped with A-4-81 rotor (Eppendorf SE, Hamburg, Germany)
Heraeus FrescoTM 17 microcentrifuge equipped with 24 × 1.5/2.0 mL rotor with ClickSealTM biocontainment lid (Thermo Scientific, Waltham, Massachusetts, USA)
Counting chamber Thoma (depth: 0.01 mm, 0.0025 mm2, Assistent, Glaswarenfabrik Karl Hecht, Sondheim vor der Rhön, Germany)
Autoclave (HMC Europe, HICLAVE, catalog number: HGS-133)
Incubator (temperature range: 10–30 °C, e.g., Heraeus Vötsch, Vienna, Austria)
Benchtop orbital incubator shaker (New Brunswick Scientific, Edison, model: Innova® 40/40R)
Chemically resistant diaphragm vacuum pump model N810 FT.18 (KNF LABOPORT, Hamburg, Germany)
Vacuum filter manifold and sterile filtration funnels (Millipore SigmaTM, Microfil®, Merck Millipore Billerica, USA)
Nitrocellulose filter (0.45 μm; diam. 47 mm, part no: 1215230, GVS North America, Sanford, USA)
Overhead shaker model REAX2 (Heidolph Instruments, type: 541-21001-00)
Heating oven, 3.8 cu ft, 230 VAC (Memmert, UNB500/230 Basic Oven)
Balance (PM4600 DeltaRange®, Mettler-Toledo, Vienna, Austria)
Microbiological safety cabinet (Platinum SF, Kojair Tech Oy, Mänttä-Vilppula, Finland)
Carton package (to guarantee incubation in darkness)
Sterile glass Petri dishes (100 mm × 20 mm Rotilabo®, Carl Roth, Karlsruhe, Germany)
Sterile plier
Sterilizable punch (that gives e.g., plugs with ~0.8 cm × 0.7 cm × 1.1 cm)
Tweezers
PCR cycler (Primus 96 advanced, Peqlab Biotechnologie, Erlangen, Germany)
Horizontal gel electrophoresis system (RunOneTM Electrophoresis Cell, EmBi Tech, San Diego, USA)
Gel DocTM EZ Imager 30016395-0000 171800 (Bio-Rad Laboratories, Hercules, California, USA)
Thermomixer® comfort 5355 (Eppendorf SE, Hamburg, Germany)
Software
Microsoft Excel version 2209
RStudio version 4.0.3 (2020-10-10)
Procedure
Pre-experiments to determine bacterial viability on wood blocks
Grow bacterial isolates in 5 mL of TS liquid medium overnight (25 °C, 220 rpm, uncontrolled light/dark conditions).
Harvest bacteria and wash them with 1 mL of 0.85% NaCl solution (3×, centrifugation at 5,900× g for 10 min).
Use a Thoma chamber to adjust the bacterial suspension to 108 CFU/mL with 0.85% NaCl solution (end volume minimum 10 mL).
Immerse the autoclaved wood blocks with bacterial suspension and incubate them in a sterile glass Petri dish overnight at 25 °C.
The next day, imprint the wood blocks gently on TSA plates, using a sterilised tweezer. Remove the wood blocks. Incubate at 25 °C overnight to 24 h.
If bacteria grow rapidly on TSA plates, they are useable for the main experiment.
Preparation of the main experiment
Establish fungus on solid agar medium (Figure 1; here, S. lacrymans was pre-cultivated on modified Melin-Norkrans (MMN) for 3–4 weeks at 25 °C).
Figure 1. Established culture of S. lacrymans. Arrow: peripheral zone
Add mycelia-covered agar plugs (use a sterilizable punch, ~0.8 cm × 0.7 cm × 1.1 cm) to vessels filled with 25 mL of MEA medium.
Note: Use the peripheral zone of the pre-culture(s). Set aside some vessels, as they serve as control (Figure 1).
Incubate in darkness for four weeks (25 °C).
In the meantime:
Number the wood blocks (50 ± 0.5 mm × 25 ± 0.5 mm × 15 ± 0.5 mm, e.g., from Picea abies).
Autoclave wood blocks (120 °C) and note their weights (maintain sterile conditions, e.g., put balance under the sterile workbench). Keep them in a sterile environment until use.
Put a minimum of four moisture control specimen wood blocks in the oven and dry them at ~50 °C for 72 h.
Weigh and record the dry mass of the moisture control wood blocks.
Number and weigh glass Petri dishes.
Setup of the main experiment
Start to prepare bacteria approximately 2–3 days before fungal cultures are ready. Culture them on TSA medium (25 °C).
Inoculate 5 mL of liquid TS medium with bacteria (25 °C, 220 rpm, overnight).
Harvest bacteria and wash them with 0.85% NaCl solution (3×, centrifugation at 5,900× g for 10 min)
Use a Thoma chamber to adjust the bacterial suspension to 108 CFU/mL (use 0.85% NaCl solution).
Immerse the numbered, autoclaved, and weighed wood blocks with the bacterial suspension, remove them immediately, drain the excess suspension (sterilised filter paper) and put one (immersed) wood block in each vessel with fungal mycelium. Prepare a minimum of five replicates.
Note: Set aside some vessels as they serve as control.
For each approach, put wood blocks in vessels filled with MEA but without mycelium, as these are additional controls (axenic bacterial controls) (minimum triplicates per bacterium).
Treat a minimum of 14 wood blocks solely with 0.85% NaCl solution and place them on top of the mycelium (axenic fungal control).
Incubate the vessels at 25 °C in darkness (e.g., put them in a carton package) for eight weeks (Figure 2).
Figure 2. Preparation and setup procedure of the wood decay experiment. Illustration of sections B and C.
Harvesting
Remove superficial mycelium from the wood blocks with sterilised tweezers (Figure 2). Collect the mycelium under sterile conditions for further use in step D3.
Imprint the wood blocks on TSA plates and incubate them at 25 °C.
Put each wood in one numbered and pre-weighed glass Petri dish; the numeration should match.
Weigh the wood blocks + Petri dish.
Dry the wood blocks at approximately 50 °C.
Weigh the mycelium collected from the wood.
Dilute the mycelium sample 1:10 with NaCl solution (0.85%).
Shake the mycelium solution for 30 min on a rotary shaker.
Plate 100 μL of the solution on TSA agar plates and incubate at 25 °C.
Use the residual solution.
For plate casting after Koch (if less than 1 mL):
i. Mix residual solution with 3 mL of TSA soft agar.
ii. Vortex gently.
iii. Pour evenly on TSA plates.
For concentration on a nitrocellulose filter with vacuum (if more than 1 mL):
i. Clean the filter unit with 96% EtOH and sterilise the tweezers.
ii. Install the sterile filtration funnel and the filter.
iii. Pour the solution on the filter.
iv. Apply vacuum.
v. Put the filter on TSA medium.
Incubate all agar plates at 25 °C and check daily for growth (Figure 3).
Figure 3. Harvesting and evaluation procedure of the wood-decay experiment. Illustration of sections D and E.
Evaluation
Weigh dried wood blocks.
The wood blocks can stay in the glass Petri dish, as the weight was determined at the start of the experiment.
Weigh them daily until they are dry (should take no longer than 2–3 days).
Check TSA plates for growth, documenting on which plates growth occurs.
Use single bacteria colonies for establishment of pure cultures (TSA agar plate, 25 °C).
Perform colony PCR and check if bacteria match with applied species by sequencing the 16S rRNA gene (Embacher et al., 2021).
i. Prepare PCR reaction: mix 12.5 μL of Red Taq DNA Polymerase Master Mix (2×) with 0.625 mL of each of the primers 27F and 1492R (10 μmol), 10.75 μL of distilled water, and 0.5 μL of BSA (2%) (total volume = 25 μL).
ii. Pick a single bacterial colony not older than three days with a sterile toothpick and add it to the PCR mixture.
iii. Transfer the reactions to the PCR cycler: PCR conditions are 95 °C for 10 min, 30 cycles of 95 °C for 30 s, 53 °C for 30 s, 72 °C for 45 s, and a final elongation step at 72 °C for 10 min.
iv. Check the PCR products by agarose gel electrophoresis (sodium borate buffer 1×, 100 V, approximately 20 min) to confirm their correct size (~1,550 bp).
v. Purify the PCR reaction(s) (PEG protocol by Travis Glenn).
1) Transfer 18 μL of the respective PCR reaction(s) into a 1.5 mL tube and mix with 16 μL of PEG (20%) by constantly pipetting the solution up and down.
2) Put it on the thermo shaker (37 °C, 800 rpm) for 15 min.
3) Centrifuge at 15,000× g for 15 min at RT (21 °C).
4) Discharge the supernatant (as it contains leftover polymerases, primer dimers, and unused dNTPs interfering with the subsequent sequencing reaction).
5) Wash the pellet with 50 μL of ice-cold 80% ethanol (-20 °C).
6) Centrifuge at 15,000× g for 2 min at RT (21 °C).
7) Remove the supernatant.
8) Repeat the centrifugation step and remove the supernatant completely.
9) Incubate the tubes on the thermo shaker (37 °C and 800 rpm) with lid open until no trace (visible drops as well as smell) of ethanol is left.
10) Add 18 μL of nuclease-free water.
11) Pipette the nuclease-free water up and down several times (this ensures that the DNA is fully resuspended in the water).
vi. Check the purified PCR products again with agarose gel electrophoresis (sodium borate buffer 1×, 100 V, approximately 20 min) (Figure 3).
vii. Send for sequencing [using e.g., the Microsynth sequencing service (Balgach, Switzerland)].
Data analysis
Wood decay rates
To ensure that the initial moisture content was uniform throughout all wood blocks and to calculate the estimated dry weight on day zero (dry m0), dry four moisture control specimen wood blocks at ~50 °C for 72 h after autoclaving and weighing (point B5). This can be done in parallel to the assay. Our specimen lost on average 0.6905 g (≙8.62%), hence 91.38% of the mass remained unchanged. Calculate dry m0 as: dry m0 = m d0 × 0.9138, with m d0 being the initial weight of the wood block after autoclaving (before exposure to microbes). Calculate mass loss (ML) difference as: mT (g) = (dry m0 - m1), where mT is the difference of ML after exposure to microorganisms, and m1 and dry m0 are the dry masses after and before degradation, respectively. The ML in % is the measure for the extent of fungal degradation [mT (%) = (dry m0 ÷ m1) × 100]. Prepare a minimum of five replicates for each approach and of three replicates per control approach. Calculate average wood weight loss in % and standard deviation for each approach and illustrate e.g., in a boxplot. Conduct statistical analysis with pairwise.t.test function for pairwise comparison using t-tests with pooled SD (p-value adjustment method: holm and Bonferroni) in the psych package v2.2.5 of the R programming language (RStudio). Additionally, calculate single-factor variance analysis [ANOVA, function aov() in RStudio].
A significantly higher weight loss in the co-cultivation approach than in the axenic fungal control is interpreted as a synergistic effect; lower weight losses are interpreted as antagonism.
Evaluation of PCR 16S rRNA placement
Check sequencing results for quality by evaluating the corresponding chromatograms. If 16S rRNA gene sequences of the applied microbes are available for comparison, compare initial sequences with the sequences of the re-isolated strains by aligning them. Alternatively, blast sequences of the 16S rRNA gene with the NCBI online tool. Limit the searching settings to 16S ribosomal RNA sequences (Bacteria and Archaea) and optimise for highly similar sequences (megablast). According to the results of the Blast search, assign the bacteria to genera and compare with the bacterial strains applied at the beginning. This allows to estimate if the bacteria survived the procedure.
Note: Some mould spores may have survived and developed; we recommend recording affected plates/tins. We excluded these specimens from the calculations.
Notes
Prepare enough tins with pre-cultured fungus, as contaminations occur easily since some steps cannot be performed entirely sterile. Before starting the experiment, we recommend careful planning and listing of what should be prepared.
Mind the controls:
Axenic fungal control: S. lacrymans with wood that is immersed with 0.85% NaCl solution.
Wood sterile control: wood that is immersed with 0.85% NaCl solution, without microbes.
Axenic bacterial control: wood that is immersed with bacterial suspension that was adjusted to 108 CFU/mL without fungus.
Minimum four moisture control specimen wood blocks (without them it is not possible to estimate the initial dry weight dry m0).
It is also possible to sterilise the wood blocks via UV light, as this would preserve the initial state of the wood better than autoclaving. The disadvantage is that more moulds and other contaminants survive this procedure, which might cause problems in the experiment.
Acknowledgments
The protocol was developed on the basis of DIN V ENV12038 (Durability of wood and wood-based products - Wood-based panels - Method of test for determining the resistance against wood-destroying basidiomycetes; German version ENV 12038:2002).
Doctoral program BioApp from the University of Innsbruck is acknowledged for funding. S.N. was funded by the Austrian Science Fund: grant Y0801-B16. We thank Manuela Seehauser and Sara Hnaien for conducting the initial experiments and helping thereby to eradicate first issues.
Competing interests
The authors declare no conflicts of interest or competing interests.
References
Boer, W. d., Folman, L. B., Summerbell, R. C. and Boddy, L. (2005). Living in a fungal world: impact of fungi on soil bacterial niche development. FEMS Microbiol. Rev. 29(4): 795–811.
Bravery, A. (1978). A Miniaturised Wood-block Test for the Rapid Evaluation of Wood Preservative Fungicides, IRG/WP 2113. International Research Group on Wood Protection. Stockholm, Sweden.
Clausen, C. A. (1996). Bacterial associations with decaying wood: a review. Int. Biodeterior. Biodegradation 37(1–2): 101–107.
Cortes-Tolalpa, L., Salles, J. F. and van Elsas, J. D. (2017). Bacterial Synergism in Lignocellulose Biomass Degradation – Complementary Roles of Degraders As Influenced by Complexity of the Carbon Source. Front. Microbiol. 8: e01628.
Embacher, J., Neuhauser, S., Zeilinger, S. and Kirchmair, M. (2021). Microbiota Associated with Different Developmental Stages of the Dry Rot Fungus Serpula lacrymans. J. Fungi 7(5): 354.
Embacher, J., Seehauser, M., Hnaien, S., Zeilinger, S., Kirchmair, M., Neuhauser, S. and Rodriguez-R, L. M. (2022). Cryptic diversity and micro-niche specialization of Microbacterium spp. associated to Serpula lacrymans. Univestity of Innsbruck.
German Institute for Standardisation (2002). DIN V ENV 12038:2002-07 Durability of wood and wood-based products - Wood-based panels - Method of test for determining the resistance against wood-destroying basidiomycetes.
Hegarty, B., Steinfurth, A., Liese, W. and Schmidt, O. (1987). Comparative Investigations on Wood Decay and Cellulolytic and Xylanolytic Activity of Some Basidiomycete Fungi. Holzforschung 41(5): 265–270.
Hutner, S. H., Provasoli, L., Schatz, A. and Haskins, C. P. (1950). Some Approaches to the Study of the Role of Metals in the Metabolism of Microorganisms. Proc. Am. Philos. Soc. 94(2): 152–170.
Kobayashi, D. Y. and Crouch, J. A. (2009). Bacterial/Fungal Interactions: From Pathogens to Mutualistic Endosymbionts. Annu. Rev. Phytopathol. 47(1): 63–82.
Lohwag, K. (1965). Zur Abbauintensität des PilzesPolystictus abietinus (Dicks.) Fr. Holz als Roh- und Werkstoff 23(1): 1–2.
McGuire, K. L. and Treseder, K. K. (2010). Microbial communities and their relevance for ecosystem models: Decomposition as a case study. Soil. Biol. Biochem. 42(4): 529–535.
Sugano, J., Maina, N., Wallenius, J. and Hildén, K. (2021). Enhanced Lignocellulolytic Enzyme Activities on Hardwood and Softwood during Interspecific Interactions of White- and Brown-Rot Fungi. J. Fungi 7(4): 265.
Tauber, J. P., Schroeckh, V., Shelest, E., Brakhage, A. A. and Hoffmeister, D. (2016). Bacteria induce pigment formation in the basidiomycete Serpula lacrymans. Environ. Microbiol. 18(12): 5218–5227.
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Fluorescence Resonance Energy Transfer to Detect Plasma Membrane Perturbations in Giant Plasma Membrane Vesicles
MS Mathew Sebastiao *
NQ Noé Quittot *
IM Isabelle Marcotte
SB Steve Bourgault
(*contributed equally to this work)
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4838 Views: 452
Reviewed by: Alessandro Didonna Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Biochimica et Biophysica Acta (BBA) - Biomembranes Jan 2023
Abstract
Disruptions and perturbations of the cellular plasma membrane by peptides have garnered significant interest in the elucidation of biological phenomena. Typically, these complex processes are studied using liposomes as model membranes—either by encapsulating a fluorescent dye or by other spectroscopic approaches, such as nuclear magnetic resonance. Despite incorporating physiologically relevant lipids, no synthetic model truly recapitulates the full complexity and molecular diversity of the plasma membrane. Here, biologically representative membrane models, giant plasma membrane vesicles (GPMVs), are prepared from eukaryotic cells by inducing a budding event with a chemical stressor. The GPMVs are then isolated, and bilayers are labelled with fluorescent lipophilic tracers and incubated in a microplate with a membrane-active peptide. As the membranes become damaged and/or aggregate, the resulting fluorescence resonance energy transfer (FRET) between the two tracers increases and is measured periodically in a microplate. This approach offers a particularly useful way to detect perturbations when the membrane complexity is an important variable to consider. Additionally, it provides a way to kinetically detect damage to the plasma membrane, which can be correlated with the kinetics of peptide self-assembly or structural rearrangements.
Key features
• Allows testing of various peptide–membrane interaction conditions (peptide:phospholipid ratio, ionic strength, buffer, etc.) at once.
• Uses intact plasma membrane vesicles that can be prepared from a variety of cell lines.
• Can offer comparable throughput as with traditional synthetic lipid models (e.g., dye-encapsulated liposomes).
Graphical overview
Keywords: Plasma membrane Lipid vesicles FRET GPMV Kinetics Membrane perturbation Microplate Cells Amyloid Membrane lytic peptides
Background
The plasma membrane is a highly dynamic, heterogenous barrier between the intracellular components and the extracellular space ( Harayama and Riezman, 2018; Kalappurakkal et al., 2020). The integrity of this barrier is paramount to maintaining normal cell function and, consequently, a great deal of importance is attributed to understanding the processes by which it is disturbed. In particular, numerous amyloidogenic proteins have been found to interact with and destabilise the plasma membrane (Sciacca et al., 2018). While the biophysics of membrane disruption during amyloid formation are typically studied using synthetic models (e.g., liposomes, lipid monolayers, black lipid membranes), these models fall short of accurately depicting the complexity of plasma membranes (Thakur et al., 2011; Serra-Batiste et al., 2016; Sciacca et al., 2020). In vivo, plasma membranes are composed of not only lipids but also a significant amount of proteins (approximately 50% w/w) and polysaccharides (Kusumi et al., 2012; Harayama and Riezman, 2018). This heterogeneity is difficult, if not impossible, to recreate in a synthetic model, and it is highly probable that the molecular composition of the model system can influence protein aggregation and subsequent disruption of the membrane (Zhang et al., 2017). Prior work has demonstrated that varying the proportions of anionic phospholipids, cholesterol, and gangliosides has dramatic effects on the permeability of liposomes (Song et al., 2014; Sciacca et al., 2016 and 2020). Similarly, an accurate prediction of the cytotoxicity based on membrane damage observed in synthetic models alone remains challenging (Cao et al., 2013).
In several recent studies, giant plasma membrane vesicles (GPMVs) were employed to investigate the biophysics of amyloid-associated membrane damage (Quittot et al., 2021; Birol et al., 2018; Schlamadinger and Miranker, 2014). GPMVs are useful models isolated from different cells, such as HeLa, HepG2, Caco-2, RBL, CHO, or INS using comparable and straightforward protocols (Sezgin et al., 2012; Zartner et al., 2021). Consequently, membrane composition of GPMVs is virtually indistinguishable from the parent cells (K.R. Levental and Levental, 2015), although it must be noted that the relative distribution between the inner and outer phospholipid leaflets—in particular those containing anionic phosphatidylserine head groups—can be altered to a non-negligible extent in the vesicle membranes (Sezgin et al., 2012). These recent approaches are limited, however, in their ability to monitor large numbers of samples in a high-throughput fashion. The protocol described here compensates for the current shortcomings of GPMV-based analyses through the use of a microplate assay, measuring the Förster resonance energy transfer (FRET) between two differently labelled vesicles in a suspension. This allows for an experimental throughput comparable to that of traditional liposomes, such that an entire 96-well microplate can be analysed in a single experiment. While this assay is relatively robust and can detect changes to membranes over time, it is limited in its inability to detect small-scale perturbations. Due to the nature of the FRET response, membranes must be in close proximity and under somewhat static conditions. Additionally, smaller defects such as the formation of transmembrane pores may not be detectable with this method. If evaluating transmembrane pore formation with GPMVs is desired, an elegant protocol was developed by Birolet al. (2018), in which cytoplasmic proteins are fluorescently labelled and whose release can be monitored (Birol et al., 2018). Nonetheless, the following approach proves itself to be a valuable source of biophysical data regarding the relative degree of membrane damage during amyloid formation. Additionally, this approach may also be applicable to other membrane-damaging compounds such as antimicrobial and lytic peptides.
Materials and reagents
Biological materials
Rat INS-1E pancreatic cells (provided as a donation by Dr. Marc Prentki at the Centre hospitalier de l’Université de Montréal. Cells can be sourced from Accegen, catalog number: ABC-TC233S)
Reagents
N-ethylmaleimide (NEM) (Sigma-Aldrich, catalog number: E3876)
Human islet amyloid polypeptide (IAPP) (synthesised in-house, can be ordered from Genscript, catalog number: RP11278)
KCNTATCATQRLANFLVHSSNNFGAILSSTNVGSNTY-NH2
Note: Disulfide bridge between C2–C7.
Rat islet amyloid polypeptide (rIAPP) (synthesised in-house, can be ordered from Genscript, catalog number: RP11280)
KCNTATCATQRLANFLVRSSNNLGPVLPPTNVGSNTY-NH2
Note: Disulfide bridge between C2–C7.
Ammonium molybdate (Sigma-Aldrich, catalog number: 277908)
Ascorbic acid (Sigma-Aldrich, catalog number: AX1775)
Phosphorus standard (Sigma-Aldrich, catalog number: P3869)
Sulfuric acid, 98% (H2SO4) (Sigma-Aldrich, catalog number: 258105))
Hydrogen peroxide, 30% (H2O2) (Thermo Fisher, catalog number: 033323-AP)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
Sodium hydroxide (NaOH) (Sigma Aldrich, catalog number: 221465)
Calcium chloride (CaCl2) (Sigma-Aldrich, catalog number: C4901)
HEPES, free acid (Sigma-Aldrich, catalog number: 391314)
HEPES, 1 M solution (Cytiva, catalog number: SH30237.01)
Sodium pyruvate (Cytiva, catalog number: SH30239.01)
Penicillin-Streptomycin, 100× (Cytiva, catalog number: SV30010)
Fetal bovine serum (FBS) (Cytiva, catalog number: SH30070.03)
RPMI 1640, with L-glutamine (Cytiva, catalog number: SH30027.01)
Hank’s buffered saline solution (HBSS), without calcium, magnesium, and phenol red (Cytiva, catalog number: SH30588.01)
Trypsin 1× with 0.05% EDTA (Wisent, catalog number: 325-042-CL)
β-mercaptoethanol (Sigma-Aldrich, catalog number: M3148)
FAST-DiO (Invitrogen, Thermo Fisher, catalog number: D3898)
FAST-DiI (Invitrogen, Thermo Fisher, catalog number: D7756)
Wash buffer (see Recipes)
1 M sodium hydroxide (NaOH) (see Recipes)
Giant plasma membrane vesicle (GPMV) buffer (see Recipes)
3.5% m/v ammonium molybdate (IV) (see Recipes)
10% m/v ascorbic acid (see Recipes)
Complete growth media (see Recipes)
8.9 N H2SO4 (see Recipes)
1 mg/mL FAST-DiO (see Recipes)
1 mg/mL FAST-DiI (see Recipes)
Recipes
Wash buffer
10 mM HEPES, 150 mM NaCl, 2 mM CaCl2, pH 7.4
In a beaker, add 400 mL of ultrapure (type 1) water.
Dissolve HEPES (1.192 g), NaCl (4.38 g), and CaCl2 (0.111 g).
Using a pH meter, slowly add 1 M NaOH under stirring to bring the pH to 7.4.
Transfer the solution to a 500 mL volumetric flask and add ultrapure (type 1) water to complete the volume.
This solution must be filtered through a 0.2 μm PES bottle filter inside of a laminar flood hood before use.
Store the bottle at 4 °C.
1 M NaOH
Weigh out 4 g of NaOH pellets.
In a beaker, add 75 mL of ultrapure (type 1) water.
Slowly add the NaOH pellets to the water while stirring until completely dissolved.
Transfer the solution to a 100 mL volumetric flask and add more ultrapure (type 1) water to complete the volume.
This solution can be stored in a brown glass bottle at room temperature.
CAUTION: The dissolution of NaOH is highly exothermic. This must be done slowly to avoid boiling. For larger volumes, consider keeping the mixing beaker on ice to avoid excessive heat generation.
GPMV buffer
10 mM HEPES, 150 mM NaCl, 2 mM CaCl2, 2 mM NEM, pH 7.4
In a 50 mL conical tube, add 12.5 mg of NEM.
Fill to the 50 mL line with wash buffer.
Once dissolved, filter this solution through a 0.22 μm syringe tip filter inside of a laminar flow hood before use.
Note: This solution must be prepared fresh each time GPMVs are to be produced, as the NEM is not stable in solution for prolonged duration.
3.5% m/v ammonium molybdate (IV)
In a 100 mL volumetric flask, add 3.5 g of ammonium molybdate (IV).
Add 50 mL of ultrapure (type 1) water and ensure that the solid is completely dissolved.
Fill to the 100 mL line with ultrapure (type 1) water.
Transfer to a brown glass bottle and store at 4 °C.
10% m/v ascorbic acid
In a 100 mL volumetric flask, add 10.0 g of ascorbic acid.
Add 50 mL of nanopure water and ensure that the solid is completely dissolved.
Fill the rest to the 100 mL line with nanopure water.
Transfer to a brown glass bottle and store at 4 °C.
Complete growth media
RPMI 1640, 100 U/mL pen-strep, 10% FBS, 10 mM HEPES, 1 mM sodium pyruvate, 50 mM β-mercaptoethanol
Inside of a laminar flow hood, add the following to a 500 mL bottle of RPMI 1640:
5 mL of penicillin-streptomycin (100×).
50 mL of FBS.
5 mL of HEPES (1 M).
5 mL of sodium pyruvate (100 mM).
1.75 μL of β-mercaptoethanol. CAUTION: β-mercaptoethanol is strongly odorous. Ensure adequate ventilation before using this product.
Store this medium at 4 °C.
Note: Working under aseptic conditions is essential here to avoid contaminating the cell culture.
8.9 N H2SO4
Measure 75.8 mL of nanopure water into a glass bottle.
Slowly add 24.2 mL of concentrated (95%–98%) H2SO4. CAUTION: The dilution of H2SO4 is exothermic and should be done with care. Keep the solution on ice and under constant stirring while slowly adding the acid.
1 mg/mL FAST-DiO
Dissolve 1.0 mg of FAST-DiO with 1.0 mL of DMSO.
Prepare aliquots of this master solution by dispensing 50 μL into separate 0.5 mL microfuge tubes.
Note: This compound is light sensitive, particularly when not incorporated into a lipid bilayer. Minimise exposure to ambient light. Using smaller aliquots minimises the number of freeze/thaw cycles for each aliquot.
1 mg/mL FAST-DiI
Dissolve 1.0 mg of FAST-DiO with 1.0 mL of DMSO.
Prepare aliquots of this master solution by dispensing 50 μL into separate 0.5 mL microfuge tubes.
Note: This compound is light sensitive, particularly when not incorporated into a lipid bilayer. Minimise exposure to ambient light. Using smaller aliquots minimises the number of freeze/thaw cycles for each aliquot.
Laboratory supplies
96-well microplates, black, non-binding surface with clear bottom (Corning, catalog number: 3651)
Silicone microplate covers (Corning, catalog number: 3090)
100 mm tissue culture dishes (Sarstedt, catalog number: 3902)
150 mm tissue culture dishes (Sarstedt, catalog number: 3903)
2 mL serological pipettes (Sarstedt, catalog number: 1252025)
5 mL serological pipettes (Sarstedt, catalog number: 1253025)
10 mL serological pipettes (Sarstedt, catalog number: 1254025)
25 mL serological pipettes (Sarstedt, catalog number: 1685020)
15 mL conical centrifuge tubes (Sarstedt, catalog number: 62.554.100)
50 mL conical centrifuge tubes (Sarstedt, catalog number: 62.547.004)
0.5 mL microfuge tubes (Sarstedt, catalog number: 72.704)
100 kDa centrifugal filter units (Amicon, Millipore, catalog number: UFC910024)
0.2 μm PES bottle filter units (ThermoFisher, catalog number: 569-0020)
10 mL syringes (BD, catalog number: 309604)
0.22 μm syringe filters (Sarstedt, catalog number: 83.1826.001)
16 mm × 150 mm glass reaction tubes (VWR, catalog number: 47729-580)
12 mm × 75 mm glass reaction tubes (VWR, catalog number: 47729-570)
Plastic plugs for 12 mm × 75 mm reaction tubes (VWR, catalog number: 60819-003)
Brown glass bottles (Sigma-Aldrich, catalog number: DWK218062454)
Microplate lids with silicone seal (Corning, catalog number: 07-200-699)
18 G needles (BD precision glide, catalog number: 305195)
Equipment
Plate reader (Molecular Devices, Spectramax i3)
Centrifuge (Thermo, IEC CL30) with swinging bucket rotor (S41*)
Laminar flow hood (VWR Microzone)
Incubator (Thermo, Heracell 150i)
Hot plate (Fisherbrand, Isotemp)
Aluminium heating block (Thermo Fisher, catalog number: 88880136)
pH meter (Fisherbrand, Accumet AE150)
Micropipettes (Gilson Pipetteman P200, catalog number: F144058M)
Vortex mixer (Scientific industries vortex genie-2, catalog number: SI-0236)
Metal bowl or glass beaker, large enough to contain a test tube rack and capable of being heated at 100 °C. Can be replaced for a water bath if one is available
Software and datasets
Prism 8 (GraphPad)
Excel, Office 16 (Microsoft)
Procedure
Cell culture
Retrieve one aliquot of 106 INS-1E cells from liquid nitrogen storage and thaw them slowly at room temperature. Once the cells have warmed enough to be safely handled, the tube can be further warmed by rolling it back and forth between the hands. CAUTION: Liquid nitrogen is cold enough to cause skin damage in cases of prolonged contact. Wear appropriate PPE when handling liquid nitrogen.
Once thawed, work under a laminar flow hood and maintain aseptic conditions. Transfer the cells to a 15 mL conical tube and add 10 mL of growth medium, pre-warmed to 37 °C. Centrifuge the cells at 500× g for 5 min at room temperature and remove the supernatant.
Resuspend the 106 cells in 10 mL of 37 °C complete growth medium and transfer to a 100 mm tissue culture dish. Store the culture dish inside of the incubator at 37 °C with 5% CO2.
After two days, passage the cells:
Remove the old growth medium and rinse with 2 × 5 mL of 37 °C HBSS.
After removing the HBSS, add 2 mL of 37 °C trypsin and incubate the cells for 5 min at 37 °C.
Retrieve the cells and inactivate the trypsin with 5 mL of 37 °C growth medium.
Collect the cells in a 15 mL conical tube and centrifuge at 500× g for 5 min at room temperature.
Discard the supernatant and resuspend the cell pellet in 10 mL of 37 °C growth media.
Transfer the cells to a new tissue culture dish and return them to the incubator.
Check on the cells daily until 80% confluency is reached and passage again.
After three passages, evenly split the cells (15 × 106 cells) into two 100 mm culture dishes.
After two days, check the confluency. If the cells are below 80% confluency, wait another day.
Once the cells have reached ≥ 80% confluency (≥ 15 × 106 cells), transfer the cells from one 100 mm culture dish into a 150 mm culture dish. Split the second culture dish into two 100 mm culture dishes (7.5 × 106 cells each).
One dish serves as a reserve for continuing the culture growth, and the other serves as a source of cells to produce GPMVs. Maintain these as normal in the incubator, changing the medium and splitting the cells as necessary.
Allow the cells in the 150 mm culture dish to reach ≥ 80% confluency (≥ 33 × 106 cells) and split them evenly into two 150 mm culture dishes. These two 150 mm culture dishes will be used to produce GPMVs in the next steps. Maintain these two 150 mm culture dishes in the incubator at 37 °C with 5% CO2.
This process is graphically summarised in Figure 1.
Note: Working under aseptic conditions is essential here to avoid contaminating the cell culture.
Figure 1. Schematic representation of cell culture and splitting to prepare cells for giant plasma membrane vesicles (GPMVs) production
GPMV formation
Retrieve the two 150 mm culture dishes from the incubator once they reach 80% confluency (33 × 106 cells). Discard the old growth medium and wash the cells twice with 10 mL of 37 °C HBSS.
Discard the HBSS and wash the cells twice with 10 mL of 37 °C wash buffer.
Discard the wash buffer and add 5 mL of 37 °C GPMV buffer to the cells.
Note: The GPMV buffer must be prepared fresh before each use, as the NEM is unstable in solution for prolonged periods of time. NEM was chosen as the GPMV-inducing agent due to its potent ability to induce GPMV formation and its low impact on peptide aggregation. Unlike other GPMV-inducing agents, NEM does not crosslink proteins but rather alkylates free cysteines (Scott, 1976; Erdinc Sezgin, 2022; I. Levental et al., 2010; K. R. Levental and Levental, 2015).
Incubate the cells in the GPMV buffer for 2 h at 37 °C with 5% CO2.
Retrieve the two 150 mm culture dishes from the incubator and carefully collect the liquid above the cells containing the GPMVs in one 15 mL conical tube per culture dish.
Centrifuge the GPMVs at 500× g for 5 min at room temperature to pellet any cell debris. Transfer the supernatant containing the GPMVs to two empty 15 mL conical tubes.
GPMVs labelling
To one 15 mL conical tube with the GPMVs, add 5 μL of the FAST-DiO solution (1 mg/mL) for a final concentration of 1 μg/mL and gently rock the tube back and forth for 10 s to mix it. To the second tube, add 2 μL of FAST-DiI solution (1 mg/mL) for a final concentration of 0.4 μg/mL and rock the tube back and forth gently for 10 s to mix.
Note: FAST-DiO and FAST-DiI stock solutions are light sensitive. To avoid degradation of the fluorophores, work in a dimly lit environment.
Incubate the GPMVs for 1 h at 37 °C to complete the labelling.
Recover the two tubes from the incubator and transfer the contents of each tube to a 100 kD Amicon centrifugal filter unit to concentrate the GPMVs and to reduce the amount of any unincorporated tracers.
Centrifuge the Amicon units at 400× g for 1 min and verify the volume remaining in the filter unit, ensuring that the entire volume does not pass through. Repeat these short centrifugations until 500 μL remain in each unit.
This process is graphically summarised in Figure 2.
Figure 2. Workflow for the isolation and labelling of giant plasma membrane vesicles (GPMVs) formed from cultured cells
Carefully recover the GPMVs retained in the filter unit using a P200 pipette, transferring the labelled membranes into 1.5 mL microfuge tubes. The GPMVs prepared this way should be used within 48 h and can be stored at 4 °C if needed.
Note: While FAST-DiO and FAST-DiI are not particularly sensitive to photodegradation after being incorporated into the membranes, it is good practice to nonetheless avoid unnecessary exposure to ambient light.
Total phosphorus assay
In order to ensure experimental reproducibility and to accurately quantify the ratio between GPMVs and peptide, the concentration of phospholipids in the GPMV suspension can be measured through the total phosphorus assay.
Preheat an aluminium block to 210 °C on a hot plate.
Prepare a calibration curve using the phosphate standard reagent using 12 mm × 75 mm glass tubes. See Table 1 for the quantities suggested.
Table 1. Phosphorus quantities of PO43- standard
Quantity of P (μmol) Volume of PO43- standard (μL)
0.00000 0
0.00650 10
0.01625 25
0.03250 50
0.04875 75
0.06500 100
Transfer 100 μL of each GPMV suspension to 12 mm × 75 mm glass tubes.
Add 450 μL of 8.9 N H2SO4 to all tubes.
For each standard and sample to be analysed, puncture a plastic stopper five times with an 18 G needle to prevent the buildup of pressure during the next steps.
Heat the tubes on the aluminium block for 25 min, ensuring that the temperature remains between 200 and 220 °C. This reaction serves to digest the organic matter in the samples.
Remove the tubes and cool for 5 min until they can be handled. Add 150 μL of H2O2 to each tube, cap, and heat at the same temperature for 30 min. This reaction serves to remove any dark coloration that may develop in the reaction tubes due to the digestion of organic material.
Remove the tubes from heat and allow them to cool at room temperature. Meanwhile, prepare a 100 °C water bath by bringing a metal bowl of water to a boil on the hotplate.
Note: It is important that the bowl is large enough to place all the reaction tubes inside at the same time. This can be accomplished using a plastic test tube rack.
Transfer the residual liquid in each tube to a clean 16 mm × 150 mm glass tube using a P1000 micropipette.
Add 500 μL of 10% ascorbic acid, 500 μL of 3.5% ammonium molybdate (IV), and 3.9 mL of nanopure water.
Gently vortex the samples, cover each tube with a glass marble, and heat in the water bath for 7 min. Here, the reduction of the mixture by ascorbic acid permits the colorimetric reaction between phosphorus and ammonium molybdate.
Allow the tubes to cool for 5 min before transferring 200 μL of each into a 96-well microplate. Measure the absorbance in each well at 820 nm and calculate the concentration of phosphate using a linear regression. See the section on data analysis for details. CAUTION: Hot H2SO4 and H2O2 are caustic and should be handled with care. Use gloves, lab coat, safety glasses, and work inside of a chemical fume hood.
FRET kinetics
Using the phospholipid concentrations obtained from D, prepare dilutions of the GPMVs at 2× concentrations (100, 25, and 6.25 μM) in the GPMV buffer.
Transfer 50 μL of each mixed GPMV suspension to a 96-well microplate (black, non-binding surface with clear bottom) in triplicate. IMPORTANT: It is crucial to prepare the appropriate blanks to be able to normalise the data afterwards.
To each GPMV sample, add 50 μL of either GPMV buffer (blanks), IAPP freshly solubilised at 25 μM in GPMV buffer, or rat IAPP (control peptide) freshly solubilised at 25 μM in GPMV buffer.
Cover the microplate with the silicone lid to prevent evaporation.
In the plate reader, measure the fluorescence due to FRET every 10 min (λexcitation = 485 nm, λemission = 570 nm). Program the kinetics to perform a single agitation for 5 s and then record the fluorescence intensity from the bottom of the plate at 10 min intervals for 24 h with gain at the medium setting.
Note: While this protocol has been developed to detect plasma membrane damage associated with amyloidogenic peptides, it is likely to work with other, non-amyloidogenic peptides as well. Additionally, the aggregation of amyloidogenic peptides is likely sensitive to the number of measurements made during the experiment. It is therefore strongly recommended being consistent in the number of wells measured and the frequency at which the data points are collected.
Data analysis
Linear regression
Using the optical densities measured during the total phosphorus assay, perform a linear regression by plotting the optical density against the quantity of Pi (μmol) for the six standards. Sample data is shown below (Table 2, Figure 3). Using these data points, fit a linear function and display the slope, intercept, and correlation value for the regression.
Table 2. Standard curve obtained after total phosphorus assay
Pi standard (μL) Pi (μmol) OD at 820nm
0 0.00000 0.0525
10 0.00650 0.0768
25 0.01625 0.1047
50 0.03250 0.1477
75 0.04875 0.1919
100 0.06500 0.2558
Figure 3. Linear regression from total phosphorus assay
From the linear regression in Figure 3, the Pi content of the GPMV samples can be calculated using the equations below.
OD = slope × Qty Pi + intercept
Qty Pi = (OD - intercept)/slope
ConcGPMVs (μM) = (Pi μmol)/(100 μL) × 106
The values obtained from these calculations are shown below in Table 3, with the final Pi concentrations highlighted in green. These values correspond directly to the phospholipid concentration.
Table 3. Sample values obtained from the linear regression
Sample Volume used (μL) OD 820nm Pi (μmol) Pi (μmol/μL) Pi (μM)
GPMVs 100 0.0958 0.01403 1.4036 × 10-4 140.4
GPMV FDiO 75 0.0764 0.00758 1.0102 × 10-4 101.0
GPMV FDiI 75 0.0822 0.00951 1.2677 × 10-4 126.8
FRET kinetics
Average the fluorescence values of each sample across the corresponding triplicate wells.
Normalise the averaged data for each measurement as described by the equation below, where Fblank is the fluorescence intensity measured in the GPMVs in buffer, and Ftreated is the fluorescence of the GPMVs treated with human or rat IAPP.
Fnorm = Ftreated/Fblank
Fit the data with a Boltzman sigmoidal curve using GraphPad Prism by selecting Fit a curve with a non-linear regression in the analysis area and finding the Boltzman sigmoidal function in the section Classic equations from prior versions of prism, as shown in Figure 4. This can also be done in Excel or any other plotting software using the following equation:
f(x) = Fmin + (Fmax - Fmin)/(1 + eV(50-x)/k)
Figure 4. Representative fluorescence resonance energy transfer (FRET) kinetics obtained from FAST-DiO- and FAST-DiI-labelled giant plasma membrane vesicles (GPMVs)
From the fitting, kinetic parameters, such as the top and bottom fluorescence values, the rate k, and the time to half max V50, can be extracted and compared as shown below in Figure 5. To compare the values between the two peptides, either a Student’s t-test or ANOVA can be used, depending on the number of conditions analysed.
Figure 5. Kinetic parameters extracted from the Boltzman sigmoidal fitting to the data in Figure 4
Validation of protocol
Results from this protocol can be viewed in Figure 10 of the published article Sebastiao et al. (2023). Data were obtained in triplicate from independent GPMV preparations. The non-amyloidogenic rat IAPP was used as a control variable in all tests, and FRET values were normalised against the corresponding membranes in the absence of any peptide. Statistical analysis was done using a two-way ANOVA (GraphPad Prism 8) on the values extracted from the sigmoidal fitting.
General notes and troubleshooting
General notes
The rat INS-1E cells should not be kept for longer than 20 passages to ensure the reliability and quality of the GPMVs produced.
This process is applicable to other cell lines as well. It has been done using Chinese hamster ovarian cells (CHO-K1) without any modifications (Quittot et al., 2021). For other cell lines, optimization of GPMV production may be required.
It is important to maintain all parameters as consistent as possible due to the sensitive nature of amyloid self-assembly. Even varying the number or frequency of measurements made during the kinetic assay can alter the aggregation behaviour of amyloidogenic peptides (Sebastiao et al., 2017).
Troubleshooting (Table 4)
Table 4. Troubleshooting
Issue Solution
No phospholipids were detected in the total P assay Vesicles are likely diluted. Either use a larger volume than 100 μL or further concentrate the vesicles using the Amicon.
Insufficient FRET signal is detected Increase the quantity of FAST-DiO from 5 to 10 μL.
Kinetics are noisy The process is inherently sensitive to many experimental parameters. Ensure that all peptide solutions are properly monomerised and completely dissolved before adding to the GPMVs.
Inability to fit a sigmoidal function The sigmoidal response is typical for amyloid aggregation. It is not necessarily the response for associated membrane disruption. If required, select a different model to fit the data.
Acknowledgments
This work was supported by the National Science and Engineering Research Counsel (NSERC) and the Quebec Research Funds for Nature and Technologies (FRQNT). The protocol was adapted from previous work (Sebastiao et al., 2023).
Competing interests
The authors declare no competing interests.
References
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A Novel Imaging Protocol for Investigating Arabidopsis thaliana Siliques and Seeds Using X-rays
R Brylie A. Ritchie
TU Theodore A. Uyeno
DD Daniel F. Rincon Diaz
Ansul Lokdarshi
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4839 Views: 887
Reviewed by: Samik BhattacharyaSwati MeghaDawid S Zyla
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Abstract
Understanding silique and seed morphology is essential to developmental biology. Arabidopsis thaliana is one of the best-studied plant models for understanding the genetic determinants of seed count and size. However, the small size of its seeds, and their encasement in a pod known as silique, makes investigating their numbers and morphology both time consuming and tedious. Researchers often report bulk seed weights as an indicator of average seed size, but this overlooks individual seed details. Removal of the seeds and subsequent image analysis is possible, but automated counts are often impossible due to seed pigmentation and shadowing. Traditional ways of analyzing seed count and size, without their removal from the silique, involve lengthy histological processing (24–48 h) and the use of toxic organic solvents. We developed a method that is non-invasive, requires minimal sample processing, and obtains data in a short period of time (1–2 h). This method uses a custom X-ray imaging system to visualize Arabidopsis siliques at different stages of their growth. We show that this process can be successfully used to analyze the overall topology of Arabidopsis siliques and seed size and count. This new method can be easily adapted for other plant models.
Key features
• No requirement for organic solvents for imaging siliques.
• Easy image capture and rapid turnaround time for obtaining data.
• Protocol may be easily adapted for other plant models.
Graphical overview
Arabidopsis siliques using the Inspex 20i X-ray machine
Keywords: X-rays Plant development and morphogenesis Arabidopsis thaliana siliques and seeds
Background
Seed development is critical for the survival of all flowering plants (Jardinaud and Petitprez, 2003). Of the current plant models, Arabidopsis thaliana, with its convenient seed pod (silique), continues to be the most widely used tool for studying fruit/seed development and morphogenesis (Bates et al., 2013; Kao and Nodine, 2021). Arabidopsis siliques contain rigid cell walls and are pigmented by chlorophyll, which makes analyses of seed morphology, anatomy, and ultrastructure challenging (Hedhly et al., 2018; Attuluri et al., 2022;). To overcome these difficulties, conventional methods employ different types of fixatives (e.g., Acryloyl-X, FPA50) and optical clearing reagents (e.g., chloral hydrate, 2,2’-thiodiethanol, sodium dodecyl sulfate) for whole-mount microscopy (Hedhly et al., 2018; Attuluri et al., 2022). Despite their pervasive use, these methods are tedious, as these clearing procedures have specific protocols that involve the use of harmful organic solvents and lengthy (> 24 h) durations (Attuluri et al., 2022). Hence, there is a growing demand for non-invasive techniques that can provide high-quality digital data with shorter turnaround times and avoid the use of toxic organic solvents. In the work presented here, we describe a new, non-invasive technique for visualizing Arabidopsis siliques with seeds at different stages of their growth using a custom X-ray imaging system. While X-ray microscopy has been used for imaging flower development in Arabidopsis (Prunet and Duncan, 2020), studies of seed and silique development continue to rely mostly on conventional histological methods. We show that with minimal sample processing, without any use of toxic organic solvents, high-quality images can be captured with our custom X-ray imaging system (in 1–2 h) for downstream processing and data analyses. Statistical tests using one-way ANOVA and frequency distribution show high reproducibility and consistency of seed count and seed diameter in dried Arabidopsis siliques across three independent experiments. Taken together, our easy-to-use X-ray system serves as a new imaging tool for visualizing Arabidopsis siliques and seeds and, with some modification, can be adapted for other plant models.
Materials and reagents
Biological materials
Arabidopsis thaliana ecotype Columbia (Col_0) plant at midflowering, complete flowering, and seed harvest stage (Boyes et al., 2001).
Laboratory supplies
Fine precision medium tipped tweezers/forceps (Fisher Scientific, catalog number: 12-000-157)
Petri dish (Fisher Scientific, catalog number: FB0875712)
Permanent ink markers
Paper towels (Fisher Scientific, catalog number: 19-040-898)
Tape (Fisher Scientific, catalog number: 15949)
X-ray source: Kevex PXS10-16W MicroFocus system
Boss LS-1416 CO2 laser cutter
Equipment
Inspex 20i X-ray machine
For custom manufacturing of the X-ray imaging cabinet, details are provided in Figure 1. The Inspex 20i X-ray machine was assembled by Kodex Inc. (Nutley, NJ).
Figure 1. Schematics of the X-ray imaging setup. (A) Cartoon sketch of the Inspex 20i X-ray machine with all the accessories. The custom steel cabinet (J) measures 64.77 cm (depth) × 127 cm (height) × 78.74 cm (width) and is lined with a lead sheet to conform to BRH 21 CFR 1020.40 and FDA 21 CFR 1020.40, in accordance with Georgia State X-ray radiation regulations. The X-ray source (G) is a Kevex PXS10-16W MicroFocus system that produces a downward facing cone-beam so that shadows can be detected on the Kodex LTX-1717 digital detection panel (E) located at the bottom of the cabinet. This amorphous silicon flat panel detector has a cesium iodide scintillator in the outer layer that allows for the conversion of X-rays to light, which is then digitized by the photodiode layer below for transmission to the computer for display on the screen. The microfocus system allows focused images to be enlarged or reduced by varying the specimen height between the source and detector (closer to the source results in a magnified image). Placing the specimen as close as possible to the source (G(a)) results in a field of view of approximately 9.5 mm2. The field of view adjacent to the detector (E) is approximately 45 times greater at 431.8 mm2 (the detector is 17 × 17 inches). Components external to the cabinet (red): (A) Power supply for the X-ray source; (B) Power supply and data transfer unit for the detector; (C) Computer to render and save images; (D) Uninterruptable power supply and surge suppressor for the X-ray machine. Components inside the cabinet (black): (E) The Kodex LTX-1717 digital detection panel, E(w) width 44.45 cm, E(d) depth 44.45 cm, E(h) distance between detector and source 104.14 cm; (F) The cone beam path of the X-ray (cone of illumination, 53°); (G) The Kevex PXS10 MicroFocus X-ray source (maximum settings of 80 kV, 0.100 mA); (H) Image collection shutter trigger; (I) Touchscreen monitor and keyboard for image viewing. (Average settings for this study: Silique specimens in this study were placed on a low-density plastic platform suspended 98.4 cm above the detector, and the source was operated at 40 kV and 0.09 Amps). (B) Picture of the X-ray machine with the front door open.
Scale for the measurement of siliques/seeds using the Inspex 20i X-ray machine
We used an X-ray semi-transparent scale that was cut from cardstock (weight 90 Lb) using a Boss LS-1416 CO2 laser cutter (70 W). We did this because the printed hashmarks on solid plastic or metal rulers are not visible in X-ray images. We designed the scale using the Adobe Illustrator vector drawing package. Only the millimeter hashmarks and the external contour of the scale were cut; the millimeter number markings were not. The kerf width of the laser cutter (aka. hashmark line width) measured an average of 0.152 mm. This was wider than we had hoped, but the centers of the marks were extremely accurate. The dimensional accuracy of the laser cutter was listed as being ± 0.0127 mm, and this high accuracy was confirmed using multiple micrometer measurements from the hashmark centers. Since the scale bars and their hashmarks could be seen beside the seeds and siliques, they proved a simple, economical, and reproducible method of calibrating X-ray image dimensions.
Software and datasets
VetView Console: All-In-One Workstation Software is used for image acquisition, processing, and editing.
DaVinci Resolve 11: Calibration software, run as a script, to dark calibrate data read from the X-ray detector while the source is not providing illumination.
FIJI ImageJ Build 2.13.1: This image processing program was used to count and measure diameters of the seeds within the silique.
GraphPad Prism Version 9.4.1: Statistical analyses and production of graphs.
Procedure
Please see Video 1 for sections A, B, and C detailing the procedure.
Video 1. Video tutorial showing the sample preparation and X-ray imaging procedure
Setting up low-density plastic platform in the Inspex 20i X-ray machine for sample tray (see Figures 1, 2 for details)
Figure 2. Sample loading and readiness of the Inspex 20i X-ray machine. (A) Large Petri dish with silique samples on damp paper towel. (B) Small Petri dish with two silique samples and scale placement using metal tweezer. (C, D) Lateral (C) and Overhead (D) view of the low-density platform held by tape at both ends. Also shown is the Petri dish carrying silique samples and scale shown in panel B. (E, F) Power supply for the X-ray source showing X-ray key enabled in panel F.
Open the front door of the X-ray machine by pulling the metal latch located at the center-right of cabinet J.
Tape the low-density plastic platform ~3.5 cm underneath the source G. We found that the highest contrast images were made when specimens were imaged on a platform made of low-density polyethylene plastic films. This is because these films are radiotransparent relative to the siliques, seeds, and paper calibration scales. However, many other plastics such as the polystyrenes and polycarbonates used in plastic Petri dishes can also be used with good results.
Close the front door by pushing the metal latch handle until the click sound confirming the interlock system has engaged is heard.
Calibration of Inspex 20i X-ray machine (see Figure 1 and Figure 2E–2F for details)
Press the power switch located on control panel A for switching the control panel A ON.
Turn ON the control panel B using the power switch.
Turn ON panel C to start the desktop by pressing the power button ON.
On the control panel A, turn the metal key counterclockwise to position ON and press the green button on panel A.
Note: The following lights should be visible now: Red color for X-ray ON, yellow for interlock, and green flashing indicating the main power ON. See Figure 2 for details.
Wait for the X-ray tube unit to warm up for ~10–15 min.
Note: As part of the warm-up sequence, the X-ray sign above the steel cabinet J will illuminate and the numbers on the control panel A (Figure 2F) will show a gradual increase in voltage and current (Amps).
Full system readiness is marked by illumination of the green light next to the standby on panel A, the readings for volts and amps set at 0, and the X-ray sign OFF (Figure 2F).
Open the front door (see step B1) and ensure that the overhead lights inside cabinet J are not illuminated. If the light is off, then close the front door and proceed to the next step.
Double click on the dark calibration tab located on the desktop to run dark calibration.
Notes:
This step is required for best image contrast.
As part of this step, a pop-up window will appear on the desktop screen and then disappear, indicating step completion.
Repeat step B8.
Harvesting Arabidopsis siliques and imaging procedure with Inspex 20i X-ray machine (see Figures 1 and 2 for details)
We used standard methods for growing Arabidopsis plants on soil. This step is a standard procedure in plant biology and various resources are commonly available.
Dampen one paper towel with deionized water and place it into a large Petri dish.
Note: The entire paper towel should be wet, then squeeze out the extra deionized water.
For harvesting siliques of the desired growth stage (Boyes et al., 2001), use tweezers to gently pluck one silique at a time and place it into a clear Petri dish.
Note: Gentle care is highly recommended for handling dried siliques as they tend to split and release the seeds.
Retrieve siliques, with pedicle attached, using fine-tipped tweezers and place on the damp paper towel until ready for next step.
Note: Siliques can stay for a maximum of 1–2 min only on the damp paper towel, as longer wait times may trigger imbibition in dry siliques.
Transfer one silique into the base of a small clean Petri dish.
Note: A maximum of 5–6 siliques can be placed side-by-side to improve throughput.
Place the 1 cm scale bar next to the siliques in the small Petri dish.
Open the front door of cabinet J and place the small Petri dish onto the low-density plastic platform.
Note: A few trial runs may be required to adjust sample positioning, focus, and overall image quality.
Close the front door and open the VetView console program on the desktop.
Click “new sample” tab and label the sample using the sample ID and sample name boxes.
Click capture and wait for a pop-up window.
Note: Ensure that the new window is set to the whole sample and then click capture.
Click the green ON button on control panel A.
On the control panel A, set volts to 40.0 and amps to 0.09 using the dial knobs below the number screen.
Click the handheld X-ray button H.
As the VetView program displays “image acquiring”, click the red button on control panel A to turn the X-ray beam OFF.
Note: This step ensures an extended life for the X-ray bulb. Leave X-ray beam on for no longer than one minute after image is acquired.
Image will appear on the desktop screen and will show a white square placed around the center of the image.
Note: The white square is a tool to assist with centering the image and cropping unnecessary blank space.
Ensure your sample and scale bar are within the marquee by left clicking on the marquee edge and dragging it to center your samples inside the marquee frame.
Note: The shape of the marquee can be adjusted by dragging either the corners or the sides.
Select “accept image” for downstream processing.
Note: For reacquiring an image, select “reject image”. Repeat step D5 by adjusting the small Petri dish with the silique samples. Click edit protocol on the desktop screen and start from step D9.
After accepting the desired image, the VetView console program will reconfigure to the home page.
Select the name provided for the sample (step D7) and click open study.
Note: This will allow you to right-click to adjust the contrast.
Click the “save” tab to save the image file and close study to return to the home page of the VetView console program.
Note: Check saved images in the destination folder before proceeding to another round of imaging.
Remove the small Petri dish with silique samples from the X-ray machine by opening the front door of cabinet J.
Repeat steps D3–D18 for acquiring additional images.
When the experiment is finished, turn off the overhead lights in cabinet J and power down the X-ray machine by switching off the desktop (panel C) and control panels A and B.
Note: Powering OFF sequence is in the opposite order as discussed in section C for powering ON.
Data analysis
Results
Traditional histological methods for visualizing Arabidopsis siliques employ organic solvents and typically require > 24 h before image capture. Our new silique imaging tool boasts minimal sample processing and faster experimental turnaround time, through the use of X-ray imaging. The X-ray images in Figure 3 and the measurements shown in Figure 4 show the suitability and reproducibility of using X-ray imaging for visualizing Arabidopsis siliques at different stages of development. We originally imaged only single siliques per X-ray exposure; however, the field of view of our setup allows for at least four siliques and a scale bar to be imaged per X-ray exposure at the highest usable image resolution (Figure 3). This feature allows for higher throughput and reducing variability between experiments.
Figure 3. Image of siliques with scale bar. Representative image of siliques at maturity stage (dry) with the scale at the bottom. White color scale bar (1 cm) is shown in the center of the image.
The results of our three independent experiments indicate that the seed count and diameter are consistently reproducible between experiments (Figure 4). In summary, the custom X-ray imaging system offers significant advantages over traditional silique and seed visualization methods relying on clearing/fixative reagents. It is envisioned that researchers may adapt this new tool for imaging silique and seed development of different plant species.
Figure 4. Silique and seed imaging using Inspex 20i X-ray machine and statistical tests for reproducibility. Representative image of silique at (A) young, (B) mature, and (C) dry stages of maturity captured by the Inspex 20i X-ray machine. Inset in each panel shows an example image of the silique at different stages of maturity. (D) Total seed count (y-axis) and (E) seed diameter (y-axis) in millimeters (mm) of dry siliques from three independent experiments (x-axis - Test 1, 2, 3). Center bold horizontal line within each test represents the median (panel D: 26–28.5; panel E: 0.39–0.41), while the range of distribution is shown with horizontal small edges at both the ends of the data points. Each test is represented by n > 20 data points, and each dot represents an average of n > 30 siliques per test. One-way ANOVA P value > 0.05 for panels D and E.
Validation of protocol
This protocol is validated by providing data to support the claims. Please refer to Figure 4, panel D & E, for details on the number of replicates and statistical tests.
Notes
In our experiments, we did not find any phenotypic changes in the Arabidopsis plants cultivated from seeds that were exposed to X-rays.
Acknowledgments
We thank the Valdosta State University, College of Science & Math and the Department of Biology for supporting undergraduate and faculty research.
Author contributions: Brylie A Ritchie: Performed the experiments, collected data and wrote the original draft of methods; Theodore A Uyeno: Designed the X-ray imaging module and scale, prepared figures, wrote the original draft for figures for reviewing and editing; Daniel F Rincon Diaz: Performed the experiments, collected data, wrote the original draft of methods and assisted in making the video tutorial; Ansul Lokdarshi: Conceptualized the project, developed the figures, analyzed data, wrote, reviewed, and edited the final manuscript.
Competing interests
Authors declare no competing interests.
References
Attuluri, V. P. S., Sánchez López, J. F., Maier, L., Paruch, K. and Robert, H. S. (2022). Comparing the efficiency of six clearing methods in developing seeds of Arabidopsis thaliana. Plant Reprod. 35(4): 279–293.
Bates, P. D., Jewell, J. B. and Browse, J. (2013). Rapid separation of developing Arabidopsis seeds from siliques for RNA or metabolite analysis. Plant Methods 9(1): 9.
Boyes, D. C., Zayed, A. M., Ascenzi, R., McCaskill, A. J., Hoffman, N. E., Davis, K. R. and Gorlach, J. (2001). Growth Stage-Based Phenotypic Analysis of Arabidopsis: A Model for High Throughput Functional Genomics in Plants. Plant Cell 13(7): 1499–1510.
Hedhly, A., Vogler, H., Eichenberger, C. and Grossniklaus, U. (2018). Whole-mount Clearing and Staining of Arabidopsis Flower Organs and Siliques. J. Vis. Exp.: e3791/56441.
Jardinaud, M. and Petitprez, M. (2003). SEED DEVELOPMENT | Embryogenesis. In B. Thomas (Ed.). Encyclopedia of Applied Plant Sciences (pp. 1225–1232). Oxford: Elsevier.
Kao, P. and Nodine, M. D. (2021). Application of expansion microscopy on developing Arabidopsis seeds. In Guichard, P. and Hamel, V. (Eds.). Methods in Cell Biology (pp. 181–195). Academic Press.
Prunet, N. and Duncan, K. (2020). Imaging flowers: a guide to current microscopy and tomography techniques to study flower development. J. Exp. Bot. 71(10): 2898–2909.
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4,840 | https://bio-protocol.org/en/bpdetail?id=4840&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Human Schwann Cells in vitro III. Analytical Methods and a Practical Approach for Quality Control
PM Paula V. Monje
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4840 Views: 600
Reviewed by: Vivien J. Coulson-ThomasDhruv Rajanikant PatelWeiyan Jia
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Original Research Article:
The authors used this protocol in Molecular Neurobiology Aug 2018
Abstract
This paper introduces simple analytical methods and bioassays to promptly assess the identity and function of in vitro cultured human Schwann cells (hSCs). A systematic approach is proposed to unequivocally discriminate hSCs from other glial cells, non-glial cells, and non-human SCs (authentication), identify hSCs at different stages of differentiation, and determine whether individual hSCs are proliferative or senescent. Examples of how to use distinct cell-based approaches for quality control and routine troubleshooting are provided to confirm the constitution (identity, purity, and heterogeneity) and potency (bioactivity) of hSC cultures from multiple sources. The bioassays are valuable for rapidly gauging the responses of hSCs to mitogenic and differentiating factors and ascertaining the cells’ basic properties before performing co-culture or cell grafting studies. The assays are image based and use adherent hSCs established in monoculture to simplify the experimental setup and interpretation of results. Finally, all sections contain thorough background information, notes, and references to facilitate decision making, data interpretation, and ad hoc method development for diverse applications.
Keywords: Bioassays Proliferation Differentiation Senescence Identification Potency Authentication Immunodetection Markers Myelin Fibroblasts
Background
Human Schwann cell (hSC) cultures are valuable in vitro models for interrogating the biology of SCs during nerve development, maturation, and regeneration in normal and disease states. These cultures are also valuable as transplantable biologics to treat the injured or dysmyelinated central (CNS) and peripheral (PNS) nervous systems [reviewed in Guest et al. (2013); Monje et al. (2021c); Vallejo et al. (2022)]. Quality control testing of the cultured hSCs is needed for numerous reasons. First, hSC cultures are seldom pure and contain other proliferative cells from the endoneurium and the connective tissue layers. Second, the cultured hSCs differ from those in the original tissue as mature hSCs rapidly dedifferentiate after isolation and adapt to the in vitro environment. Third, non-cellular impurities and factors introduced during culture, such as animal serum and coating reagents, may alter hSC function. Altogether, these issues emphasize the importance of implementing robust analytical methods to promptly (1) confirm the hSC phenotype and the level of purity of the cell cultures; (2) evaluate the state of differentiation and biological activity of the hSCs; and (3) recognize and quantify myelin debris and other visible and subvisible impurities. Selecting suitable methodologies to address these elements will depend on the intended application. Cultured cells are manipulated biologics regardless of the manufacturing method used and need to be scrutinized thoroughly. Contrary to research-grade hSC cultures, those used clinically must meet the highest possible quality standards for identity and function (Bunge et al., 2017; Khan et al., 2021).
The goal of this paper is to describe easy-to-run analytical methods to reveal the purity, bioactivity, and often-changing constitution of donor-relevant hSC cultures by proposing the following: (1) to effectively identity hSCs using live and fixed cultures and discriminate them from cellular and non-cellular contaminants such as myelin, fibroblasts, non-human cells, and glial cells of CNS origin; and (2) to determine hSC function by means of proliferation, differentiation, and senescence assays (Figure 1). As explained in Protocol 1, immunological methods are recommended to reveal the phenotype and level of purity of the hSC cultures. Once the composition of the cultures is confirmed and the cells are authenticated for the species and tissue of origin (Protocol 2), investigators are encouraged to further interrogate their products using specific bioassays designed to evaluate the cellular responses to agonists known to exert an action on SC proliferation and differentiation (Protocol 3). Each protocol contains background information and notes to guide technical or logistical decisions regarding the analysis and interpretation of results. The troubleshooting section defines issues that researchers may encounter, along with appropriate approaches for problem resolution. Assessing cell viability, metabolic activity, microbial contamination, and cell transformation is important, but assays are not described here because they can be found elsewhere. Regarding contamination with cancerous cells, it should be noted that transformation of the hSCs in culture, or amplification of tumor cells imported from the tissue of origin, are unlikely events in hSC cultures from normal tissues (Bastidas et al., 2017).
The experimental conditions described in the following sections were optimized for adult nerve-derived hSC monocultures (adherent cells) obtained by traditional methods, irrespective of the source, passage, and stage of differentiation of the cells. However, our approach may be equally suitable for the analysis of other types of hSC cultures if enough cells (1 million or more) are available for the initial seeding. The use of multi-well plates is recommended for direct image analysis by fluorescence or light microscopy to reduce time, labor, and cost during experimentation and facilitate data interpretation. However, it is advisable to implement a systematic investigative approach (explained above) including more than one measure (or assay) to confirm the results. The suggested strategy is not intended to replace longer term in vitro (e.g., traditional co-culture systems) or in vivo studies (e.g., cell grafting). Some complex cellular responses requiring the interactions of hSCs with other cell types and the physical environment cannot be recapitulated under the simplified conditions of these experiments. Notwithstanding, the assays described here can be used as screening or discovery platforms to tackle various questions, since no consensus has been reached on how to approach quality control measures for SCs in culture. Future directions include scaling up the assays for a larger number of samples and developing quantitative approaches for specific readouts (e.g., myelin gene expression) relevant to nerve regeneration, therapy, or disease.
Figure 1. Quality control assessments for cultured human Schwann cells (hSCs). Measures of identity and bioactivity described in this study were selected based on their cell type specificity and reliability for scrutinizing hSC cultures in basic and translational research applications. Image: mitogen- and serum-starved hSCs stained with CellTrackerTM Green, a vital fluorescent dye.
Materials and reagents
Our cell culture protocols use research-grade materials, reagents, and solutions that are endotoxin-free and suitable for cell culture. Still, researchers are encouraged to assess the quality of relevant materials by appropriate methods to identify potentially toxic or incompatible elements in the culture medium, coating reagents, and plasticware.
All assays can be implemented using basic cell culture equipment and labware typically available in research labs. The description of products provided below is for reference only. Researchers may find comparable items from other providers. An exception is the primary antibodies (Table 1) because they have been validated carefully in our lab using cultured hSCs and nerve tissues. If other antibodies are selected, take into consideration that the ones used for rodent SC research may fail to recognize the respective antigens from humans. For this reason, it is important to confirm the specificity and reactivity of all antibodies in-house using proper positive and negative controls. Some of the suggested monoclonal antibodies were produced in our laboratory from hybridoma cell cultures but alternative ones are available from commercial sources (see Table 1). Lastly, the list below is not fully comprehensive. Additional technical details can be found in the accompanying papers (Aparicio and Monje, 2023; Monje, 2023).
Table 1. Useful antibodies to characterize human Schwann cells (hSCs) and non-glial cells established in cell culture. These antibodies were validated using traditional adult nerve-derived hSC monocultures. Staining with NGFR, Sox10, and S100B antibodies, alone or together with fibronectin, SMA (α-smooth muscle actin), and FAP (fibroblast activation protein/seprase) antibodies, is recommended for the initial characterization of hSCs and non-glial cells. We have not found a ubiquitous non-glial cell marker. Protocol 1C is suitable for staining with all antibodies except for anti-O1 and anti-O4, which can be accomplished only in live cells (Protocol 1B). A 1:200–1:500 starting dilution is suitable for most antibodies; the optimal concentration should be determined by the end user in the target cells. (*) Indicates that the antibodies are produced from hybridoma cell lines. MBP: myelin basic protein; MAG: myelin-associated glycoprotein; MPZ: myelin protein zero; GFAP: glial fibrillary acidic protein.
Name of marker & subcellular localization Product information Expected results
NGFR
• Cell membrane
• Mouse monoclonal 8737-IgG. ATCC # HB8737*
• Rabbit monoclonal, EP1039Y. Abcam, catalog number: ab52987.
• Highly specific hSC marker. Expressed at high, homogeneous levels in all cultured hSCs.
Sox10
• Nuclear
• Rabbit monoclonal. Abcam catalog number: ab155279. • Highly specific hSC marker. Expressed at high, homogeneous levels in all hSCs.
S100β
• Cytoplasmic (typical) and nuclear (rare)
• Rabbit polyclonal. DAKO, catalog number: Z0311. • Mouse monoclonal. Sigma, catalog number: S2657 • Highly specific hSC marker. High, homogeneous levels in all hSCs with occasional nuclear localization.
Sox2
• Nuclear
• Rabbit polyclonal. Santa Cruz, catalog number: sc-20088. • Highly specific hSC marker, heterogenously expressed in individual hSCs and reduced by CPT-cAMP stimulation.
Nestin
• Cytoplasmic
• Mouse monoclonal. EMD Millipore, catalog number: MAB353. • Highly specific hSC marker. Heterogeneously expressed (donor- or batch-dependent) and reduced by CPT-cAMP stimulation.
GFAP
• Cytoplasmic
• Rabbit polyclonal. DAKO, catalog number: Z0334. • Specific hSC marker expressed at low levels in expanded hSCs.
MAG
• Cell membrane and myelin
• Mouse monoclonal. Chemicon, catalog number: MAB1567. • Mostly in myelin debris. Expressed at low or undetectable levels in expanded hSCs. Induced with CPT-cAMP.
MBP
• Myelin
• Rat monoclonal. MAB386 (Millipore, former Chemicon) • Mostly in myelin debris. Non-myelin-associated MBP is rarely detectable in expanded hSC cultures.
MPZ/P0
• Cell membrane and myelin
• Chicken polyclonal. AB9352 (Millipore, former Chemicon) • Mostly in myelin debris. Non-myelin-associated MPZ is rarely detectable in expanded hSC cultures.
Erg2/Krox20
• Nuclear
• Rabbit polyclonal (non-commercial)
• Expressed at low levels in hSCs. Enhanced with CPT-cAMP.
O4
• Cell membrane and myelin
• Mouse monoclonal. O4-IgM * • Expressed in all hSCs right after isolation but only in a proportion of the expanded hSCs. Enhanced with CPT-cAMP.
O1
• Cell membrane and myelin
• Mouse monoclonal. O1-IgM* • Expressed in hSCs right after isolation. Expanded hSCs are O1- regardless of cAMP levels.
Vimentin
• Cytoplasmic
• Rabbit monoclonal. Cell Signaling, catalog number: D21H3, 5741. • Equally expressed in hSCs and fibroblasts at high, homogeneous levels.
CD44
• Cell membrane
• Mouse monoclonal. Cell Signaling, catalog number: 156-3C11. • Equally expressed in hSCs and fibroblasts at high, homogeneous levels.
Fibronectin
• Extracellular (typical) and cytoplasmic
• Mouse monoclonal. Santa Cruz, catalog number: sc-8422.
• Mouse monoclonal. Sigma, catalog number: HFN 36.3 (89062006)*
• Filamentous or punctuated staining with significantly higher levels in fibroblasts as compared to hSCs.
SMA
• Cytoplasmic
• Mouse monoclonal. Thermo Fisher: catalog number: MS113-PO.
• Rabbit monoclonal. Cell Signaling, catalog number: D4K9N 19245.
• Expressed at high levels in a proportion of non-glial cells, possibly pericytes. hSCs display low levels of SMA.
FAP
• Cell membrane
• Rabbit Monoclonal. Cell Signaling, catalog number: 66562. • Expressed at high levels in a proportion of non-glial cells. hSCs do not typically express FAP.
Thy1/CD90
• Cell membrane
• Rabbit monoclonal. Abcam. Catalog number: 92574 Abcam • Expressed at variable levels in a proportion of non-glial cells. Hard to detect by simple immunostaining. hSCs do not typically express Thy1.
Supplies and consumables
Polypropylene conical-bottom centrifuge tubes, 15 and 50 mL (Corning, catalog numbers: 430791 and 430290)
Serological pipettes, 5, 10, and 25 mL, polystyrene, sterile (VWR)
Pasteur pipettes, polystyrene, individually wrapped for liquid disposal (VWR, Argos Technology, catalog number: 10122-560)
Laminin-coated cell culture dishes for cell expansion. 100 mm × 20 mm plates, polystyrene (Corning, catalog number: 353003) coated with a laminin substrate, as described in Andersen and Monje (2018)
Laminin-coated multi-well plates for analytical assays. Cell culture–treated 24-well plates, flat bottom, polystyrene (Corning, catalog number: 3524) coated sequentially with PLL and laminin, as described in (Andersen and Monje, 2018). (Optional) Use commercially available 24-well plates coated with poly-L-ornitine (PO) and laminin (BD Biosciences, catalog number 354659). Do not plate hSCs on uncoated surfaces. Polystyrene plates or chamber slides are preferred. Coverslips are not suitable since the hSCs are unstable and display an abnormal morphology on any glass surface
Paraformaldehyde (PFA) 20% stock solution (Electron Microscopy Sciences, catalog number: 15713)
Methanol (Sigma, catalog number: 154903) maintained at -20 °C for cell permeabilization. (Optional) 0.1% (v/v) Triton X-100 (Sigma, catalog number: 11332481001) prepared in D-PBS and stored at 4 °C
Laboratory wrapping film (Parafilm, catalog number: PM-996) and aluminum foil
Media, supplements, and other cell culture products
Distilled water, cell culture grade (Fisher Scientific, Gibco, catalog number: 15-230-147)
Dulbecco’s phosphate-buffered saline (DPBS) with calcium and magnesium, pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 14190)
Hank’s balanced salt solution (HBSS) formulated without calcium or magnesium and containing phenol red, pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 14170-112)
Dulbecco’s modified Eagle’s medium (DMEM) with high glucose and phenol red, pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 11965092)
DMEM, Nutrient Mixture F-12 (DMEM/F-12), no glutamine, with phenol red (Thermo Fisher Scientific, Gibco, catalog number: 21331020)
De-complemented fetal bovine serum (FBS) (HyClone, catalog number: SV 30014.03), stored in aliquots at -80 °C
100× GlutaMAX supplement (Thermo Fisher Scientific, Gibco, catalog number: 35050061)
Gentamycin 50 mg/mL, 1,000× stock solution (Thermo Fisher Scientific, Gibco, catalog number: 15750-060)
HEPES buffer solution 1 M (Thermo Fisher Scientific, Gibco, catalog number: 15630-080)
Normal goat serum (GeneTex, catalog number: GTX73206), stored in aliquots at -80 °C
Forskolin (Sigma-Aldrich, catalog number: F68861); for a detailed protocol on preparation, storage, and use of forskolin stock solution, see Andersen and Monje (2018)
Heregulin-β1 (referred to as heregulin), HRG1-B1177-244 recombinant peptide (Preprotech, catalog number: G-100-03); for a detailed protocol on preparation, storage, and use of heregulin stock solution, see Andersen and Monje (2018)
CPT-cAMP stock solution (5 mM in DMEM), prepared from adenosine 3′,5′-cyclic monophosphate, 8-(4-chlorophenylthio), sodium salt (Calbiochem, catalog number: 116812); for a detailed protocol on preparation, storage, and use of CPT-cAMP stock solution, see Monje (2018)
Low proliferation medium (LP) (see Recipes)
High proliferation medium (HP) (see Recipes)
Starvation or D1 medium (DMEM/F12 - 1% FBS) (see Recipes)
PFA-based fixation solution (see Recipes)
Blocking solution (see Recipes)
Antibodies, dyes, and commercially available detection kits
Mouse monoclonal antibodies from hybridoma cell lines. Anti-nerve growth factor receptor (NGFR), anti-O4, and anti-O1 (Sommer and Schachner, 1981) in the form of conditioned medium produced from HB-8737 cells (also known as 200-3-G6-4, obtained from the American Type Culture Collection, ATCC), O4 cells, and O1 cells (kindly provided by Dr. Melitta Schachner), respectively. Researchers can refer to our publication (Ravelo et al., 2018) for technical details on our hybridoma culture protocols. Briefly, transfer the cell content of a hybridoma stock (1 × 106–2 × 106 cells/cryovial) directly into a T-75 flask containing Iscove’s modified Dulbecco’s medium (with phenol red) supplemented with 10% FBS and antibiotics. Culture the cells in suspension inside a CO2 incubator until the cultures are sufficiently dense and the medium becomes slightly acidic. Next, separate the culture supernatant from the cellular content by centrifugation to obtain conditioned medium enriched in monoclonal antibodies. The conditioned medium is often used without dilution, but the specificity and reactivity of each batch should be tested using appropriate positive control cells or tissues
Primary antibodies. pERK1/2/MAPK mouse monoclonal antibody (Santa Cruz, catalog number: sc7383); pAkt-Ser-473 rabbit polyclonal antibody (Santa Cruz, catalog number: sc7985); Akt rabbit polyclonal (Cell Signaling, catalog number: 9272); ERK2/MAPK rabbit polyclonal (Santa Cruz, catalog number: sc154); human nuclei (HNA), mouse monoclonal (Sigma, MAB1281, clone 235-1) or HNA mouse monoclonal (Abcam, ab191181). Other antibodies are listed in Table 1
Fluorescent secondary antibodies of the appropriate species and class, Alexa FluorTM-conjugated (Molecular Probes). Fluorochromes should be chosen and combined as optimal for visualization
FM4-64FX, fixable membrane stain (Invitrogen, catalog number: F34653). (Optional) FluoroMyelinTM (red or green) myelin stain (Invitrogen, catalog number: F34652)
Hoeschst-34580 (Molecular probes, catalog number: H21486). (Optional) 4′,6-Diamidino-2-Phenylindole, Dilactate, (DAPI, Invitrogen, catalog number: D3571)
Click-iT EdU (5-ethynyl-2′-deoxyuridine) Alexa-Fluor-594 Imaging Kit (Life Technologies, catalog number: C10339)
Senescence-associated (SA) β-Galactosidase (SA-β-Gal) staining kit (Cell Signaling Tech, catalog number: 9860)
VectaShieldTM antifade liquid mounting reagent for fluorescence (Vector Laboratories, catalog number: H-1000). (Optional) Prepare a homemade mounting reagent, as suggested in Ravelo et al. (2018), to control photobleaching and preserve the stained cultures. Add a sufficient volume of mounting reagent to fully cover the fixed cells (300 μL/well in a 24-well plate) or a couple of drops if a glass coverslip is mounted on top
Equipment
Biological safety cabinet, BL2 level (Thermo Scientific 1300 series class II, 1300 series, type A2)
Cell incubator set at 37 °C and 8%–9% CO2 (Thermo Scientific Forma, series II, water-jacketed)
Inverted phase contrast microscope (VWR) equipped with 10× to 40× phase contrast objectives and attached digital camera (VWR, V5MP)
Benchtop refrigerated centrifuge (Beckman CoulterTM, Allegra X-12R) equipped with swing bucket rotor (SX4750) and adapters for 50 and 15 mL tubes
Automated cell counter for the image-based counting of cells in suspension (Bio-Rad, TC20) or a hemocytometer for manual cell counting
Inverted fluorescence microscope (Olympus, IX71) for brightfield, phase contrast, and fluorescence microscopy; equipped with standard UV, FITC, and TRITC filter sets and attached digital camera
Procedure
Protocol 1: Analysis of identity and purity
The characterization of cultured SCs has changed over time. Early investigations relied on histochemical and ultrastructural visualization of cells (Askanas et al., 1980) and biochemical measurements for the enzymatic activity of 2′,3′-cyclic nucleotide 3′-phosphohydrolase (Reddy et al., 1982). Currently, investigators can implement various methods to unequivocally understand the constitution of cell cultures. High-throughput sequencing methodologies, such as RNA-seq performed on whole populations or single cells, are thus far the most sensitive and comprehensive approaches. However, simple image-based immunological tests using cell type–specific antibodies are fast, informative, and cost effective to analyze cultured cells. These tests can aid in making crucial decisions, such as changing the culture conditions, expanding the cells further, or subjecting them to purification.
The hSC phenotype from normal mature peripheral nerves can be identified based on the expression of markers such as NGFR (also known as the Low-Affinity Neurotrophin Receptor p75), the S100 Calcium Binding Protein, Beta (S100B), and the SRY-Box Transcription Factor 10 (Sox10) (Scarpini et al., 1986; Assouline and Pantazis, 1989a and 1989b). It is important to estimate the type and proportion of non-glial cells, because endoneurial fibroblasts, endothelial cells, and pericytes can contaminate the hSC cultures in various proportions (Hoyng et al., 2015; Weiss et al., 2016; Peng et al., 2020; Khan et al., 2021). hSCs in vitro derived from non-pathological human nerves or the skin encompass defined phenotypic characteristics (Stratton et al., 2017; Chu et al., 2022); however, the cultured hSCs can change if passaged extensively or subjected to certain experimental treatments. A rigorous phenotyping analysis cannot be overlooked when the source of hSCs is unknown, the tissue material used to derive the cells is abnormal, or the hSCs are obtained artificially by in vitro differentiation or other methods [discussed in Monje (2020)].
The sections below include basic protocols for the routine analysis of hSC cultures. Procedure A describes a generic protocol to plate hSCs in multi-well dishes for direct image analysis. Procedure B encompasses a quick protocol (i.e., < 1 h long) to identify hSCs based on live-cell immunostaining. Procedure C uses fixed cells to detect hSC markers alone or together with markers of non-glial cells (see Table 1). Procedure D enables the detection of extracellular and intracellular myelin debris, which may be undesirable in certain applications. Representative results from these procedures are shown in Figures 2–3 and 5–7.
Finally, it should be mentioned that the methods for quantification of fluorescent cells and the parameters for assay design, such as the selection of positive and negative controls, are not described here. Please refer to publications from our group for additional experimental data and information (Monje et al., 2018; Monje, 2020; Peng et al., 2020).
Cell plating (common to all protocols)
Prepare a single-cell suspension of hSCs, count the cells, and estimate their viability by the method of choice. For a reference to detailed protocols, see Ravelo et al. (2018).
Plate the cell suspensions at mid-density in HP medium (see Recipe 2) inside a 24-well plate using 50,000 cells/well or an equivalent density (25,000 cells/cm2) as determined by the available surface. Use plastic dishes coated with PLL (or PO) and laminin to ensure prompt adhesion and survival of the cells (see Materials and reagents). Consider that enough replicates are needed to achieve statistical significance in all measurements that compare control vs. treatment conditions. Triplicate samples are sufficient for most assays.
Culture the cells in the CO2 incubator for 24–72 h to obtain a culture at 40%–60% confluence, as determined empirically by periodic image analysis by phase contrast microscopy.
When the cells are ready to be stained (or fixed), remove the culture medium by gentle aspiration inside a biosafety cabinet, working quickly to avoid unnecessary exposure of cells to the air flow.
(Optional) Wash the cells with D-PBS or LP medium (see Recipe 1) if floating myelin or cellular debris is evident by phase contrast microscopy.
Proceed with the description provided in Protocol 1B–1D.
Live cell immunolabeling
Add a sufficient volume of conditioned medium containing NGFR or O4 antibodies (e.g., 500 μL for each well of a 24-well plate). Alternatively, use an appropriate dilution of a commercially available antibody (see Table 1). Properly scale up or down the suggested measures if other plate formats are used in this and all subsequent steps.
Incubate the cells at room temperature (RT) for 20 min and leave them undisturbed inside the biosafety cabinet (see Note a).
Remove the conditioned medium and quickly wash the cells 2× with an excess of D-PBS to eliminate unbound antibodies while ensuring that cells do not detach.
Fix the cells for 20 min with fixation solution (see Recipe 4) and wash them 3× with D-PBS to remove traces of PFA.
Prepare a 1:300–1:1,000 dilution of Alexa-conjugated secondary antibody (as determined by the primary antibody) in blocking solution (see Recipe 5) and add it to the cells in an adequate volume (see Protocol 1B, step 1). For instance, use Alexa Fluor 488-conjugated anti-mouse IgG to detect NGFR (from HB-8737 conditioned medium) or anti-mouse IgM to detect O4 (see Note b).
Add a general nuclear stain, such as DAPI or Hoechst-33342 (1:1,000) to the secondary antibody solution. Alternatively, the nuclear staining can be done independently before mounting after a brief 10 min incubation at RT using dyes (1:1,000) diluted in D-PBS.
Cover the plates with aluminum foil and place them on an orbital shaker for 30–60 min at slow motion (20–30 rpm), as the cells can peel off from the dishes even after fixation.
Wash the cells 3× with D-PBS to eliminate unbound secondary antibodies.
Add mounting reagent in a sufficient volume to cover the cells (see Materials and Reagents). Mount the cells with a glass coverslip if visualization is performed at 40× or higher power objectives.
Image the cultures using an inverted fluorescence microscope and a suitable filter set (see Equipment) starting at 10× to obtain a panoramic view of the cultures (Figure 2A–2B) before visualization at higher magnification (Figure 2C–2G).
Figure 2. Identification of human Schwann cell (hSC) cultures using distinct antibody combinations. Low (A–B) and high (C–G) magnification images of typical hSC cultures are shown to reveal the reactivity and specificity of some recommended antibody combinations. Cultures containing a proportion of non-glial cells (white arrowheads) were selected for display. The morphology of hSCs is varied according to the plating density, the formulation of the culture medium, and the level of fibroblast contamination, but the antibody reactivity and specificity are unchanged. All cultures were grown in HP medium except for those in panel G, which were grown in differentiation medium for seven days (Protocol 3C). Refer to Table 1 for technical details and interpretation of results. Nuclei were stained with DAPI (blue) in all images.
Estimate the proportion of NGFR+ and O4+ cells in relationship to total cells (DAPI+ or Hoechst+) to inform on the percentage of glia/Schwann (NGFR+) and non-glial cells (NGFR-), and of mature (O4+) and immature (O4-) hSCs. This can be achieved by manual or automated counting methods.
Immunolabeling of fixed cells
Gently aspirate the culture medium, fix the cells for 20 min with fixation solution (see Recipe 4), and wash them 3× with D-PBS.
Permeabilize the cells by incubation with cold methanol (preferred) or 0.1% Triton X-100 in D-PBS for 10 min. Permeabilization of the cell membranes is required only for the staining of intracellular antigens exhibiting cytoplasmic or nuclear localization. Methanol is preferred because detergents can damage the cellular structure if timing is not carefully controlled.
Aspirate the methanol (or the Triton X-100 solution) quickly and wash the cells 3× with D-PBS.
Add blocking solution (see Recipe 5) for at least 30 min at RT.
Remove the blocking solution and replace it with either of the following: (1) conditioned medium from NGFR hybridoma cells (see above); (2) anti-S100B (1:300 in blocking solution), or (3) anti-Sox10 (1:300 in blocking solution) to identify hSC-specific markers with localization to the plasma membrane, the cytoplasm, and the nucleus, respectively. (Optional) Refer to Table 1 for an alternative selection of primary antibodies.
Incubate the cells overnight at 4 °C with gentle agitation (20–30 rpm in an orbital shaker) to maximize immunolabeling detection.
Proceed as described in Protocol 1B, steps 5–11, using a corresponding selection of secondary antibodies and a nuclear stain to estimate the proportion of hSCs (NGFR+, S100B+, Sox10+) and non-glial cells (NGFR-, S100B-, Sox10-) by imaging analysis.
Labeling of myelin debris
Perform a live staining of the cell cultures using anti-O1 antibodies (see Materials and Reagents) following the protocol described for anti-O4 (Protocol 1B, steps 1–3) (see Note c).
Fix the cells with fixation solution (see Recipe 4), as described in Protocol 1B, step 4.
Prepare a labeling solution containing AlexaFluor488-conjugated anti-mouse IgM antibodies (1:500) in combination with FM4-64FX (1:1,000) or FluoroMyelinTM-red (1:500) in an adequate volume of D-PBS (see Notes c and d). (Optional) Incorporate a nuclear stain such as DAPI or Hoechst-33342 to this mixture.
Add the abovementioned labeling solution directly to the cells, cover the plates with aluminum foil, and incubate them for 30–60 min with gentle agitation (20–30 rpm in an orbital shaker). Monitor the cells under the fluorescence microscope to confirm the levels of staining, as larger diameter myelin granules require a longer labeling time with FM4-64FX or FluoroMyelinTM.
Remove the solution and wash the cells 3× with D-PBS to remove traces of the fluorophores.
Perform image analysis to discriminate and quantify extracellular (O1+) and total (FM4-64FX+/FluoroMyelinTM+) myelin granules. The observation of granular spots heavily stained with O1 antibodies may be interpreted as extracellular myelin. In such cases, confirm that O1+ granules are non-cellular by showing they are not associated with intact nuclei. Total myelin can also be revealed by immunostaining with antibodies against myelin proteins (see Table 1) in fixed cell cultures (see Protocol 1C).
Notes:
Prolonged incubation with primary antibodies (> 30 min) leads to internalization or capping of the antibodies, which is seen as patchy, granular (instead of smooth) staining. Placing the cells on ice reduces the capping effect; however, lowering the temperature should be done with caution because the temperature shock can impair hSC adhesion. More information on troubleshooting and staining protocols can be found in Ravelo et al. (2018).
NGFR immunostaining can be performed using live or fixed cells (Protocol 1C) with similar results. However, a live-cell labeling protocol is required for lipid antigens, as PFA fixation leads to non-specific membrane binding of O4 and O1 (galactocerebroside/GALC) antibodies. Whereas NGFR expression is homogeneous and constant in all hSCs regardless of the culture conditions, O4 expression is heterogeneous and depends on cAMP stimulation (Peng et al., 2020). For best results, culture hSCs in CPT-cAMP-containing medium for at least three days (Protocol 3) before performing O4 immunostaining. The expression of myelin lipids declines rapidly after hSC isolation from the nerves. It has been reported that hSCs from explant cultures can maintain GALC expression in 30% of the cells two weeks after isolation (Turnbull et al., 2001). Yet, GALC+ and O1+ cells are rarely found in established hSC cultures even under optimal conditions for differentiation (Monje et al., 2018).
hSC cultures from mature nerves contain a variable proportion of myelin debris in the extracellular environment and within the cells themselves, because hSCs effectively engulf myelin fragments before and after isolation from the nerve tissue. Extracellular myelin can be washed out easily; however, intracellular myelin is retained inside the hSC’s cytoplasm. Whereas extracellular myelin stains heavily with O1 and O4 antibodies, intracellular myelin requires staining with myelin fluorophores. An important point to take into consideration is that cultured, isolated hSCs do not contain intact or newly formed myelin sheaths. If myelin-like structures are observed in primary or established hSC cultures, they are likely derived from the original tissue. Myelin debris do not interfere with hSC function in vitro but can inhibit axon growth and trigger an immune response if myelin-containing cultures are used for cell grafting in vivo.
FM4-64FX and FluoroMyelinTM enable quick and selective myelin labeling in live and fixed hSC cultures and can be used in combination with antibody staining. FM4-64FX is preferred because it emits strong fluorescence and can be fixed permanently with aldehyde-based buffers. These dyes are believed to work via lipophilic interactions. However, membranes other than myelin and intracellular lipid droplets can be stained as well.
Protocol 2: Authentication of hSC cultures
Authentication is performed to correctly identify the cell material by a series of established methods. An authentication guide has been established for most common mammalian cell lines. However, a similar guide for cultured hSCs has not yet been developed. Traditionally, SC cultures were derived directly from a nerve fascicle (or ganglia), which simplified the identification of the neural (Schwann) phenotypes by virtue of knowing the tissue of origin. At present, however, culturing technologies have expanded, and hSC-like cells can be created by various in vitro techniques (Huang et al., 2020). This shift in culturing technologies represents a major challenge in identifying the hSCs and discriminating them from other glial and non-glial cell types.
State-of-the-art technologies, such as RNAseq, can readily determine the constitution of cell cultures. However, the time and cost associated with these high-resolution approaches can be limiting. Immunodetection methods arehighly reliable for quickly confirming (or disproving) the species and tissue of origin of whole cell cultures or individual cells within mixed populations. For instance, antibodies with selective reactivity to human NGFR and MPZ proteins have been used to differentiate human from non-human SCs in cell culture and within grafted tissues (Levi and Bunge, 1994). Alternatively, this discernment has been achieved by concomitantly detecting a SC-specific marker (e.g., with MPZ or NGFR antibodies) and a ubiquitous human marker with nuclear localization (Bastidas et al., 2017). Researchers have discriminated hSCs from central glial cells (namely, astrocytes) based on NGFR immunodetection, as this membrane receptor is expressed only in PNS glia (Assouline and Pantazis, 1989a). Individual myelin-associated hSCs, i.e., either mature (myelin-forming) or repair (myelin-engulfing), can be discriminated from CNS glia (oligodendrocytes) by immunodetection of the major peripheral myelin glycoprotein MPZ (Levi and Bunge, 1994).
Here, we suggest a simple approach to validate hSC cultures by incorporating at least two elements of support, (1) for the human origin and (2) the PNS origin of the hSCs, using combined antibody- and ligand-based fluorescent labeling methods (Table 2). This approach may be useful in the following scenarios: (1) when the cells deemed to be SCs are not obtained from human peripheral nerves or ganglia; (2) when the origin of the tissue or the cell cultures is dubious or unknown; (3) when contamination with spinal cord or CNS tissue cannot be ruled out; and (4) when there is a known or suspected pathology affecting the donor tissue used to derive the hSCs. Authentication is also recommended when cells initially confirmed to be normal hSCs depart from the expected phenotype while in culture. This observation may signal an underlying issue, such as fibroblast overgrowth, transformation of the hSCs, or cross-contamination with cancerous cell lines. Importantly, results from our suggested bioassays (Protocol 3) should be interpreted under the assumption that the cells under investigation faithfully represent the PNS-derived hSC phenotype. This author recommends a stepwise approach to cell identification (Protocol 1) and authentication (Protocol 2) when researchers believe that this general assumption should be challenged.
Table 2. Quick staining protocols for human Schwann cell (hSC) validation in culture. Table 2 presents simple strategies for verifying both the human and the SC background of cultured cells. Different staining methods using human-selective and multispecies-selective NGFR antibodies are proposed. NGFR-8737 (mouse monoclonal), S100B (rabbit polyclonal), and GFAP (rabbit polyclonal) antibodies are described in Table 1. HNA monoclonal antibodies are described in Materials and Reagents.
Antibody or fluorophore combination Purpose of the staining Interpretation of results
NGFR-8737 (human-specific) + S100B (multispecies) • Discriminate SCs that are human (NGFR+) from total SCs (S100B+) regardless of species
• Cells that are S100B+/NGFR+ can be considered hSCs if derived from peripheral nerve
• NGFR-/S100B+ cells should not be regarded hSCs. Suspect of contamination with CNS glia
NGFR (multispecies) + HNA (human nuclei only) • Discriminate hSCs (NGFR+/HNA+) from other human cells (NGFR-/HNA+)
• Cells that exhibit an NGFR-/HNA+ phenotype cannot be considered hSCs
• NGFR+/HNA- cells may consist of nonhuman SCs
NGFR-8737 + GFAP (multispecies) • Discriminate hSCs (NGFR+) from astrocytes (GFAP+) • NGFR-/GFAP+ cells may consist of CNS glia (astrocytic)
NGFR-8737 + FM4-64X (or FluoroMyelinTM) • Confirm the hSC phenotype among NGFR+ cells • FM4-64X+ granules inside NGFR+ cells are strong evidence of the adult, repair-like hSC phenotype
Plate cells from a mixed culture or unknown source as described in Protocol 1A. Limit the numbers to 20,000 cells/well (approx. 10,000 cells/cm2) to achieve a low-density culture. Ideally, the cells should be separated from each other to identify them individually.
Gently aspirate the culture medium, fix the cells for 20 min with fixation solution (see Recipe 4), and wash them 3× with D-PBS.
Permeabilize the cells by incubation with cold (-20 °C) methanol.
Aspirate the methanol quickly and wash the cells 3× with D-PBS.
Add blocking solution (see Recipe 5) for at least 30 min at RT.
Replace the blocking solution with the selected antibody combinations presented in Table 2.
Proceed as described in Protocol 1C, steps 6–7, using the appropriate selection of secondary antibodies and fluorescent dyes (Table 2), followed by image analysis (see Notes a and b).
Notes:
NGFR-8737 reacts strongly and specifically with human/primate NGFR without cross-reacting with NGFR from other species. Cultured NGFR-8737+ cells can be considered human Schwann-like glia if they originate from PNS tissues. Nevertheless, refer to Table 1 to develop alternative antibody combinations to confirm the hSC phenotype. As shown in Figure 3C, S100B+ cells may not be considered Schwann-like without confirming double-labeling with NGFR antibodies. We have not identified the presence of NGFR+, S100B- in established hSC cultures (Peng et al., 2020).
The hSC phenotype can be confirmed by co-immunostaining with HNA and SC-specific antibodies (Tables 1 and 2, Figure 3B). An alternative human-specific nuclear protein is NuMA, which has proven useful for localizing hSCs in the environment of a xenografted host (Bastidas et al., 2017).
Figure 3. Quality control testing of known and unknown cell cultures. A, B. Discrimination of human and rat Schwann cells (SCs) using human-specific NGFR-8737 (A) and HNA (B) antibodies. The proportion of hSCs and rat SCs has differed in A (1 human: 9 rat) and B (9 rat: 1 human). C–I. Immunological assessment of an unidentified cell culture using multiple antibody combinations against glia (NGFR, S100B, and GFAP) and non-glial cell markers (SMA, FN). Cells were treated in the absence (control, C–G) and presence of CPT-cAMP (H, I), an agent that drives SC differentiation (see Protocol 3C). The unidentified human-derived (HNA+) cell culture contained only a few hSC-like cells, as evidenced by co-expression of S100B and NGFR (arrowheads in C2). Contamination with CNS glia was suspected considering the presence of S100B+ (C1–C2) and GFAP+ (G–I) cells that did not co-express NGFR. Notice the strong staining for FN (E) and the induction of SMA in the CPT-cAMP condition, which is a response not expected of hSCs.
Protocol 3: Analysis of bioactivity
For over five decades, researchers have relied on in vitro systems to address the neuron-supportive function and other typical SCs responses during nerve development, maturation, and repair. Traditional bioassays have involved placing hSCs in co-culture with dorsal root ganglia (DRG) neurons to evaluate SC–neuron interactions, such as SC-elicited axon growth (Stratton et al., 2017), axon contact-driven SC proliferation (Morrissey et al., 1995c), and myelin sheath differentiation (Morrissey et al., 1995b). However, researchers noticed that cultured hSCs reduced the viability of neuronal cells and did not differentiate effectively under conditions that supported myelination by non-human (rat) SCs (Morrissey et al., 1995c), and that expanded hSCs failed to align along axons, proliferate, and differentiate when placed in co-culture with DRG neurons despite their strong pro-regenerative phenotype (Monje et al., 2018). Other studies indicated that only a subset of nerve-derived hSCs activated the promyelinating factor POU3F1 and myelinated PNS axons after transplantation (Stratton et al., 2017), and that myelination of axons in the spinal cord was achieved when grafting cultured SCs from rats rather than humans (Bastidas et al., 2017).
The determination of potency for cultured hSCs is complex because there is no common analytical method or bioassay able to address the multiple functions of SCs in vivo. FDA regulations state that surrogate markers that correlate with bioactivity may be used alone or together with cell-based assays for the functional characterization of cellular products (FDA, 2008). In this scenario, an argument is made that the levels of expression and activation of key membrane receptors, intracellular molecules (e.g., kinases), and transcription factors deemed relevant to SC lineage specification and differentiation may be considered surrogate bioactivity indicators for hSCs in vitro. One example from the literature is the consistent use of the transcriptional regulators Egr2/Krox20 and cJun, which drive and inhibit myelination, respectively, to highlight the properties of SCs from intact and injured nerves (Arthur-Farraj et al., 2012).
Therefore, this author recommends a stepwise characterization of hSC identity and function by implementing simple, neuron-free assays prior to launching more sophisticated studies involving co-culture systems or xenotransplantation. We have found that kinase activation (phosphorylation), proliferation, differentiation, and senescence assays (Monje et al., 2018) provide useful information on the characteristics of donor-relevant hSC populations. As explained in the following sections, kinase (Protocol A) and proliferation (Protocol B) assays are designed to evaluate the magnitude of cellular responses to stimulation with heregulin alone and in combination with forskolin, a reversible adenylyl cyclase activator. Differentiation assays (Protocol C) evaluate the expression of myelination-associated markers in response to high doses of cAMP analogs, a potent pharmacological trigger for myelin-related SC differentiation in vitro (Monje, 2018). Senescence assays (Protocol D) evaluate senescence-associated growth arrest, which can be linked to extended passaging or stress-inducing treatments in cultured hSCs.
The proposed cell-based assays share the following features: (1) they are SC-specific, i.e., they inform on the properties expected of SCs rather than other cell types; (2) they are quantitative or semi-quantitative, i.e., the output from the assays is (or can be converted into) a measurable readout; and (3) they assess the magnitude of a cellular or molecular response as a function of a stimulatory signal or experimental condition (i.e., heregulin in kinase activation assays, cAMP in differentiation assays, and rounds of subculture in senescence assays) in reference to a negative control condition (i.e., vehicle-stimulated cells or cells not subjected to passaging). Bioactivity assays should be performed only with purified hSC cultures or cultures whose identity is thoroughly understood (Protocols 1 and 2). Deviations from the normal hSC phenotype, or the presence of contaminating cells, may alter or misrepresent the results. Please, refer to our publications (Monje et al., 2018; Peng et al., 2020) for additional data, technical details, and alternative approaches regarding the optimization and use of the bioassays. Whilst image analysis using fluorescence or light microscopy (this protocol) is useful as the first approach, other methods (western blot, ELISA, and q-RT-PCR) may be advantageous to confirm the results or increase the detection sensitivity.
Determination of β1-heregulin/ErbB-elicited ERK and Akt activation
This procedure describes how to evaluate ERK and Akt phosphorylation using pathway-selective, phospho-specific antibodies in β1-heregulin-stimulated hSC cultures, as reported previously (Monje et al., 2006; Monje et al., 2021b). The responsiveness to β1-heregulin, and the consequent activation of ErbB/HER2 and ErbB/HER3 receptors, are key bioactivity measures for cultured SCs in general. β1-heregulin (a member of the NRG family) is the most potent mitogenic factor described for hSCs (Levi et al., 1995) and induces proliferation by promptly activating ErbB-mediated signaling via Ras-ERK and PI3K-Akt pathways (Monje et al., 2006 and 2008). ERK and Akt phosphorylation are equally reliable readouts for (and correlate with) β1-heregulin-induced cell cycle progression (Monje et al., 2006). This response is expected to be SC-specific because other nerve-resident cell types, such as endoneurial fibroblasts, lack the expression of the ligand binding partner ErbB3 and are unresponsive to the β1-heregulin stimuli. In these assays, a recombinant β1-heregulin peptide is used as a mimetic of the natural axon-bound heregulin/NRG family members known to mediate axon-contact driven SC mitogenesis (Morrissey et al., 1995c). Detection of activated (phosphorylated) ERK and Akt can be completed within 3 h by immunofluorescence microscopy imaging, as per the protocol described below.
Plate the cells as explained in Protocol 1A.
Starve the cells by progressively removing mitogenic factors and serum from the culture medium, as follows: the day after plating, remove the HP medium and replace it with an equal volume of LP medium; 24 or 48 h later, replace the LP medium with starvation medium (D1, see Recipe 3), and incubate the cells overnight before stimulation (see Note a).
Induce ERK and Akt activation by stimulating the cells with recombinant β1-heregulin. To do so, replace the culture medium with DMEM/F12 containing vehicle (control) or β1-heregulin at 10 nM (treatment, single dose). (Optional) Provide β1-heregulin in a range of concentrations from 0.1 to 10 nM to ascertain dose-dependent changes (see Note b).
Incubate the cells for 10 min (fixed time point) in a CO2 incubator. The activation of ERK and Akt by β1-heregulin is maximal at 5–10 min and declines thereafter. However, P-ERK is maintained for 24 h after β1-heregulin addition in hSCs (Monje et al., 2006), so the time course of these experiments can be extended for several hours or even days (Monje et al., 2008).
Rapidly remove the medium by gentle aspiration and fix the cells with methanol (-20 °C) for 15 min.
Proceed as described in Protocol 2C, steps 3–7, except use anti-Phospho(P)-ERK/Total ERK and/or anti-P-Akt/Total Akt antibodies, 1:200–1:300 each in blocking solution. (Optional) If an expedited reading is needed, incubation with primary antibodies can be reduced to 1–2 h at RT.
Analyze the cells by fluorescence microscopy to identify ERK and Akt phosphorylation levels in reference to total kinase levels in β1-heregulin-treated and control cells. Refer to Table 3 for data interpretation and Figure 4 for representative images. See Monje et al. (2006) for additional results.
Table 3. Schwann cells (SC)-specific bioassays. Table 3 summarizes our suggested assays to reveal the unique responses of cultured hSCs. All assays rely on the detection of one or more inducible molecular readouts while exploiting the experimental fine-tuning of an activating signal (input) driving a measurable change (output) through a SC-specific mechanism of action.
Bioassay Expected results Interpretation
Kinase activation (heregulin-
dependent)
• Cultured hSCs respond to β1-heregulin by rapidly activating ERK and Akt.
• The differential immunolabeling between control (without heregulin) and treated cells (heregulin-stimulated) should be obvious in all hSCs.
• Inter-experimental variability is expected due to the rapid kinetics of ERK and Akt phosphorylation, but the overall responses should be consistent across hSC populations. For a reference, include positive controls to reveal maximal ERK and Akt activation.
Proliferation (heregulin and forskolin-dependent)
• β1 heregulin is sufficient to increase the percentage of dividing hSCs. This response is synergistically enhanced by forskolin but forskolin alone does not increase hSC proliferation.
• hSCs incorporate EdU in a nonsynchronous manner starting roughly at 18–20 h post stimulation under these conditions.
• The proportion of proliferating (EdU+) cells is donor- and passage-dependent, but it is usually < 40% in early-passage cultures under the suggested conditions.
• High basal proliferation in the control condition may indicate an excess of fibroblasts or, in rare cases, cancerous (heregulin-independent) hSC proliferation.
Differentiation (CPT-cAMP dependent)
• CPT-cAMP induces upregulation of certain myelin-associated markers (e.g., O4, Krox20, PRX) and downregulation of immature hSC markers (e.g., cJun) within 3–5 days. Corresponding changes in mRNA expression occur at earlier time points.
• CPT-cAMP treated hSCs become post-mitotic and morphologically dissimilar to untreated cells.
• Select the markers that provide the highest resolution for detection. Assessment of myelin gene expression is only relevant when comparing control (no CPT-cAMP) and CPT-cAMP-treated conditions. The magnitude and kinetics of the expression of different gene products are variable. Substantial batch variability is also expected.
• Molecular changes may be evident even when morphological differentiation is not apparent.
Senescence (passage-dependent)
• hSC populations contain a proportion of senescent cells even at passage-zero. Late-passage cultures may consist only of senescent hSCs.
• Fibroblasts do not get senescent under these conditions and can be discriminated from hSCs by their negative SA-β-Gal staining.
• Normal hSC cultures (nerve-derived) become senescent, usually after four rounds of passaging.
• The percentage of senescent cells varies from culture to culture or donor to donor, even when same-passage cultures are compared.
Figure 4. Heregulin-elicited ERK activation in human Schwann cells (hSCs). hSCs were plated, deprived of mitogenic factors and serum, and stimulated as per Protocol 3A. The cells were stained with antibodies against the phosphorylated (P-ERK) and total forms of ERK, respectively. hSCs rapidly respond to β1-heregulin by inducing P-ERK and its shuttling to the nucleus. ERK staining is strong and homogeneous in the hSC’s cytoplasm (right panels, control).
Determination of heregulin-dependent proliferation with and without forskolin
As mentioned above, cultured hSCs are responsive to heregulin/NRG because they constitutively express ErbB2-ErbB3 receptors in the absence of other ErbB/EGFR family members (Morrissey et al., 1995c; Peng et al., 2020). SCs are among the few known cellular types whose proliferation is enhanced, rather than reduced, by pharmacological agonists of intracellular cAMP at low doses. In addition, the effect of cAMP-elevating agents on heregulin-induced mitogenesis is synergistic in SCs. This synergism was discovered during early investigations using in vitro cultured SCs from rats (Stewart et al., 1991) and humans (Levi et al., 1995) and can be regarded as a distinctive SC response. Nerve-derived fibroblasts are not heregulin-sensitive and respond to cAMP agonists by decreasing their rate of cell division (Levi et al., 1995).
The assays described below are designed to determine cell cycle progression (DNA synthesis) in response to β1-heregulin and forskolin, alone and in combination, as reported in Monje et al. (2006, 2008 and 2018), and Peng et al. (2020). These assays are well-suited to determine a hSC-specific outcome in populations confirmed to be pure and non-senescent. The requirement of β1-heregulin as a mitogenic factor is a defining feature of the cultured hSC phenotype. Cells that do not require exogenous β1-heregulin for propagation in vitro should be further scrutinized for their identity or transformed phenotype.
Plate the cells as described in Protocol 1A and starve them of mitogenic factors (Video 1) as described in Protocol 3A.
Video 1. Human Schwann cells (hSCs) deprived of mitogenic factors.hSCs were plated on a PLL-laminin-coated dish in LP medium and imaged using IncuCyte ZOOMTM using 20× objective lenses. This condition serves as a common control for the proliferation (Video 2) and differentiation (Video 3) assays. Notice the fast adhesion of the cells, the changes in cell morphology, and the progressive cell-cell alignment. hSCs proliferated moderately (see mitotic figures) due to serum factors present in the LP medium. Individual phase contrast images were taken every 30 min starting 4 h post-plating. Some relevant elements were highlighted in the video.
Stimulate the cells with β1-heregulin and forskolin. Replace the culture medium with D1 medium containing vehicle (control), 10 nM β1-heregulin (main mitogen), 2 μM forskolin (adjuvant), and 10 nM β1-heregulin plus 2 μM forskolin (growth factor combination to observe a synergistic effect in the number of proliferating cells). (Optional) These assays can be performed in the presence of 10% FBS for maximal hSC viability without hindering the synergistic effect of forskolin (Monje et al., 2018) (see Note c).
Label the newly synthesized DNA with the thymidine analog EdU. Add EdU (1 μM) to all wells to label cells undergoing S-phase entry 4–18 h post stimulation with β1-heregulin and forskolin. (Optional) Add EdU concurrently with the stimulating factors in Protocol 3B step 2 (see Note d).
Incubate the cells in a CO2 incubator for 48–72 h.
Fix the cells and detect DNA-bound EdU by following the manufacturer’s instructions for fluorescent detection of EdU+ nuclei. Add a nuclear dye such as Hoechst-33342 or DAPI to label all nuclei.
Image the cultures by fluorescence microscopy and count the number of EdU+ cells in reference to the total number of cells (DAPI+ or Hoechst+) to estimate the percentage of cell division. Refer to Table 3 for data interpretation, and Videos 1 and 2 for representative results.
Video 2. Human Schwann cell (hSC) proliferation in response to the mitogenic factors heregulin and forskolin. Cells were plated in LP medium for 4 h before being stimulated with heregulin and forskolin (Protocol 3B) and recorded via IncuCyte ZOOMTM, as described in Video 1. Notice the high levels of proliferation (mitotic figures), cell-cell alignment, and migration of hSCs under the influence of mitogenic factors. This culture can be considered confluent and ready for analysis by the end of the recording period.
Determination cAMP-dependent differentiation
hSCs growing in the absence of neurons maintain a highly immature, proliferative phenotype, unless induced to differentiate with a potent stimulus, such as high doses of cAMP analogs. This unique response of hSCs allows the measurement of myelin-associated gene expression in response to a single stimulatory signal without the need to introduce neurons. Researchers should consider that SCs established in culture express negligible or undetectable levels of myelin proteins and lipids. Thus, a strong rationale exists for challenging the hSCs with differentiating factors before measuring the expression of myelin markers.
Our hSC differentiation protocol is based on prolonged incubation with high doses of the cell-permeable, phosphodiesterase-resistant analog of cAMP, CPT-cAMP (Monje et al., 2018). These simple bioassays can substitute for the laborious assessment of myelin sheath formation in neuron-SC culture systems. Use highly viable, non-senescent populations and fresh reagents (e.g., CPT-cAMP stocks) for optimal results. Additional technical details on the preparation of cells, substrates, and media, and the optimization of experimental variables can be found in Monje (2018).
Plate the cells in HP medium as explained in Protocol 1A.
The day after plating, replace the HP medium with an equal volume of D1 or LP medium containing vehicle (DMEM) or CPT-cAMP (250 μM in DMEM), to maintain the cells exhibiting an immature phenotype (control) or to promote differentiation, respectively.
Incubate the cells in a CO2 incubator for 3–5 days with daily observations to identify morphological changes associated with CPT-cAMP stimulation (see Note e). (Optional) Add EdU labeling reagent by the second- or third-day post-CPT-cAMP addition, to concurrently determine myelin gene elevation and cell cycle exit.
Proceed to stain the cells with antibodies that recognize the state of differentiation of the hSCs, e.g., by O4 labeling (Protocol 1B) or immunostaining for myelin-related markers (Protocol 1C). See Table 1 and Figure 5 for antibody options.
Figure 5. Differentiation assays. Human Schwann cells (hSCs) were stimulated and analyzed per Protocol 3C and stained with antibodies per Table 1. Low (A–B) and high magnification (C–J) images of control (non-stimulated cells in D1) and cAMP-differentiated hSCs (cells treated with 250 μM CPT-cAMP in D1 for 7 days) are shown. Notice the phenotypic conversion of the hSCs from an elongated, roughly bipolar shape to an enlarged, flattened morphology in the CPT-cAMP condition. CPT-cAMP treatment does not change S100B (A–D) and NGFR (C–D, I–J) expression. However, a proportion of the CPT-cAMP-treated hSCs express higher levels of Krox20 (G–H), periaxin (PRX, E–F), and O4 (E–H) along with lower levels of nuclear cJun (I–J), as denoted by the white arrowheads. Heterogeneity in the expression levels of myelin markers is expected (H–F). The S100B negative cells highlighted by the black arrowheads in panels A–B are fibroblasts. Nuclei were stained with DAPI (blue, C–F). The PRX antibody was courtesy of Peter Brophy. The Krox20 antibody was courtesy of Dies Meijer.
Add a nuclear dye such as Hoechst-33342 or DAPI to label all nuclei.
Image the cultures by fluorescence microscopy and count the number of O4+ cells (or the marker of choice) in reference to the total cells to estimate the differentiation efficiency.
Refer to Table 3 for data interpretation. See Figure 5 and Videos 1 (control) and 3 (CPT-cAMP) for representative results.
Video 3. Human Schwann cell (hSC) differentiation in response to CPT-cAMP.Cells were plated in LP medium for 4 h before being stimulated with CPT-cAMP (250 µM) per (Protocol 3C). The plates were transferred to IncuCyte ZOOMTM for live cell imaging immediately after CPT-cAMP stimulation. Notice that CPT-cAMP inhibits hSC proliferation, alignment, and migration while inducing a flat, expanded cytoplasm that features transient vacuoles of different sizes. This culture is ready to be analyzed for the presence of myelin-related markers by the end of the incubation period. Some relevant elements were highlighted in the video. For a reference, compare this video with Video 1 (control condition).
Determination of passage-dependent senescence
SCs of human origin undergo senescence as they are passaged in vitro or exposed to various stressors, including environmental changes. The causes of senescence are ill-defined. Yet, senescence can be considered an hSC-specific attribute, as senescence does not affect human fibroblasts from nerve tissues (Peng et al., 2020). Intriguingly, cultured SCs established from rat nerves can expand almost indefinitely without getting transformed or acquiring senescence (Mathon et al., 2001). A hypothetical hSC culture that fails to senesce under standard in vitro conditions is likely not human or not SC-related. By determining the proportion of senescent hSCs, one can predict the progression (expandability) of donor-relevant cell cultures.
We recommend a generic enzymatic SA-β-Gal test for senescence detection because markers indicative of hSC senescence are elusive (Monje et al., 2021a). Senescent and non-senescent cultures are hard to discriminate (Figure 6 and Video 4). Perform senescence assays on populations that have been expanded for several passages, become unresponsive to proliferate with SC-specific mitogens, or manifest morphological or functional changes, such as excessive cell clumping, detachment, or floating debris.
Figure 6. Discrimination of expandable and non-expandable (senescent) human Schwann cell (hSC) cultures. This figure shows an example of how to functionally discriminate non-senescent (Donor-1) vs. senescent (Donor-2) cultures based on results from SA-β-Gal (A–B) and EdU incorporation (C–D) assays in cultures growing in HP medium. hSCs from these two donors are visually undistinguishable based on their morphology, pattern of alignment, and levels of SC-specific markers such as S100B (E–F). Most cells from Donor-2 are non-proliferative and stain positive for SA-β-Gal (black arrowheads in B). Estimating the percentage of senescent cells, alone or together with EdU staining, can help to predict the characteristics of cells from subsequent rounds (Monje et al., 2018). Notice the dissimilar levels of EdU labeling in individual cells, indicating the asynchronous incorporation of DNA. Some individual cells (white arrowheads) have undergone more than one cycle of cell division.
Video 4. Senescent human Schwann cell (hSC) cultures.The cells were plated in HP medium and recorded via IncuCyte ZOOMTM every 20 min. Notice that senescent hSCs are highly viable and migratory but fail to efficiently expand in number despite the presence of heregulin and forskolin. The appearance of vacuoles (seen clearly at the onset), the expanded cytoplasm, and the lack of cell-cell alignment are morphological evidence of senescence. However, senescence is defined by the lack of proliferation and should be confirmed with appropriate assays. The senescent phenotype is irreversible. Some relevant elements were highlighted in the video. For a reference, compare this video with Video 2, featuring a highly proliferative hSC culture.
Plate the cells in HP medium as explained in Protocol 1A. Include a known population of senescent cells in all SA-β-Gal experiments to serve as positive control. hSCs from passage-4 or higher and/or hSCs treated with H2O2 for >3 days are suitable, as environmental stressors readily drive hSC senescence.
Incubate the cells in a CO2 incubator for at least 24 h to ensure all cells are well-adhered and extend processes on the substrate (see Video 4).
Fix the cells with fixation solution (see Recipe 4) for 20 min as explained in Protocol 1B.
Reveal the enzymatic activity of SA-β-Gal by following the manufacturer's recommendations.
Add a nuclear dye such as Hoechst-33342 or DAPI to label all nuclei.
Image the cultures by light microscopy (SA-β-Gal) and fluorescence microscopy (nuclei) and count the number of SA-β-Gal+ cells in reference to the total number of cells. Refer to Table 3 for data interpretation. Figures 6 and 7, and Video 4 show representative results.
Figure 7. Identification of fibroblast contamination and senescent cells. Examples are shown of how to recognize cultures enriched in fibroblasts (left panels) and senescent cells (right panels) by immunostaining with cell type–specific markers (FN, green; S100B, red) and enzymatic SA-β-Gal staining (blue precipitate inside the hSCs), respectively. The cultures were grown in LP medium in the absence (control, A–B) or presence of 250 μM CPT-cAMP (Protocol 3C, C–D) for seven days before fixation and immunostaining (Protocol 1C). Even though hSCs differentiated effectively as denoted by S100B staining, the fibroblasts proliferated extensively (C–D). In E–H, hSCs (passage-5) were plated in HP medium, fixed, and stained with SA-β-Gal after three days (Protocol 3D). Observe the differential behavior of the cells in areas of low (E–F) and high (G–H) density. Senescent hSCs are migratory and have the capacity to form aggregates that are prone to detachment. Recommendations: (1) purify the cells from A–D; (2) use or cryopreserve the cells from E–H but do not continue the culturing, as nearly all hSCs are senescent.
Notes:
Progressive removal of mitogenic factors and serum is needed to lower the endogenous levels of phosphorylated kinases and induce hSC quiescence without massive apoptotic cell death triggered by loss of trophic support. Starved hSCs maintain their alignment and general morphology, even though their processes become thinner (Figure 1 and Video 2).
This protocol is equally suitable to determine the activation status of other β1-heregulin-responsive kinases such as ErbB-2, ErbB-3, Raf, MEK, and RSK (Monje, Bartlett Bunge et al. 2006). Always include a positive control condition to activate ERK and Akt. The phorbol ester PMA, an activator of PKC, can be used to elicit maximal ERK activation in all cells. Purified PIP2,3 can be used to activate Akt in a membrane receptor-independent manner (Monje, Athauda, et al.2008).
Alternative ErbB- and cAMP-inducers may be considered. ErbB-dependent hSC proliferation can be elicited with axolemmal preparations enriched in membrane-bound NRG, a mitogen for cultured hSCs (Sobue et al., 1984). Cholera toxin, pertussis toxin, and phosphodiesterase-resistant analogs of cAMP (e.g., db-cAMP) are effective in promoting a synergistic effect on heregulin-induced hSC proliferation (Levi et al., 1995; Monje et al., 2006)
EdU incorporation assays are a suitable alternative to radioactive (e.g., tritiated thymidine) and antibody-based (e.g., BrDU) methods. EdU detection uses mild fixation and can be multiplexed with antibody labeling. Results from EdU incorporation assays are best interpreted when done in conjunction with immunostaining (Protocols 1B–C) and senescence assays (Protocol 3D). We routinely stain cells with antibodies (e.g., anti-NGFR or anti-O4) before developing the EdU reaction to readily discriminate proliferative hSCs from other cell types (Monje et al., 2018).
Typically, hSCs lose their spindle-shaped morphology and acquire a large, reticulated shape coincidently with cell cycle exit within 2–3 days after CPT-cAMP treatment (Video 3). If morphological differentiation is confirmed by phase contrast microscopy, the cells can be considered ready for analysis. If morphological differentiation is not appreciated by the third or fourth day of treatment, new medium containing CPT-cAMP should be added, and additional time should be allowed for differentiation to occur.
Recommendations and troubleshooting
Clinical investigations have revealed that with optimized standard operating procedures, clinical-grade reagents, and systematic approaches, it is possible to consistently achieve hSC cultures at high yields (e.g., 108 cells at passage-2) and purity (> 90% hSCs) without introducing purification steps (Khan et al., 2021). In non-clinical settings, however, the level of control is much lower, starting with the conditions for tissue procurement, which are usually incomplete or unknown to the investigators. The importance of applying good cell culture practices cannot be more emphasized. Routine monitoring and periodic testing of the cultures using benchmark verification tests and assays can allow researchers to understand and correct problematic issues as soon as they arise. Unlike rat SCs, hSC cultures are sensitive to various experimental variables, such as changes in the media or substrate. Considerable donor- or batch-related variability is expected. Additionally, the cells within single cultures are heterogenous, thus indicating the need to use various analytical tools to evaluate the quality and stability of donor-relevant hSC cultures.
This section was elaborated to assist in the detection and resolution of technical problems that are commonly seen in hSC cultures (Table 4). Running appropriate tests (Protocols 1–2) and bioassays (Protocol 3) is most important in the following scenarios: (1) when the cultures acquire an unusual appearance or show a distorted alignment pattern, (2) when there is excessive floating debris, and (3) when the rate of proliferation changes or cell growth becomes density independent. For a reference, Figures 6–7 and Videos 4–5 present examples of cultures that merit further scrutiny due to their atypical features attributable to senescence and fibroblast contamination.
Table 4. Troubleshooting guide for hSC cultures
Potential problem Possible Cause Corrective measure
Cultures do not expand at the expected rate Ineffective mitogenic cocktail, or arrival to senescence (Figures 6-7). Test the activity of heregulin and forskolin using specific bioassays (Monje et al., 2021b) or simply replace the stock solutions and perform a senescence test.
Cultures do not acquire the typical pattern of growth or bipolar morphology This is normal if cultures are not yet confluent but may reflect problems such as over-proliferation of fibroblasts (Figure 7), cytotoxicity, or senescence. Continue the culturing and provide new HP media to expedite hSC division. If this problem persists, perform a purity check along with viability assays.
Cells do not adhere or extend membrane processes after plating This is likely the result of poor adhesion and may indicate a problem with the substrate. Short or retracted processes may indicate stress and poor viability (e.g., due to over-trypsinization). Perform a quick viability test. If cells are alive, transfer the cell suspensions to a fresh laminin-coated dish within 2–4 h post-plating. Otherwise, hSCs would undergo apoptosis triggered by lack of adhesion (anoikis).
There is excessive floating debris, and the medium contains particulate material This is a usual phenomenon in some hSC cultures but may also indicate cytotoxicity or microbial contamination. Wash the cells with D-PBS to remove the debris, add new HP medium, and perform a sterility test to discriminate this condition from microbial contamination.
Signs of microbial contamination become clear This can be manifested at any time due to unknown contamination of the nerve explant or an improper aseptic technique during the culturing steps. Discard the contaminated hSC cultures. Perform proper tests to identify the type and source of contamination. Decontamination attempts are futile. Discard the stocks if contamination is traced to the stocks.
The cells migrate to certain areas, form clumps, and detach from the substrate This is usual in hSC cultures and may indicate stress due to factors in the medium or substrate. Clumping may result from over confluence or senescence (Figure 7). This may be corrected by replating the cells into a new dish. The culture may be terminated if the cells are senescent.
Cultures become enriched in senescent cells Cells have reached their expansion limit, or environmental factors have triggered premature senescence. Discard the cultures or use them as such. Senescent cells are highly viable and bioactive, but proliferation is irreversibly halted (Figures 6 and 7).
Cells do not adhere after being seeded from a cryogenic stock, or there is an excess of apoptotic cells This can indicate an adhesion problem (substrate), cytotoxicity (to the DMSO) during and after plating, or poor cryopreservation (i.e., suboptimal storage conditions). Confirm the viability of the stocks. Take precautions to work fast and eliminate traces of DMSO after plating. Recover the culture after removing the apoptotic cells.
Cultures do not contain O4+ hSCs This is usual. O4+ cells decline with expansion. O4 expression is not constitutive but induced in response to cAMP agonists and axon contact. Confirm the identity of the cultures by immunostaining. Add forskolin or other cAMP-stimulating agents for at least 72 h to restitute O4 expression (Figure 5).
Fibroblasts proliferate fast and overtake the culture This issue occurs when the selective pressure of cAMP-stimulating agents is removed (Figure 7 and Video 5). Fibroblasts are resistant to undergo senescence. Restitute the mitogenic combination to maintain high levels of cAMP and reduce fibroblast growth. Purify the cells if fibroblast contamination becomes problematic.
Cells align well and cover the available surface, but cell counts remain low This common observation indicates poor proliferation or enrichment in senescent cells. Test the bioactivity of the mitogenic factors and perform a senescence test.
Cells proliferate in the absence of heregulin and lose density-dependent control of proliferation This is unusual in normal hSC cultures. It can indicate transformation or cross-contamination with a highly proliferative, cancerous cell line. Perform identity and authentication tests to verify the source of the proliferative cells. If the cells are confirmed to be hSCs, evaluate the possibility of transformation via appropriate tests.
Video 5. Fibroblast contamination. Human Schwann cells (hSCs) were plated in LP medium and imaged via IncuCyte ZOOMTM every 30 min. This culture contained a high proportion of fibroblasts, which can be recognized by their expanded (low contrast) cytoplasm and conspicuous nuclei. The hSCs extended processes but failed to align with one another, possibly due to the presence of fibroblasts concentrating in the central area. This culture has abundant cellular debris. The over-proliferation of fibroblasts and the debris can signal an underlying issue with the hSC culture (see Table 4). Some relevant elements were highlighted in the video. For a reference, compare this video with Video 1, which displays a pure hSC culture under the same culture conditions.
Recipes
Low proliferation medium (LP)
Reagent [stock concentration] Final concentration Volume
DMEM (or DMEM/F12) n/a 445 mL
FBS (100%) 10% 50 mL
GlutaMax 200 mM (100×) 1× 5 mL
Gentamycin (1,000×) 1× 0.5 mL
Total n/a 500 mL
Note: Balance the pH to make it slightly acidic (pH = 6), sterilize it using a 0.22 μm filter, and store it at 4 °C. Follow the guidelines provided in the accompanying paper (Monje, 2023).
High proliferation medium (HP)
Reagent [stock concentration] Final concentration Volume
Low proliferation medium n/a 500 mL
Heregulin-β1177-244 (25 μM) 10 nM 200 μL
Forskolin (15 mM) 2 μM 69.25 μL
Total 500 mL
Note: Balance the pH to 7, filter, and store the HP medium as indicated for the LP medium. More information can be found in Monje (2023).
Starvation or D1 medium (DMEM/F12 – 1% FBS)
Reagent [stock concentration] Final concentration Volume
DMEM/F12 n/a 45 mL
FBS (100%)
GlutaMax 200 mM (100×)
HEPES 1 M (100×)
Gentamycin 50 mg/mL (1,000×)
Total
1%
1×
1× (10 mM)
1×
n/a
0.5 mL
0.5 mL
0.5 mL
0.05 mL
50 mL
Note: Balance the pH to 7, filter, and store as indicated for the LP medium. This medium is used only for controlled starvation, as hSCs are affected by the absence of serum. Do not freeze.
PFA-based fixation solution
Reagent [stock concentration] Final concentration Volume
PFA (20%) 4% 4 mL
DPBS 1× 16 mL
Total 20 mL
Note: The stocks are maintained frozen at -20 °C until use. Avoid repeated freeze-thaw cycles. Working solutions can be stored at 4 °C.
Blocking solution
Reagent [stock concentration] Final concentration Volume
Normal goat serum 5% (v/v) 5 mL
DPBS 1× 1× 45 mL
Total 50 mL
Note: The stocks are maintained frozen at -20 °C until use. Avoid repeated freeze–thaw cycles.
Acknowledgments
Natalia Andersen, Ketty Bacallao and Kaiwen Peng contributed to the experimental work. Patrick M. Wood, Mary Bunge, and Linda White provided expert guidance, troubleshooting tips, and banked cells during the initial investigations. The development of these protocols was supported by grants (to P.V.M.) from the National Institutes of Health (NS084326), the Craig H Neilsen Foundation (339576), The Miami Project to Cure Paralysis and the Buoniconti Foundation from the University of Miami, the Charcot Marie Tooth Association, and the Indiana State Department of Health (33997, 43547). Grant NIH/NS009923 (to M.B., P.M.W and P.V.M) provided seminal funding for developing proliferation and differentiation assays and the creation of cell stocks. The contents of this article are the responsibility of the author and do not necessarily represent the official views of the funding agencies. Valeria Nogueira assisted with illustrations, Gabriela Aparicio with manuscript and figure formatting, Thomas Dolan with video editing, and Louise Pay with English editing. Lingxiao Deng contributed fruitful scientific discussions and laboratory support. We are greatly indebted to the generosity of the anonymous individuals and their families for providing tissues for research.
Competing interests
P.V.M. is the founder of GliaBio LLC., a consulting company that focuses on glial cell research. The author declares that the research described here was conducted in the absence of commercial or financial relationships that could be construed as a potential conflict of interest.
Ethics
The experiments were performed using anonymized, banked hSC cultures from non-pathological tissues without regard to the gender or age of the donor. This research was deemed to constitute non-human-subject research by the Institutional Human Subjects Research Offices of the University of Miami (2003–2018) and Indiana University (2019–2022). The human cells used in these studies were derived from postmortem tissues made available by the Life Alliance Organ Recovery Agency (LAORA) of the University of Miami to the laboratory of P.M.W. Requests for information and materials (cells, antibodies, other) should be directed to P.V.M. Investigators are encouraged to contact P.V.M. for feedback on the reported protocols.
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Production, Extraction, and Solubilization of Exopolysaccharides Using Submerged Cultures of Agaricomycetes
LG Lina R. Dávila Giraldo
PB Paula X. Villanueva Baez
CF Cristian J. Zambrano Forero
WA Walter Murillo Arango
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4841 Views: 609
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Abstract
Macrofungi, also known as mushrooms, can produce various bioactive compounds, including exopolysaccharides (EPS) with distinct biological properties and subsequent industrial applications in the preparation of cosmetics, pharmaceuticals, and food products. EPS are extracellular polymers with diverse chemical compositions and physical properties secreted by macrofungi in the form of capsules or biofilms into the cellular medium. Submerged cultivation is an industrially implemented biotechnological technique used to produce a wide variety of fungal metabolites, which are of economic and social importance due to their food, pharmaceutical, and agronomic applications. It is a favorable technique for cultivating fungi because it requires little space, minimal labor, and low production costs. Moreover, it allows for control over environmental variables and nutrient supply, essential for the growth of the fungus. Although this technique has been widely applied to yeasts, there is limited knowledge regarding optimal growth conditions for filamentous fungi. Filamentous fungi exhibit different behavior compared to yeast, primarily due to differences in cell morphology, reproductive forms, and the type of aggregates generated during submerged fermentation. Furthermore, various growing conditions can affect the production yield of metabolites, necessitating the development of new knowledge to scale up metabolite production from filamentous fungi. This protocol implements the following culture conditions: an inoculum of three agar discs with mycelium, agitation at 150 rpm, a temperature of 28 °C, an incubation time of 72 h, and a carbon source concentration of 40 g/L. These EPS are precipitated using polar solvents such as water, ethanol, and isopropanol and solubilized using water or alkaline solutions. This protocol details the production procedure of EPS using submerged culture; the conditions and culture medium used are described. A detailed description of the extraction is performed, from neutralization to lyophilization. The concentrations and conditions necessary for solubilization are also described.
Key features
• Production and extraction of EPS from submerged cultures of mycelial forms of macrofungi.
• Modification of the method described by Fariña et al. (2001), extending its application to submerged cultures of mycelial forms of the macrofungi.
• Determination of EPS production parameters in submerged cultures of mycelial forms of macrofungi.
• EPS solubilization using NaOH (0.1 N).
Graphical overview
Keywords: Fungal exopolysaccharide Fungi Submerged culture Macrofungi Polysaccharide
Background
Macrofungi are fungi that produce reproductive bodies that are visible to the naked eye. Some macrofungi known for their production of polysaccharides are the Shiitake (Lentinula eodes) or the mushroom turkey tail (Trametes versicolor). Macrofungal polysaccharides are a group of compounds of great interest in bioprospecting studies of both medicinal and edible fungi, owing to their broad spectrum of bioactivities, including cytotoxic, antiproliferative, antitumor, immunomodulatory, anti-inflammatory, antioxidant, and stimulatory effects, on the colon microflora (Arora and Tandon, 2015; X. Meng, 2016; Nowakowski et al., 2021). These bioactivities are related to the chemo-preventive action of macrofungal polysaccharides against different types of cancers such as colorectal cancer and other chronic diseases. Macrofungi polysaccharides are complex carbohydrates found in the cell walls and extracellular matrices. These polysaccharides play a crucial role in the structural integrity and functionality of fungal organisms. They can be homopolysaccharides that contain repeating units of the same monosaccharide or heteropolysaccharides, containing several types of monosaccharides. Polysaccharides can also include other groups, such as sulfate or glucuronate, and adopt simple, triple, or random helical conformations depending on the conditions of the culture medium (Huang and Nie, 2015; Gong et al., 2020), significantly affecting their solubility in water or marginally basic aqueous solutions. Solubility and the type of conformation adopted are crucial for biological activity and must be controlled in different assays at the in vitro level because of their effects on the expected results. Structural conformation is also responsible for the great water solubility of polysaccharides: for example, scleroglucan secreted by Sclerotium fungi adopts a highly ordered, rigid, triple helical tertiary structure when dissolved in water at room temperature and low concentrations of alkali, usually below 0.15 M NaOH (Castillo et al., 2015). Water solubility of fungal exopolysaccharides (EPS) depends, to a large extent, on monosaccharide composition and structure and size of the respective molecules (Jaros et al., 2018). These variables, together with differences in the type of bonding and branching, tertiary structure, electrical charge and conformation, and solubility generate a diversity of physical and chemical properties, as well as reported biological activities (Bacic et al., 2009).
Polysaccharides produced by macrofungi can be classified into three main groups based on their location in the cell: (1) cytosolic polysaccharides that provide a source of carbon and energy for the cell; (2) cell wall polysaccharides that include the peptidoglycans, teichoic acids, and lipopolysaccharides; and (3) extracellular polysaccharides that are exuded in the form of capsules or biofilms, also known as EPS, which constantly diffuse into the cell wall and cell cultures (Kambourova et al., 2015; Wang et al., 2022). For the production of EPS, the submerged culture technique is used at both laboratory and industrial scales, where the mycelium of the fungus grows while floating as small spheres called pellets when there is agitation (Papagianni, 2004). The submerged fermentation process is widely used for producing different types of metabolites on a large scale or under flask conditions in which organisms can be obtained in limited physical spaces under conditions controlled and optimized for the production of biomass; another advantage is the ease of separating the mycelium from the EPS in the culture medium, obtaining a product of uniform quality for their purification in a further step. On the other hand, it provides several advantages such as higher productivity and yields, lower labor costs, and lower contamination risk. Submerged culture offers the potential advantages of faster production of EPS within a smaller space, reduced possibility of contamination, and better control of cultivation conditions such as carbon source and temperature (Dudekula et al., 2020).
In addition to submerged cultures, several other methods have been employed for the production of EPS. Some of these methods are: (1) surface culture, in which the microorganisms are grown on the surface of a solid substrate, such as agar or other suitable media; the EPS is produced and secreted by the microorganisms onto the surface of the substrate (Breitenbach et al., 2022); (2) solid-state fermentation, which involves the growth of microorganisms on solid substrates without the addition of free-flowing water; the microorganisms utilize the nutrients present in the solid substrate to produce EPS; common solid substrates used include agricultural residues, cereal grains, and sawdust (Isikhuemhen et al., 2014); (3) biofilm reactors, which provide an ideal environment for the growth of biofilms, allowing the microorganisms to attach to a surface and form a structured matrix; EPS production occurs with the biofilm structure (Seneviratne et al., 2008); (4) immobilized cell systems, in which the microbial cells are immobilized or attached to a solid support matrix, such as alginate beads, polyurethane foam, or glass beads; the immobilized cells produce EPS while attached to the support matrix, allowing for easy separation and recovery of the EPS (Li et al., 2017); and (5) membrane bioreactors, which combine the use of submerged culture with the retention of microbial cells and EPS using a membrane filtration system; the membrane allows the culture medium to pass through while retaining the microbial cells and EPS, which can then be harvested and recovered (Brito et al., 2019). It is important to note that the method of choice for EPS production depends on various factors, including the microorganism used, the type of EPS desired, and the scale of production. Different methods offer distinct advantages and disadvantages in terms of yield, productivity, cost, and ease of operation.
Extensive investigations have been conducted to enhance the yield of EPS production. These studies have focused on optimizing various factors, including the assessment of different carbon sources (e.g., glucose, sucrose, fructose) and nitrogen sources (e.g., peptone, yeast extract, ammonium nitrate). The influence of pH levels and incubation time on scleroglucan production has also been thoroughly examined. Notably, Fariña et al. (2001) successfully identified the optimal conditions for scleroglucan production by Sclerotium rolfsii. Their research determined the most advantageous carbon and nitrogen sources, optimal pH range, and incubation time, resulting in significant improvements in scleroglucan yields. Our approach differs from the method described by Fariña et al. (2001), as we evaluate the impact of two carbon sources. Moreover, we consider the contribution of the mushroom species as a crucial factor in EPS production. Incorporating a comprehensive range of tests during initial evaluation enables the identification of optimal strains for bioactive EPS production (Mahapatra and Banerjee, 2013; Angelova et al., 2022). Among agaricomycetes, Agaricales and Polyporales are the two most prominent and extensively used orders in biotechnology. To validate this protocol, representative strains from Agaricales and Polyporales were selected, namely Schizophyllum radiatum and Lentinus crinitus, respectively.
Materials and reagents
Petri dishes (90 mm × 15 mm) (Nest, catalog number: 752001)
Polypropylene microcentrifuge tubes (2.0 mL) (Eppendorf, Biologix, catalog number: 80-0020)
Paper towel
Conical tubes (50 mL) (Falcon, catalog number: 602002-1)
Erlenmeyer flask (250 mL)
Beakers (10 and 500 mL) (Glassco Aleman, catalog number: 229.202.02, 229.202.08A)
Borosilicate glass bottle with screw cap with rubber/Teflon liner (250 mL) (Glassco, catalog number: 6679.269.245.01)
Nichrome inoculating needle (Citoplus, catalog number: 33210001B)
Filter paper, pore size: 3 μm (Munktell Ahlstrom, catalog number: 391)
Filter paper, pore size: 10 μm (Ahlstrom Munksjo, catalog number: 389)
Pistil (Axygen, Pes15b)
Core samplers (diameter: 5 mm)
Magnetic stir bar (KartellTM, catalog number: 0076800)
Biological materials
Schizophyllum radiatum Fr. (Fungario Universidad del Tolima; Laboratory collection, strain 030, voucher: LRD27)
Lentinus crinitus (L.) Fr. (Fungario Universidad del Tolima; Laboratory collection, strain 147, voucher: ZF37)
Reagents
Agar agar (Sigma-Aldrich, catalog number: BCBS4444V, 05040)
Peptone (EMD, Millipore, catalog number: WM879031-906BD)
Yeast extract (OXOID, catalog number: LP0021)
Malt extract (OXOID, catalog number: LP0039)
D-Glucose (EMD Millipore, catalog number: K51794437 004)
D-Sucrose (Fisher Scientific, catalog number: S5-500)
Sodium hydroxide (NaOH) (EMD Millipore, catalog number: B1101198-436)
Magnesium sulfate heptahydrate (MgSO4·7H2O) (Honeywell Specialty Chemicals Seelze, catalog number: 10314903)
Iron (II) sulfate heptahydrate (FeSO4·7H2O) (Sigma-Aldrich, catalog number: F7002)
Ammonium sulfate [(NH4)2SO4] (PanReac Química SLU, catalog number: A1032)
Anhydrous dipotassium hydrogen phosphate (K2HPO4) (Mallinckrodt Baker, catalog number: 3252-01)
Ethanol, 96% (Fábrica de Licores Tolima)
Solutions
Malt agar medium (see Recipes)
SYAMI culture medium (see Recipes)
Sodium hydroxide (NaOH) 0.1 N (see Recipes)
Recipes
Malt agar medium
Reagent Final concentration
Peptone 3 g/L
Malt extract 30 g/L
Agar-agar 15 g/L
SYAMI culture medium
Reagent Final concentration
MgSO4·7H2O 0.05 g/L
K2HPO4 2.0 g/L
FeSO4·7H2O 0.2 g/L
(NH4)2SO4 0.15 g/L
Yeast extract 2.0 g/L
Carbon source (glucose or sucrose) 40 g/L
Sodium hydroxide (NaOH) 0.1 N
Reagent Quantity
NaOH 1 g
H2O 250 mL
Equipment
Centrifuge (HERMLE Labortechnik GmbH, Z326K, catalog number: 311)
Orbital shaker (IKA LABORTECHNIK, KS250 basic)
Fume hood (Jpinglobal, catalog number: JPCEGH1200-BB-PP)
Laminar flow cabinet, horizontal (Purificación y análisis de fluidos)
Ultra-low temperature freezer, -85 °C (Kaltis, catalog number: 390)
Freeze dryer (Buchi, model: Lyovapor L-200)
Autoclave (All American, model: 1941X)
Refrigerator (COLDLINE, model: FORTE V13-RHC)
Barnstead Smart2Pure UV/UF (Thermo Scientific, type: Smart2Pure 3 UV/UF)
Hot plate magnetic stirrer (Stuart, catalog number: UC152)
Analytical balance (KERN, type: ACS 220-4)
Blender (Recco, model: 242748)
Drying stove (Memmert, catalog number: ref 1470)
Desiccator (Metacrilato 250 mm)
Sieve (Endecotts Ltd., ASTM E-11, catalog number: 65803- TAN AC8-1-1/2, 37,3)
Borosilicate glass filtration system (Glassco, catalog number: COL-764-260.202.01)
Incubator, 28 °C (Binder, catalog number: BD023UL)
pH meter (SI Analytics, type: HandyLab 100)
Water bath (Buchi, catalog number: B-480)
Software and datasets
Microsoft Excel for calculations
RStudio Team, 2015. RStudio: Integrated Development Environment for R. Boston, MA. Available at: http://www.rstudio.com/
Procedure
Production of EPS using submerged cultures
S. radiatum and L. crinitus culture preparation.
Inoculate each fungal strain from a 4 °C distilled water stock onto solid malt agar medium in a 90 mm × 15 mm Petri dish and incubate at 28 °C until the plate attains full growth for 4–7 days (see Graphical overview, step 1). It is also possible to use other Petri dish sizes.
After full growth, cut the solid malt agar medium with a sterile core sampler (5 mm diameter) and transfer three discs of each fungal strain to 2.0 mL polypropylene microcentrifuge tubes, using a sterile nichrome inoculating needle.
Add 1 mL of sterile distilled water and disperse the agar discs using a pistil. Another option to disperse is to use a stainless-steel immersion blender for 3 s.
Transfer the mixture to a 250 mL Erlenmeyer containing 50 mL of SYAMI culture medium previously autoclaved at 15 psi for 15 min and incubate by shaking at 150 rpm and 28 °C for 72 h or another specific duration; this time depends on the experimental design used and the study strain. For example, S. radiatum strain can produce EPS at 72 h and its maximum production is reached at 144 h. L. crinitus strain can produce EPS at 144 h. Other fungal spices may require more time for EPS production.
Note: Work under laminar flow cabinet.
EPS extraction
Neutralization and enzyme inactivation.
After the incubation time, add the culture to a 500 mL beaker and measure the pH. The pH can be between 4 and 5.
Add sterile distilled water in a 2:1 ratio (100 mL of H2O to 50 mL of culture).
Homogenize the cultures (mycelia and liquid) using immersion blender (two pulses).
Neutralize the mixture (pH = 7.0) using NaOH (0.1 N).
Transfer the mixture to a 250 mL borosilicate glass bottle and incubate in a water bath for 30 min at 80 °C.
Note: Work under fume hood.
Weighing mycelial biomass and EPS.
Place 20 mL of the mixture in 50 mL conical tubes. In the following steps, the weight of the mycelial biomass and EPS will be determined.
Centrifuge at 10,610× g for 30 min at 4 °C.
Transfer the supernatant to a fresh 50 mL conical tube. Add cold ethanol (99.8%) to the supernatant in a 1:1 ratio for EPS precipitation. Store this mixture and biomass pellet at 4 °C for 12 h.
Transfer the precipitated EPS and biomass pellets obtained by centrifugation to quantitative filter paper discs and place them on 100 mm × 15 mm Petri dishes that were previously weighed. Place each sample on a different filter/Petri dish.
Incubate the Petri dishes in a drying stove at 100–110 °C for 2 h.
Place the Petri dishes in a desiccator for 30 min.
Subsequently, weigh the Petri dishes for each sample (EPS precipitated and biomass pellet).
Use the following formula to calculate the dry weight of biomass and EPS:
Dry weight = (C - A)/(B - A) × 100
where A is the weight of the empty filter paper/Petri dish, B is the weight of the filter paper/Petri dish with the fresh sample, and C is the weight of the filter paper/Petri dish with the dried sample.
EPS purification.
Filter 130 mL of the mixture derived from step B1e using a quantitative paper (pore size: 10 μm). Use a borosilicate glass filtration system.
Repeat the filtering step twice using a quantitative paper (pore size: 3 μm). Use a borosilicate glass filtration system. This prevents the passage of small fragments of mycelium.
Transfer the mixture to 50 mL conical tubes.
Centrifuge at 10,610× g for 30 min at 4 °C.
Transfer the supernatant to a 500 mL borosilicate glass bottle.
Add cold ethanol (96%) in a 1:1 ratio for EPS precipitation. Incubate at 4 °C for 12 h (Figure 1a).
Figure 1. Precipitation, lyophilization, and solubilization of exopolysaccharides (EPS) obtained from submerged cultures of macrofungi
Separate the EPS using a 40 μm sieve. The EPS will be suspended on top of the sieve. Use a container under the sieve to collect the ethanol (see Graphical overview, steps 8, 9, and 10). It is also possible to use filter paper of the same pore size.
Add sterile distilled water and repeat steps B3f and B3g.
Place the obtained EPS in an ultra-low temperature freezer (-85 °C, 2 h).
Freeze-dry the EPS (see Table 1).
Table 1. Parameters for lyophilization of macrofungi exopolysaccharides (EPS)
Dried material Strains Freezing parameters Drying shelf temperature regulation Pressure of the chamber Drying time
EPS S. radiatum and L. crinitus -50 °C up to 60 °C 0.01 mbar 72 h
Determine the weight of the freeze-dried EPS (Figure 1b).
Solubilization
Preparation of concentrated EPS solution.
Weigh 100 mg of lyophilized EPS in a 50 mL beaker.
Add 1.25 mL of NaOH (0.1 N) and 10 mL of sterile distilled water.
Dissolve the mixture using a magnetic stirrer for 1–2 h. Do not heat.
Add sterile distilled water to a final volume of 25 mL.
The final concentration is 4 mg/mL.
The final concentration of NaOH is 0.005 N.
You can prepare solutions at different concentrations as required by assays.
Consider the volume of NaOH (0.1 N) used because it can modify the EPS structure. In this case, it is recommended to use the smallest volume possible and not exceed the final concentration of 0.005 N.
The prepared EPS solution can be stored at 4 °C (Figure 1c).
Data analysis
Establish an experimental design for EPS production using a submerged culture. Use a central or factorial design and analyze the data using analysis of variance or response surface methodology (RSM) (Cui and Zhang, 2012; Supramani et al., 2019; Yu et al., 2022).
Consider that the production of EPS is affected by several variables, such as inoculum properties, time, and culture conditions (Hamidi et al., 2022).
Use a negative control of the culture medium (without inoculum) for later use in the design and for calculating productive parameters.
Carbon sources are among the most crucial variables for EPS production. It is important to utilize at least two different carbon sources when producing EPS, as the production can vary significantly. Glucose or sucrose are proposed in this protocol; however, other conventional and unconventional sources can be used. The concentration of the carbon source varies among different fungal species. The use of 40 g/L is proposed in this protocol; however, other authors reported values up to 120 g/L (Mahapatra and Banerjee, 2013).
During the purification process, filtration is crucial for separating biomass residues. EPS can be used to test cell lines or other biological activity assays.
Calculate the dry EPS, dry biomass, volumetric productivity, specific productivity, practical yield, and recovery efficiency (Fariña et al., 1998; Q. Meng et al., 2021) (Table 2).
Table 2. Fermentation parameters corresponding to the exopolysaccharides (EPS) production using submerged culture
Parameters Equation
Dry EPS (g/L)
Dry biomass (g/L)
Volumetric productivity (VP) Dry EPS (g/L)/Fermentation time (h)
Specific productivity VP (g/L/h)/Dry biomass (g/L)
Practical yield Dry EPS (g/L/)/Initial carbon source substrate (g/L)
Recovery efficiency Lyophilized EPS (g/L)/Dry EPS (g/L)
Analyze the data using RStudio or MATLAB. The data correspond to the determination of the equations in Table 2.
Determine the purity of the EPS by quantifying total polysaccharides, phenols, and proteins. The methodologies for these determinations are available in Parimelazhagan (2016).
Validation of protocol
Results for dry EPS are presented in Table 3. The results are reported as mean ± standard deviation, n = 3 for this work.
Table 3. Comparison of dry exopolysaccharides (EPS) corresponding to its production by macrofungi strains using submerged culture
Macrofungi Reference Condition culture Dry EPS (g/L)
Schizophyllum radiatum Fr This work pH 5.5, SYAMI medium, 40 g/L of sucrose, 7 days, 150 rpm 10.58 ± 3.28
Lentinus crinitus (L.) Fr This work pH 5.5, SYAMI medium, 7 days, 40 g/L sucrose, 150 rpm 4.61 ± 1.30
Schizophyllum radiatum Fr Meng et al., 2021 pH-, seed medium, 6 days, 40 g/L of glucose, 1.0 g/L of Tween 80 3.77
Lentinus tigrinus (Bull.) Fr. He et al., 2016 Lactose 60 g/L, yeast extract 4 g/L; pH = 5.0, 12 days, 0.2 % Tween 80. 4.10 ± 0.27
General notes and troubleshooting
The experimental procedures necessary to carry out the protocol involve the use of chemical substances with a low risk of ignition and irritation by contact with the skin, eyes, and mucous membranes, so all protection elements and safety measures must be used to avoid and prevent accidents.
Acknowledgments
This study was supported by Project No. 58653, Alianza Colombia Científica, Gobierno Nacional de Colombia y Banco Mundial, Proyecto Betaglucanes, Contrato FP448442-211-2018 of 2018. This protocol was adapted from the work carried out by Fariña et al. (2001). We thank Dr. Julia Fariña, PROIMI, CABBIO and MINCIENCIAS for the course entitled: “Mycoprospecting: from nature to biotechnological product. Production in bioreactors and downstream processing.”
Competing interests
The authors declare that they have no conflicts of interest.
Ethical considerations
The specimens used in this study were collected under a permit conceded for access to biological resources for non-commercial purposes (Permiso Marco de Recolección, Resolución 2191 de 2018, Universidad del Tolima) and contract acknowledged for access to genetic resources and derivate products for non-commercial purposes (Contrato Marco de Acceso a Recursos Genéticos y sus Productos Derivados No. 294 de 2020. Expediente RGE0353-3, Universidad del Tolima).
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4,842 | https://bio-protocol.org/en/bpdetail?id=4842&type=0 | # Bio-Protocol Content
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In vitro Quality Assessments of Perfluorocarbon Nanoemulsions for Near-infrared Fluorescence Imaging of Inflammation in Preclinical Models
JJ Jelena M. Janjic
RM Rebecca McCallin
LL Lu Liu
CC Caitlin Crelli
AD Amit Chandra Das
AT Anneliese Troidle
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4842 Views: 559
Reviewed by: Alessandro Didonna Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal of Neuroinflammation Dec 2021
Abstract
Tracking macrophages by non-invasive molecular imaging can provide useful insights into the immunobiology of inflammatory disorders in preclinical disease models. Perfluorocarbon nanoemulsions (PFC-NEs) have been well documented in their ability to be taken up by macrophages through phagocytosis and serve as 19F magnetic resonance imaging (MRI) tracers of inflammation in vivo and ex vivo. Incorporation of near-infrared fluorescent (NIRF) dyes in PFC-NEs can help monitor the spatiotemporal distribution of macrophages in vivo during inflammatory processes, using NIRF imaging as a complementary methodology to MRI. Here, we discuss in depth how both colloidal and fluorescence stabilities of the PFC-NEs are essential for successful and reliable macrophage tracking in vivo and for their detection in excised tissues ex vivo by NIRF imaging. Furthermore, PFC-NE quality assures NIRF imaging reproducibility and reliability across preclinical studies, providing insights into inflammation progression and therapeutic response. Previous studies focused on assessments of colloidal property changes in response to stress and during storage as a means of quality control. We recently focused on the joint evaluation of both colloidal and fluorescence properties and their relationship to NIRF imaging outcomes. In this protocol, we summarize the key assessments of the fluorescent dye–labeled nanoemulsions, which include long-term particle size distribution monitoring as the measure of colloidal stability and monitoring of the fluorescence signal. Due to its simplicity and reproducibility, our protocols are easy to adopt for researchers to assess the quality of PFC-NEs for in vivo NIRF imaging applications.
Keywords: Near-infrared fluorescence (NIRF) Perfluorocarbon (PFC) Nanoemulsions (NEs) Colloidal stability Quality control In vitro validation Fluorescence stability
Background
Macrophages with their pleiotropic functions, ranging from protective to pathological (Ross et al., 2021), are important therapeutic and diagnostic targets (Ardura et al., 2019). Macrophages are the ultimate link between pain and inflammation. They are major drivers of pathological processes in most inflammatory diseases from rheumatoid arthritis, osteoarthritis, and inflammatory bowel disease (IBD) to several types of cancer. Imaging macrophages in vivo has the potential to open a wide variety of research and clinical opportunities, as this would allow a more accurate assessment of inflammatory conditions and therapeutic response. Perfluorocarbon (PFC) nanoemulsions (PFC-NEs) labeling macrophages in vivo for the purpose of 19F MRI detection of inflammation (Temme et al., 2012) and tracking immune cells (Ahrens and Bulte, 2013) has been extensively studied in a multitude of preclinical models of IBD (Kadayakkara et al., 2010 and 2012), neurological diseases (Zhong et al., 2015), arthritis (Balducci et al., 2012), and organ rejection (Hitchens et al., 2011).
The introduction of fluorescent dyes to PFC-NEs serves two purposes: 1) detecting PFC-NE-labeled cells in ex vivo tissue samples by fluorescence microscopy, and 2) providing a complimentary tracer for near-infrared fluorescence (NIRF) imaging in vivo and ex vivo. There are two distinct methods for fluorescent-dye incorporation into PFC-NEs. The first method requires chemical modification of dyes with fluorous tags (Lim et al., 2020) or their direct conjugation to the PFC oil (e.g., perfluoropolyether, PFPE) (Janjic et al., 2008; Patrick et al., 2013). The second method requires PFC-NEs formulated as “triphasic” NEs, which consist of PFC (fluorous phase) and hydrocarbon oil (synthetic or natural/organic phase) combined into an internal phase and then dispersed into water (aqueous), as the third phase (Janjic et al., 2014). These types of NEs allow for high and stable incorporation of one or more fluorescent dyes (Patel et al., 2013a and 2013b; Nichols et al., 2021).
Our lab has reported multiple examples of triphasic theranostic NEs used to encapsulate and deliver non-steroidal anti-inflammatory drugs (e.g., celecoxib) (Herneisey et al., 2019; Liu et al., 2020) and natural products (resveratrol, curcumin) (Herneisey et al., 2016; Herneisey and Janjic, 2023), rendering them multimodal (19F MR/NIRF) imaging and drug delivery agents or theranostics. When theranostic NEs are used in vivo, NIRF imaging tracks macrophages as they respond to the delivered drug by monitoring changes in their infiltration patterns at the site of inflammation caused by injury, surgery, or other insults. Specifically, we demonstrated that COX-2-inhibiting theranostic PFC-NEs have both anti-inflammatory and analgesic effects in vivo in rodent models of neuropathic (Janjic et al., 2018; Saleem et al., 2019) and inflammatory pain (Liu et al., 2020). NIRF imaging was used in these studies to measure macrophage infiltration changes due to the drug delivered. Ex vivo immunofluorescence confirms the theranostic PFC-NE associated with the intended target, where the fluorescent nanoemulsion is co-registered with the specific cell marker (Saleem et al., 2019; Liu et al., 2020). It is critical that PFC-NEs retain their droplet size distribution and fluorescence signal from the time of manufacture, during storage, and during use in animal studies. Therefore, it is important to regularly test the prepared formulations for both stability and any batch-to-batch variations. Any differences in fluorescence signal intensity between PFC-NE batches and in the presence of the drug could lead to ambiguous or inconclusive in vivo imaging results. We developed a series of tests that serve as quality control assessments for PFC-NEs as NIRF imaging agents. Figure 1 shows the conceptual framework for the presented protocols and how they are utilized in validating PFC-NEs ex vivo for the purpose of in vivo macrophage tracking in preclinical models, followed by ex vivo immunofluorescence in tissues of interest. Based on our prior work and by utilizing quality-by-design (QbD) methodologies, we established critical quality attributes (CQAs) that, when met, render PFC-NEs effective in vitro and in vivo as NIRF imaging agents for macrophage tracking (Herneisey et al., 2019; Saleem et al., 2019; Liu et al., 2020; Nichols et al., 2021; Herneisey and Janjic, 2023). In the current protocol, we describe stress tests and associated instrumental measurements that can be easily performed to obtain reliable results on colloidal and fluorescence stability of PFC-NEs under varied conditions: filtration, storage at physiological temperatures, and centrifugation. These conditions model the types of stressors the PFC-NEs are subjected to during their use as either ex vivo cell-labeling agents or in vivo drug delivery and imaging agents. Observed changes in PFC-NE size distribution and fluorescence during the stress tests must fall within the CQA specifications for the formulation to be effective for in vitro and in vivo biological testing as a NIRF imaging agent. In this protocol, we provide a list of quality tests and describe their methodology and associated CQAs for NIRF-labeled PFC-NEs.
Figure 1. Role of quality assessments of fluorescently labeled perfluorocarbon nanoemulsions (PFC-NEs) in preclinical imaging of inflammation in rodent models. The PFC-NEs are tested for size distribution, colloidal and fluorescence stability (lower left panels), and subjected to cell culture testing to measure macrophage uptake and viability (data not shown). Following PFC-NEs tail vein injection in rodents, monocyte-derived macrophages uptake the PFC-NEs and carry them to the site of inflammation. As PFC-NE-labeled macrophages accumulate in the region of inflammation, the near-infrared fluorescent (NIRF) signal also increases. This can be monitored in real time using preclinical imagers, either in vivo or ex vivo. After the experiment, excised tissues can be examined using fluorescence microscopy for the presence of PFC-NE-labeled macrophages. NIRF rat image (top right) reproduced from (Vasudeva et al., 2014); macrophages’ immunofluorescence image from diabetic mice (bottom right) reproduced from (Nichols et al., 2021). Both images are reproduced under the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by-nc/4.0/).
Materials and reagents
Dulbecco’s modified Eagle’s medium (DMEM) (Corning, catalog number: 10-013-CV), store at 4 °C
Fetal bovine serum (FBS) (ATCC, catalog number: 30-2020TM), aliquot and store at -20 °C
Freshly prepared, 100–200 nm sized nanoemulsions (please check references for the method of preparation of nanoemulsions)
24-well plates (Corning, VWR, catalog number: 10062-896)
96-well plates (Corning, VWR, catalog number: 29442-056)
Microcentrifuge tubes (Fisherbrand Premium Microcentrifuge Tubes, catalog number: 05-408-129)
MilliporeSigmaTM MillexTM-GS Sterile Syringe Filter Unit, MCE, 0.22 and 0.45 μm (catalog numbers: SLGSR33SB and SLHA033SS)
FisherbrandTM disposable cuvettes (Fisher Scientific, catalog number: 14-955-127)
Parafilm (Sigma-Aldrich, P7793)
Type I ultrapure HPLC grade water (18.2 MΩ·cm) generated using PURELAB® Option-Q system
DiR [1,1’-Dioctadecyl-3,3,3’,3’-Tetramethylindotricarbocyanine Iodide, DiIC18(7)] (Thermo Scientific, catalog number: D12731)
DiD [DiIC18(5); 1,1’-dioctadecyl-3,3,3’,3’- tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate salt] (Thermo Scientific, catalog number: D7757)
Fluid thioglycolate medium (Millipore, catalog number: STBMCTM12)
Trypticase soy broth (Millipore, catalog number: STBMTSB12)
Equipment
Odyssey Classic Imager (Li-COR Inc., Lincoln, NE)
Odyssey M Imaging System (Li-COR Inc., Lincoln, NE)
Pearl Small Animal Imaging System (Li-COR Inc., Lincoln, NE)
37 °C incubator (Fisher Scientific, IsoTemp® 500 Series)
Zetasizer NanoZS (Malvern, UK)
Prism R Refrigerated Microcentrifuge (Labnet International Inc.)
Software and datasets
Image Studio Ver 5.2 (Li-COR Biosciences, U.S.)
Zetasizer Software 7.13 (Malvern Panalytical)
GraphPad Prism, 10.0.0
Procedure
Defined quality control attributes (CQAs) and quality control (QC) measures listed in Table 1 encompass routine assessments of the manufactured nanoemulsions to ensure colloidal stability over time after refrigerated (4 °C) storage. We present procedures adapted from Herneisey and Janjic (2023). Nanoemulsion droplet size, polydispersity index (PDI), and fluorescence signal are measured throughout the lifetime of the nanoemulsion. At the same time, the other listed stress tests can be done within the first couple of weeks after manufacturing. All colloidal characterization and stress tests are elaborately described throughout this manuscript. Our lab has utilized this battery of tests in numerous publications for nanoemulsion quality assurance. CQAs and QC for our formulations are listed in Table 1, which was adapted from Herneisey and Janjic (2023). Figure 2 summarizes the representative results of CQAs and QC obtained after manufacturing two-color fluorescently labeled PFC-NEs.
Table 1. Summary of diameter and polydispersity index (PDI) critical quality attribute (CQA) and quality control (QC) testing performed on perfluorocarbon nanoemulsions (PFC-NEs).Reproduced from Herneisey and Janjic (2023), Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by-nc/4.0/)
Description CQA or QC Diameter specification PDI Specification
Initial diameter CQA 140–180 nm N/A
Initial PDI CQA N/A < 0.25
% Diameter change or PDI value after:
Filtration (0.22 μm) CQA < ± 5% < 0.25
Cell culture conditions CQA 0–10% < 0.3
Storage at 4 °C for 95 days CQA < ± 5% < 0.25
Storage at 4 °C for 215 days QC N/A N/A
Incubation at 80 °C for 7 days QC N/A N/A
Figure 2. In vitro stability testing of near-infrared fluorescent (NIRF)-labeled perfluorocarbon nanoemulsions (PFC-NEs). A) Droplet-size-distribution overlay comparison between day 1 and 8 months after manufacturing. B) pH stability over 21 days. C) PFC-NE serum stability in three different biological media (Water, DMEM, and 20% FBS in DMEM) was measured at time 0 and after 72 h incubation at 37 °C (p = 0.514, p = 0.777, p = 0.969, respectively, Student’s t-test, n = 3). D) PFC-NE centrifugation stability. PFC-NE (undiluted) size was measured before and after centrifugation (p = 0.097, Student’s t-test, for n = 3). E) PFC-NE stability before and after sterile filtration with a 0.22 μm membrane filter (p = 0.642, Student’s t-test, n = 3). F) Fluorescence signals of the PFC-NE with the NIRF dye DiR were compared between week 1 and week 20 after manufacturing. PFC-NE was diluted at a PFC-NE to water ratio of 1:4. Li-COR Odyssey Imager was used for collecting the fluorescence signal. GraphPad Prism 9.0 used for statistical analysis; data represents an average ± SD with at least three replicates per measurement. Data reproduced from Nichols et al. (2021) under the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by-nc/4.0/).
Dynamic light scattering (DLS) measurement of size and zeta potential
This analytical technique is used to measure particle size (nm), polydispersity index (PDI), and zeta potential (mV). We have utilized this method in numerous publications to describe and characterize our nanoemulsions (NEs). Our method begins with a 60 s equilibrium step for the DLS measurements, followed by three rounds of typically 15 runs at 25 °C. A number of runs needed for the optimal size distribution measurement is automatically selected by the Zetasizer Software 7.13 based on particle counts during each run. Variations in size and PDI over time or after stress indicate destabilization of colloidal infrastructure. Size measurements using Malvern’s Zetasizer provide size distribution curves that can indicate particle size uniformity within a dilution. Samples are prepared as dilutions in a medium. New nanoemulsion formulations are diluted 1:20 v/v to 1:160 v/v to determine the dilution that achieves the highest intensity size distribution. The nanoemulsions are diluted at either 1:40 v/v or 1:80 v/v. Once selected, the same dilution ratio (v/v) should be used in all tests requiring size, PDI, or zeta potential measurement. Standard size, PDI, and zeta potential measurements are diluted in HPLC (Type 1 water). Dilution medium is subject to change, pending the type of test intended to be run. The presented methods detail the simultaneous measurement of size and zeta potential by creating a dilution at a 2 mL volume of nanoemulsion in water. However, a 1 mL dilution can be prepared if only size or PDI is measured. An example data is shown in Figure 2A.
Method:
Dilute nanoemulsion in HPLC grade water 1:40 v/v or 1:80 v/v to create a total volume of 2 mL in a plastic cuvette compatible with Zetasizer Nano.
Insert the cuvette into the Zetasizer and measure size and PDI.
After each completed measurement, remove the cuvette and flush 1 mL of nanoemulsion dilution through a zeta potential measurement cuvette to wet the electrodes.
Fill the zeta cuvette with the remaining dilution to the zeta cuvette fill line.
Place the zeta cuvette into the Zetasizer Nano and measure the zeta potential.
Filtration test
For nanoemulsions intended for in vivo use, it is important to have a method of sterilization that does not affect the colloidal stability of the nano-formulation. Within the confines of a sterile hood, nanoemulsions are filtered through a 0.22 or 0.45 μm syringe filter. Filter size depends on the size of the particles to be filtered. For example, it is difficult to filter particles of ~200 nm and above using a 0.22 μm filter because particles may not easily flow through the filter, and the stress of forcing particles through a small filter pore may affect colloidal structure and stability. In these cases, a larger filter size can be used, such as 0.45 μm. Most of our nanoemulsions with a particle size <140 nm and PDI <0.15 can be filtered using 0.22 μm filters. Nanoemulsions falling outside this size and PDI range must be filtered using 0.45 μm pore size filters. See Figure 2A for typical measurements of a filtration test. Example data are shown in Figure 2E.
Method:
Measure size and PDI of unfiltered nanoemulsion in HPLC grade water.
Filter a sample of the nanoemulsion within a sterile hood using a 0.22 or 0.45 μm filter.
Using the same dilution ratio (v/v) as in step 1 (Method A), measure the size and PDI of the filtered nanoemulsion.
Compare the size values before and after filtration to see if there is a significant change in the size value.
Note: Size variation of ±10% is within the acceptable range (for details, see references: Janjic et al., 2018; Herneisey et al., 2019; Saleem et al., 2019; Liu et al., 2020; Nichols et al., 2021; Herneisey and Janjic, 2023).
Laboratory grade sterility testing before use in animal models
PFC-NEs are manufactured non-sterile and are sterile-filtered through either a 0.22 or 0.45 μm syringe filter, depending on nanoemulsion droplet size. To validate nanoemulsions for use in animal studies, we utilize a sterility protocol adapted from the USP 34 microbiological tests, article 71 [(71) Sterility Tests, 2016]. It utilizes changes in pH before and after incubation in culture media to determine sterility. Statistically significant pH deviations are determined by a t-test calculated using GraphPad Software. A significant pH change indicates that the nanoemulsion is not sterile. Selected media offer desirable growth conditions for biological contaminants such as bacteria and fungi. Fluid thioglycolate medium is used to detect only anaerobic bacteria, and trypticase soy broth is used to detect aerobic and anaerobic bacteria and fungi.
Method:
Dissolve nanoemulsions into the following media at 1:40 v/v ratios (0.5 mL of nanoemulsion to 19.5 mL of medium).
Fluid thioglycolate medium: in a water bath, 30–35 °C.
Trypticase soy broth: incubated at 22.5 ± 2.5 °C.
Divide the dilutions into two portions, one for initial pH measurement and the other portion for incubation over time.
Take a photograph of the appearance of the sample in the media before incubation begins. This will act as the medium control.
Measure pH for both medium control and sample dilutions.
Seal all the sample tubes and incubate the nanoemulsions in the media solutions for 14–28 days.
Visually assess media for microorganism growth (turbidity), photograph again and compare to photograph from step 3, and then measure pH again.
Centrifugation test
To assess colloidal stability of nanoemulsions in physically adverse conditions, such as emulating rough shipping conditions and aggressive handling, centrifugation is used. Nanoemulsion colloidal resilience to centrifugation is also important because we utilize centrifugation to measure a formulation’s drug content and separate nanoemulsion treatments from cells during in vitro cell culture assays (Herneisey et al., 2019). In our experience, centrifugation at 3,000 rpm (840× g) for 30 min has not led to significant size changes. An example of centrifugation test results is shown in Figure 2D, showing differences in droplet size before and after centrifugation.
Method:
Isolate 500 μL of undiluted and filtered nanoemulsion into 1.5 mL microcentrifuge tubes.
Centrifuge both tubes at 1,100 rpm (113× g) for 5 min at ambient temperature (25 °C).
Pipette out 100 μL of the samples and measure their size and PDI in HPLC grade water.
Centrifuge the remaining 400 μL of sample in the tubes at 3,000 rpm (840× g) for 5 min at 25 °C.
Remove enough of the sample from each tube to prepare a dilution in HPLC grade water for size and PDI measurement (Nichols et al., 2021).
Compare the sizes of all the samples (non-centrifuged, filtered samples serve as controls).
Note: The centrifugation step should not lead to significant size change for stable nanoemulsions, with any change being typically less than 10%.
Serum stability
Colloidal and fluorescence stability needs to be carefully evaluated to avoid formulation failure in vivo. While it is easy to convincingly show how the formulation performs under in vitro conditions, it is extremely challenging to predict in vivo behavior of nanoemulsions once they come into contact with complex biological media. Such media contains various components such as lipids, proteins, electrolytes, etc., all of which can determine the fate of the nanoemulsion, impacting biodistribution, toxicity, and the pharmacokinetic profile. The serum stability test is crucial if the nanoemulsions are intended for biomedical testing in animals (Moore et al., 2015). An example of serum stability test results is shown in Figure 2C, showing differences in droplet size before and after incubation for 72 h in serum-containing media at body temperature.
Method:
Prepare triplicate 1.0 mL dilutions of nanoemulsion in 1.5 mL microcentrifuge tubes with the following:
HPLC grade water
DMEM
20% FBS in DMEM
Measure size of the first set of dilution using DLS. This is the control, 0-hour time point reading.
Incubate the remaining two dilutions at 37 °C.
Measure size of the second dilution after 24 h and the size of the third dilution after 72 h. There is no need to further dilute the samples prior to DLS measurement because the nanoemulsion is diluted enough during initial preparation in step 1.
Compare any size differences between the measurements.
Note: An alternate version of the serum stability test can be carried out in HPLC grade water, 1× PBS, DMEM medium, 10% FBS in DMEM, and 20% FBS in DMEM and size measurements followed up after 24 h, 48 h, and 72 h.
Thermal cycling
Thermal cycling studies are performed to study the temperature sensitivity/tolerance of nanoemulsions and to accelerate the aging of the formulation. This test can also help determine the appropriate storage conditions and predict shelf life for the prepared nanoemulsions (Herneisey et al., 2019).
Method:
Aliquot 5.0 mL of undiluted, filtered nanoemulsion to a 15 mL vial.
Seal the vials using parafilm and store them at 4 °C.
After a 24 h incubation at 4 °C, transfer the vials to an incubator set at 50 °C and incubate for another 24 h.
Cycle the vials between 4 °C and 50 °C for four thermal cycles per temperature. Each thermal cycle is 24 h, total thermal cycling time is eight days.
After the cycles are completed, allow the samples to equilibrate to room temperature for 1 h and then measure the size and PDI.
Measurements of freshly prepared filtered nanoemulsions continuously stored at ambient temperature are used as control.
Fluorescence stability
To determine the long-term fluorescence stability of NIRF-labeled nanoemulsions, it is important to quantify the fluorescence at regular intervals to check for the loss of fluorescence. Our lab protocols were established using Li-COR Odyssey Classic Imager (Nichols et al., 2021) but can be easily adapted for other fluorescent plate readers (Herneisey and Janjic, 2023). Imager focus and intensity settings are adjusted to obtain the highest fluorescence signal without oversaturating and bleaching samples. Multiple measurements are obtained to determine the optimal setting selection. The Odyssey M Imaging system can also obtain fluorescence measurements using focus offset. Focus offset is determined the same way as on the Odyssey Classic. Once initial optimal settings are determined, the same settings are used for all follow-up measurements. We assess initial fluorescence in vitro to assure that the nanoemulsion signal is high enough to be detected in vivo during animal studies. Example of in vitro fluorescence stability monitoring is shown in Figure 2F, where serial dilutions of nanoemulsions are imaged and fluorescence quantified with Image Studio Ver 5.2. Figure 3 shows example data for monitoring inflammation in vivo by measuring fluorescence in mice on the Li-COR Pearl imager up to 40 days following tail vein injection of NIRF-labeled PFC-NEs. Fluorescence differences correspond to changes in macrophage infiltration patterns in response to the drug celecoxib (CXB) delivered in a drug-loaded NIRF-labeled PFC-NE (CXB-NE) (Liu et al., 2020).
Method:
HPLC grade water was used to prepare six serial dilutions of nanoemulsions starting from 1:5 v/v (200 μL of nanoemulsion with 800 μL of water) through 1:160 v/v, in a 24-well plate.
Three replicates of 100 μL from each dilution were transferred to a clear, flat-bottomed 96-well plate and the plate was read on the Li-COR Odyssey Imager.
The imaging parameters such as focus offset (e.g., 0.5–1.0) and intensity (e.g., 0.5) are adjusted to each plate/nanoemulsion combination used for the freshly prepared nanoemulsions and are kept constant for all follow up measurements.
DiR [1,1′-Dioctadecyl-3,3,3′,3′-Tetramethylindotricarbocyanine Iodide, DiIC18(7)] is our most preferred near-infrared dye, which can be detected using an 800 nm fluorescent channel.
Combinations of dyes, such as DiR (800 nm) and DiD (700 nm) [DiIC18(5); 1,1′-dioctadecyl-3,3,3′,3′- tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate salt], can be incorporated and measured in a single formulation if the dyes have different detection wavelengths (nm).
Repeat the fluorescence measurements on D0 (Day 0), D7, D15, D30, D60, D90, and so on as per the study requirements to check for fluorescence stability. The fluorescence loss should not be greater than 10% of the initial value (Herneisey and Janjic, 2023).
Note: Our lab has shown stable nanoemulsion fluorescence for a maximum period of 215 days, in vitro (Herneisey and Janjic, 2023) and that PFC-NEs can be detected in excised tissues after a period of 40 days following their in vivo administration (Liu et al., 2020).
Figure 3. Long-term in vivo fluorescence imaging of PFC-NE–labeled macrophages in a mouse inflammation model. (A) Drug-free (DF-NE) and (C) Celecoxib (CXB-NE) nanoemulsions showing stable fluorescence in vivo for 40 days in a male mouse model; (B, D) NIRF imaging of female mice treated with same DF-NE and CXB-NE nanoemulsions, respectively. (E) DF-NE and CXB-NE associated signal distribution in males for 40 days of follow up. CXB-NE treated animals show a decreasing signal over time, though not statistically different from DF-NE treatments. Data represents average ± SEM; two-way ANOVA was used to establish statistical significance between all groups. Reproduced from Liu et al. (2020) under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by-nc/4.0/).
Data analysis
The differences in size must be within the range specified in Table 1 and according to the CQA and QC parameters determined by the experimenter.
In vivo fluorescence data must be analyzed using two-way ANOVA, as shown in Figure 4. Statistical analysis and graphing were performed using GraphPad Prism 10.0.0 software. The values are represented as average ± SEM. Consider a P value of < 0.05 as statistically significant.
Figure 4. Laboratory grade sterility testing for perfluorocarbon nanoemulsions (PFC-NEs). Fluctuations of pH before and after 28 days of incubation in A) trypticase soy broth and B) fluid thioglycolate medium. A t-test was used to determine statistical significance of pH change over time. Nanoemulsions passed sterility qualifications with non-significant pH changes and lack of turbidity in culture media.
Validation of protocol
M. Herneisey and J.M. Janjic* “Multiple Linear Regression Predictive Modeling of Colloidal and Fluorescence Stability of Theranostic Perfluorocarbon Nanoemulsions” Pharmaceutics 2023, 15 (4), 1103.
J.M. Nichols, C.V. Crelli, L. Liu, H.V. Pham, J.M. Janjic*, A.J. Shepherd* “Tracking macrophages in diabetic neuropathy with two-color nanoemulsions for near-infrared fluorescent imaging and microscopy” Journal of Neuroinflammation (2021), Vo. 18, Article number: 299.
L. Liu, H. Karagoz, M. Herneisey, F. Zor, T. Komatsu, S. Loftus, B.M. Janjic, V.S. Gorantla and J.M. Janjic “Sex Differences Revealed in a Mouse CFA Inflammation Model with Macrophage Targeted Nanotheranostics” Theranostics 2020, 10 (4), 1694.
M. Herneisey, L. Liu, E. Lambert, N. Schmitz, S. Loftus, and J.M. Janjic* Development of Theranostic Perfluorocarbon Nanoemulsions as a Model Non-Opioid Pain Nanomedicine Using a Quality by Design (QbD) Approach AAPS PharmSciTech 2019, 20: 65.
Acknowledgments
The presented protocols and work have been supported in part by CDMRP awards: W81XWH-20-1-0854, W81XWH-20-1-0276, W81XWH-19-1-0828 and W81XWH-20-1-0730.
Composite schematic image Created with BioRender.com.
Competing interests
The authors declare no competing interests.
References
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Herneisey, M., Liu, L., Lambert, E., Schmitz, N., Loftus, S. and Janjic, J. M. (2019). Development of Theranostic Perfluorocarbon Nanoemulsions as a Model Non-Opioid Pain Nanomedicine Using a Quality by Design (QbD) Approach. AAPS PharmSciTech 20(2): e1208/s12249-018-1287-6.
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Janjic, J. M., Srinivas, M., Kadayakkara, D. K. K. and Ahrens, E. T. (2008). Self-delivering Nanoemulsions for Dual Fluorine-19 MRI and Fluorescence Detection. J. Am. Chem. Soc. 130(9): 2832–2841.
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Isolation and Culture of Neural Stem/Progenitor Cells from the Hippocampal Dentate Gyrus of Young Adult and Aged Rats
MA Mina Afhami
MB Morteza Behnam-Rassouli
AG Ali Gorji
SK Saeed Karima
KS Koorosh Shahpasand
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4843 Views: 646
Reviewed by: Vivien J. Coulson-ThomasMunenori Ishibashi Anonymous reviewer(s)
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Abstract
Adult neural stem/progenitor cells (NSPCs) in two neurogenic areas of the brain, the dentate gyrus and the subventricular zone, are major players in adult neurogenesis. Addressing specific questions regarding NSPCs outside of their niche entails in vitro studies through isolation and culture of these cells. As there is heterogeneity in their morphology, proliferation, and differentiation capacity between these two neurogenic areas, NSPCs should be isolated from each area through specific procedures and media. Identifying region-specific NPSCs provides an accurate pathway for assessing the effects of extrinsic factors and drugs on these cells and investigating the mechanisms of neurogenesis in both healthy and pathologic conditions. A great number of isolation and expansion techniques for NSPCs have been reported. The growth and expansion of NSPCs obtained from the dentate gyrus of aged rats are generally difficult. There are relatively limited data and protocols about NSPCs isolation and their culture from aged rats. Our approach is an efficient and reliable strategy to isolate and expand NSPCs obtained from young adult and aged rats. NSPCs isolated by this method maintain their self-renewal and multipotency.
Key features
• NSPCs isolated from the hippocampal dentate gyrus of young adult and aged rats, based on Kempermann et al. (2014) and Aligholi et al. (2014).
• Maintenance of NSPCs isolated from the dentate gyrus of aged rats (20–24 months) in our culture condition is feasible.
• According to our protocol, maximum growth of primary neurospheres obtained from isolated NSPCs of young and aged rats took 15 and 35 days, respectively.
Graphical overview
Isolation and expansion of neural stem/progenitor cells
Keywords: Aging Neural stem/progenitor cells Dentate gyrus Neurogenesis Hippocampus Proliferation
Background
Neural stem/progenitor cells (NSPCs) in neurogenic areas of the adult mammalian brain, i.e., the subventricular zone (SVZ) of the lateral ventricle and the subgranular zone of the hippocampal dentate gyrus (DG), divide symmetrically and asymmetrically to preserve their lineage (Walker and Kempermann, 2014; Gage, 2019). In a symmetric division, the number of progenitor cells increases, as two identical daughter cells originate from a progenitor cell or stem cell. The asymmetric division of progenitor cells produces two distinct types of cells. For example, one radial-glial cell gives rise to one radial-glial cell and one neuron (Zhao and Moore, 2018). Neurogenesis in the dentate gyrus contributes to learning, memory, and mood regulation, which decline during aging. Isolation of NSPCs from rat dentate gyrus provides valuable adult-cell-based models for identifying cellular and molecular mechanisms of neurogenesis in the hippocampus and to assess the effects of extrinsic factors and drugs on their functions outside their niches. There are different intrinsic regulatory mechanisms for NSPCs in the DG and SVZ (Takei, 2019). With increasing age, a repertoire of intrinsic and extrinsic factors drives most of the NSPCs to enter the senescent stage, limiting their proliferative capacity, and leading to subsequent dysfunctions (Obernier and Alvarez-Buylla, 2019). Slow proliferation and declining differentiation capacity are features of NSPCs in aging, which make finding strategies for improvement of neurogenesis during aging important (Kase et al., 2020).
The study of NSPCs function and their responses to aging, regeneration purposes, and molecular and cellular mechanisms of neurogenesis in various cell-culture-based models of different neurological disorders, including neurodegenerative and neurovascular diseases, requires isolation and culture of NSPCs from adult neurogenic niches (Akers et al., 2014; Christian et al., 2014; Bond et al., 2015). Among various approaches, neurosphere culture provides a 3D environment-like niche of neural stem cells, increases cell division, and offers conditions permissible for more passage numbers (Azari et al., 2010; Guo et al., 2012; Aligholi et al., 2014). However, recognition of cell morphology is difficult in the neurosphere culture system. Cells in the center of neurospheres may differentiate slowly with low survival and yield (Sun et al., 2011). In monolayer cell culture, cells have easy access to nutrients and their morphology could be easily detected. However, cells in this method might differentiate spontaneously, and self-renewal ability declined after more passages (Reynolds and Rietze, 2005; Ray and Gage, 2006).
As the proliferation and differentiation potential of the hippocampal DG differs from the SVZ, developing a complete protocol to obtain high-yield NSPCs from rat DG is needed. There is controversy regarding hippocampus-derived stem cells from adult rats and whether they have unlimited self-renewal or limited proliferative and differentiating potential (Gobbel et al., 2003; Chen et al., 2007). In this method, the expansion of isolated cells from aged rats has been introduced and discussed.
Using this isolation and culture method, we produced a robust and reproducible number of DG neurospheres from young adult and aged rats after 14–18 days and 25–30 days, respectively. In our method, 1.5 × 105 neural progenitor cells yield 120 spheres of 200–250 µm for young adult rats, and 7 × 104 neural progenitor cells yield 90 spheres of 100 µm for aged rats on average. An appropriate approach providing optimal regional-specific NSPCs could improve utilizing cells in a clinical setting for the treatment of neurological disorders as well as boost experimental aging research.
Materials and reagents
Animals
Animal housing was performed in accordance with the Guidelines of the Shefa Neuroscience Research Center, Tehran, Iran. Young adult male Wistar rats between 2 and 3 months of age (180–230 g) and aged rats between 20 and 24 months (400–450 g) were used.
General materials
Ketamine (alfasan, catalog number: 36408/3000)
Xylazine (alfasan, catalog number: 36408/3007)
Glass Petri dish (Nest Biotechnology, catalog number: 704001)
Phosphate buffer saline (PBS) (tablet) (Thermo Fisher Scientific, GibcoTM, catalog number: 18912014)
0.2, 0.5, and 1.5 mL Eppendorf tubes (Nest Biotechnology, catalog number: 613111)
6, 12, 24, 48, and 96 adherent well plates (treated) (JetBiofil, catalog number: TCP011096, TCP011012, TCP011048)
Non-adherent well plates: 6 wells (Falcon, catalog number: 353046) and 12 wells (Falcon, catalog number: 351143)
Adherent T-25 flask (JetBiofil, catalog number: TCF012050)
Non-adherent T-25 flask (JetBiofil, catalog number: TCF012600)
0.2 μm filter (PES membrane) (BioFil, catalog number: FMC 2011018)
Parafilm (Sigma, catalog number: P7793)
96% ethanol (Merck, catalog number: 100971)
70 μm cell strainer (BD Falcon, catalog number: 352340)
Petri dish culture (JetBiofil, Germany, MCD000035)
15 and 50 mL Falcon tubes (JetBiofil, catalog number: CFT011500)
Pipette tips blue 1,000 μL (Universal, catalog number: 2451-1K0), yellow 100 μL (Universal, catalog number: 200-B-YL), and 10 μL (Labcon Eclipse, catalog number: 490007-952)
Reagents
0.05% Trypsin/EDTA (Biowest, catalog number: L0931-100)
Digestive enzymes and materials:
Papain (Roche, catalog number: 108014)
Dispase (Gibco, catalog number: 17105-041)
DNase (Roche, catalog number: 11284932001)
Accutase (Sigma, catalog number: A6964)
L-Cysteine (Sigma, catalog number: 52904)
EDTA (Sigma, catalog number: 17892)
Hanks’ balanced salt solution (HBSS) (Thermo Fisher Scientific, GibcoTM, catalog number: 24020117)
Fetal bovine serum (FBS) (Thermo Fisher Scientific, GibcoTM, catalog number: 10270106)
Trypan blue solution 0.4% (Sigma-Aldrich, catalog number: T8154)
4% Paraformaldehyde (powder) (Sigma-Aldrich, catalog number: P6148)
Triton X-100 (Sigma-Aldrich, catalog number: T8787)
Bovine serum albumin (BSA) (powder) (Sigma-Aldrich, catalog number: A2058)
Normal goat serum (Thermo Fisher Scientific, catalog number: 50197Z)
Mouse anti-Nestin (Merck Millipore, catalog number: MAB 353)
Rabbit anti-Sox2 (Santa Cruz, catalog number: SC20088)
Mouse anti-GFAP (Sigma, catalog number: G3893)
Goat anti-rabbit IgG (Abcam, catalog number: ab9717)
Goat anti-mouse (Abcam, catalog number: ab6785)
4’,6-Diamidino-2-phenylindole (DAPI) (Thermo Fisher Scientific, catalog number: 62248)
Coating
Laminin (Sigma, catalog number: L2020)
Poly-L-ornithine solution (Sigma, catalog number: P4957)
Culture medium (see Recipes)
Dulbecco’s modified Eagle’s medium (DMEM)/F12 (Gibco, catalog number: 11320033)
N2 supplement (Gibco, catalog number: 17501), used for serum-free culture condition of neuroblastoma and primary cells of the central nervous systems. It is composed of vitamins, hormones, insulin, and transferrin. These components maintain neural stem cells in an undifferentiated state and support the proliferation of cells in a serum-free culture
B27 supplement (Gibco, catalog number: 12513-010), a supplement as a cocktail of antioxidants, vitamins, and proteins to reduce reactive oxygen damage and for the long-term survival of neural stem cells. It is a widely used supplement for serum-free culture conditions. Therefore, this supplement contains defined components necessary for maintaining the health of the culture and reducing variability
Heparin (Sigma, catalog number: H3149)
Penicillin/Streptomycin (Pen/Strep) (Sigma, catalog number: P4333)
Amphotericin B (Biowest, catalog number: L0009-050)
Glutamax (Gibco, catalog number: 35050-061)
Basic fibroblast growth factor (bFGF) (Sigma, catalog number: F0291)
Epidermal growth factor (EGF) (Sigma, catalog number: E4127)
Solutions
70% ethanol (see Recipes)
Digestive solution (sterile, 0.2 μm filtered) (see Recipes)
Blocking solution (see Recipes)
Antibody solution (see Recipes)
Growth medium (culture medium) (see Recipes)
Coating solution (sterile, 0.2 μm filtered) (see Recipes)
Dissection solution (see Recipes)
Ketamine (80 mg/mL) and xylazine (10 mg/mL) working solutions (see Recipes)
PBS (1×) (see Recipes)
Recipes
70% ethanol
Dilute 365 mL of 96% ethanol with 135 mL of distilled water to make up a volume of 500 mL of 70% ethanol.
Digestive solution (sterile, 0.2 μm filtered)
Dissolve 40 μL of Dispase (2 mg/mL), 80 μL of DNase (0.2 mg/mL), 100 μL of Papain (0.5 mg/mL), 1 mg of L-cysteine, and 100 μL of EDTA in 2 mL of DMEM/F12. This volume of solution is appropriate for one rat.
Preparation of papain (0.5 mg/mL): add 100 μL of papain stock solution (10 mg/mL) to 1.95 mL of DMEM/F12 basic medium to prepare 2 mL of 0.5 mg/mL papain working solution.
Preparation of Dispase (2 mg/mL): add 100 μL of Dispase stock solution (10 mg/mL) to 1.9 mL of DMEM/F12 basic medium to prepare 2 mL of 2 mg/mL Dispase working solution.
Preparation of DNase (0.2 mg/mL): add 80 μL of DNase stock solution (5 mg/mL) to 1.96 mL of PBS (1×) to prepare 2 mL of 0.2 mg/mL DNase working solution.
Add 1 mg of L-cysteine and 100 μL of EDTA to the solution.
Blocking solution
Dissolve 10% normal goat serum, 1% BSA, and 0.01% Tween in PBS (1×). To prepare 500 µL of blocking solution, add 50 µL of normal goat serum, 5 µL of BSA, and 0.5 µL of Tween to 444.5 µL of PBS (1×).
Antibody solution
Dilute primary antibodies in blocking solution:
To prepare 350 μL of 1:200 Anti-Nestin solution, add 1.75 μL of Nestin antibody to 348 μL of blocking solution. Prepare before use.
To prepare 350 μL of 1:100 Sox2 solution, add 3.5 μL of Sox2 antibody to 346.5 μL of blocking solution. Prepare before use.
To prepare 250 μL of 1:250 GFAP solution, add 1 μL of GFAP antibody to 249 μL of blocking solution. Prepare the solution before use.
Antibody Stock concentration Dilution range Final concentration (dilution used for staining)
Sox2 200 μg/0.1 mL 1:50–1:500 1:100
Nestin 1.25 μg/mL 1:20–1:200 1:200
GFAP 2.5–5 μg/mL Not available for immunocytochemistry 1:250
Growth medium (culture medium)
DMEM/F12
2% B27 supplement
1% N2 supplement
20 ng/mL bFGF
20 ng/mL EGF
2 μg/mL heparin
1% Penicillin/streptomycin
1% Glutamax
Coating solution (sterile, 0.2 μm filtered)
PLO working solution (20 μg/mL):
Dissolve 2 mL of PLO stock solution (0.1 mg/mL) in 8 mL of PBS (1×) to prepare 10 mL of PLO with the final concentration of 20 μg/mL (store at 4 °C).
Laminin working solution (5 μg/mL)
Dissolve 250 μL of stock solution (1 mg/mL) in 49.75 mL of PBS (1×) to get 50 mL of laminin solution with a final concentration of 5 μg/mL.
Dissection solution
To prepare PBS or HBSS or DMEM/F12 containing 3% Pen/Strep and 1% Amphotericin B: add 0.3 mL of Pen/Strep and 0.1 mL of Amphotericin B to 9.7 mL of PBS (1×).
Ketamine (80 mg/mL) and xylazine (10 mg/mL) working solutions
To prepare 3 mL (concentration of 80 mg/mL) of ketamine working solution:
Add 2.4 mL of ketamine stock solution (100 mg/mL) to 0.6 mL of PBS (1×).
To prepare 3 mL (concentration of 10 mg/mL) of xylazine working solution:
Add 1.5 mL of xylazine stock solution (20 mg/mL) to 1.5 mL of PBS (1×).
PBS (1×)
One tablet of PBS in 500 mL of distilled water used to produce PBS (1×) solution. Then the solution is autoclaved.
Equipment
Laminar hood (Jal Tajhiz, catalog number: 1229)
37 °C incubator shaker (IKA, model: KS 4000i Control)
Centrifuge machine (Hettich Lab Technology, model: Universal 320R)
Hemocytometer (Sigma-Aldrich, catalog number: Z359629)
CO2 cell culture incubator (Memmert, model: INC108 T2T3)
Inverted fluorescence microscope (Optika, model: XDS-2FL)
Small forceps surgical tools (Fine Science Tools, catalog number: 11050-10)
Stereomicroscope (Olympus, catalog number: 7L19549)
Dissection tools
Large scissors (Fishersci, catalog number: 10633652)
Small scissors (Fishersci, catalog number: 15207266)
Forceps (Fishersci, catalog number: 10098140)
Curved forceps (Fishersci, catalog number:10213941)
Angled forceps (Fishersci, catalog number: 22-079-762)
Spatula (Fishersci, catalog number: 11750229)
Autoclave (Kavoosh, catalog number: 2589224)
Oven (Memert, catalog number: 1-100-800)
Rodent guillotine (DCAP, Germany)
Rongerus (Fine Science Tools, catalog number: 16004-16)
Software
Infinity software (version 4.6)
Procedure
A schematic diagram depicting the whole experimental procedure is shown in Figure 1.
Figure 1. Picture demonstrating the brain removal, dentate gyrus (DG) dissection, and DG digestion
Poly-L-ornithine/Laminin coating of cell culture plates layer
Add the appropriate volume of Poly-L-ornithine (PLO) solution (20 μg/mL) into cell culture well plates to completely cover the surface (1.5 mL for 6-well plates, 450 mL for 24-well plates, and 60 μL for 96-well plates) (see Recipes).
Incubate at 37 °C for 2 h.
Tilt the plates and remove the PLO by pipetting at the corner of each well, then add the appropriate volume of PBS (1×) to each well (2 mL for 6-well plates, 1 mL for 24-well plates, and 100 μL for 96-well plates), and incubate for 2 min. Wash and incubate with PBS (1×) three times.
Add laminin so that its concentration in each well is 5 μg/mL, and incubate overnight (16 h) at 37 °C.
Use the plates immediately before letting them dry. It is optional to wash each well with PBS (1×) before seeding cells. Seed the cells 1–2 min after removal of laminin or after washing with PBS (1×).
Harvest rat brain tissue (Figure 2)
Figure 2. Harvest of the adult rat brain. A. Surgical tools. B. A rodent guillotine was used to cut the head. C. 70% alcohol was sprayed onto the head and a skin incision was made along the midline using fine scissors. D. Cutting of the remaining muscle tissues was performed by large scissors. E and F. Removal of occipital bones using rongeur is presented. G. Exposed cerebellum is presented. H. Access to the hemispheres was achieved by cutting the bone throughout the midline. I. Placing the flat blade of the scissors beneath the right parietal plate and pressing against the skull’s inner surface. J. Exposed right hemisphere is shown. K. The same procedure for the left one was performed. L. Two hemispheres are depicted. M and N. Removal of frontal bones using rongeur is shown, starting at the site of the interfrontal suture and continuing to cut small fragments of bones. O. Frontal parts of the brain have been accessed. P and Q. Separation of the brain from the skull was carried out by inserting a round spatula at the edges of the exposed brain and moving laterally in backward and forward motions and detaching the nerves through sliding the spatula beneath the brain. R. Scooping the brain out, it was transferred into a 50 mL Falcon tube containing 3–5 mL of cold PBS (1×) containing 3% Pen/Strep and 1% Amphotericin B kept on icy water.
Use two separate sets of sterile surgical tools to reduce the risk of contamination with hair and blood. One set for harvesting brains from adult rats and another for microdissection of the dentate gyrus from the brain. If the following steps are performed correctly, the brain is isolated from the skull without any damage.
Anesthetize rats using intraperitoneal (i.p.) injection of a mixture of ketamine (80 mg/mL, i.p.) and xylazine (10 mg/mL, i.p.) (see Recipes) according to the following stages:
Prepare the anesthesia solution.
Lift the rat from the home cage and cover its head with a towel. Place it quietly on the cage lid to grasp its bars and become immobilized.
Grasp with one hand near the neck of the rat under the forelimb while holding the tail with the other hand. In this situation, the rat’s head is kept immobilized. Lift upward the shoulder and expose the abdomen, then inject anesthesia solution (i.p.) on the lower right quadrant of the abdomen.
Cut the neck close to the head (decapitation) with a rodent guillotine.
Sterilize the head, dissection area, and surgical tools by spraying 70% alcohol.
Hold the head in place with the lateral parts (temporal bones) between two fingers.
By using fine scissors, cut the skin out at the middle of head by a single transverse cut and retract the skin. Separate remaining muscles and tissues at the base of the skull with large scissors. Start removing bones from occipital to frontal parts.
Remove the supraoccipital bones over the cerebellum with the rongeur until it is exposed.
Now open the bones over the hemispheres in sequential steps: firstly, place the flat blade of a pair of scissors beneath the right parietal plate and force from the inside to drive away from the brain. Separate the left parietal plate in the same way.
Proceed rostrally, and remove the frontal skull plates by rongeur, starting from the site of the interfrontal suture. Remove bones to expose frontal parts of the brain (see Note 1).
To remove meninges, pierce fine scissors into the interhemispheric fissure and with a fine forceps take away the meninges over the hemispheres.
Loosen the exposed brain laterally in forward and backward motions with a round spatula.
Insert spatula inside of the skull floor and slide it to push brain away from the skull, cutting attached nerves with the same spatula, then invert brain immediately into a 50 mL Falcon tube containing 3–5 mL of cold PBS (1×) with 3% Pen/Strep and 1% Amphotericin B on water ice. Observation of the whole brain without any damage indicates its intact structure.
DG microdissection (Figure 3)
Figure 3. Dentate gyrus microdissection. A. Surgical tools. B and C. Dissection area. D. Transfer of isolated brain onto filter paper with curved forceps. E. Intact brain under a stereomicroscope. F. Cutting the cerebellum off with a scalpel. G. Splitting the brain into two hemispheres along the midline. H. Turning over each hemisphere onto the medial surface with forceps. I. Removal of the diencephalon inside of each hemisphere (indicated by a black line). J. The hippocampus inside of the cortex is visible. Middle part indicated by a black line is the dentate gyrus. K, L, and M. Hippocampus was separated from brain tissues by inserting a spatula at the isolating border and releasing into the dissection area. N. Hippocampus under stereomicroscope. O. Insertion of the spatula into the hippocampal fissure to isolate the dentate gyrus. P. Dentate gyrus (indicated by a black line) is observed in the center.
Place a Petri dish lid on the top of ice under a stereomicroscope.
Cover the Petri dish with filter paper and rinse it completely with 2 mL of cold dissection solution. Place an illuminator on the right side and focus it on the dissection area.
Transfer the isolated brain from a 50 mL Falcon tube onto the dissection area using curved forceps and rinse it with dissection solution (2 mL for each brain).
Roll the brain onto its ventral surface. Firstly, by using a scalpel, cut off the cerebellum while holding the brain with forceps, and then cut the brain along the sagittal plane into two hemispheres.
Turn over each hemisphere upward to view the medial side and hold them in place using forceps.
Initially, remove the diencephalon inside of each hemisphere at the center by inserting a spatula at the inner border of the fornix and diencephalon, and then sliding the spatula from posterior to anterior, finally removing the diencephalon.
Now the hippocampus is visible on the medial side. Separate cortex overlays from the hippocampus throughout the corpus callosum with spatula. Isolate the hippocampus from the septo-temporal direction and detach it from other brain tissues with spatula, then turn over the entire hippocampus onto the dissection area. Cut the remaining fragments of cortex and white matter tissue.
To dissect DG from the hippocampus, refocus the microscope, and observe the hippocampal fissure (in some rats it is not as visible).
In most brains, a blood vessel in the hippocampal fissure, the hippocampal arteriole, is apparent and serves as a landmark to separate the DG from the CA1 area. The DG is in the middle area when the hippocampus is observed from the medial plane. While holding the hippocampus in place using forceps, slide along the hippocampal fissure with a fine spatula to cut the DG off, then transfer it to the microtube containing 1 mL of cold PBS (1×) or DMEM/F12 (1:1).
DG tissue dissociation (Figure 4)
Figure 4. Dentate gyrus (DG) dissociation. A. Transfer the microtube containing DG tissues under a laminar hood. B. Add 1 mL of PBS to wash the tissue. C. Use a sterile scalpel to cut the tissue up. D. Mince completely. E. Add enzyme solution. F. Collect tissue fragments into a Falcon tube. G. Add enzyme solution according to the amount of tissue fragments. H. Keep on the shaker. I. Cell suspension is visible, looking milky. J. Pellet of single cells after centrifugation is shown.
Transfer the microtube containing DG tissues under a laminar hood.
Empty the pieces of DG into a Petri dish under the laminar hood and add 1 mL of PBS (1×) containing 1% Pen/Strep antibiotic for two dentate gyruses to wash the tissue twice, aspirate the solution out of the Petri dish, and add 500 μL of PBS (1×) to cover the tissue. Keep tissue fragments wet.
Use a sterile scalpel to cut the tissue up and mince it completely for 2–3 min according to the tissue amount. Rotation of the Petri dish during mechanical digestion makes dissociation more uniform. Cut 1 mm of the initial segment of 1 mL pipette tip with a scalpel to broaden the hole of the tip and fill it with tissue fragments.
Transfer minced tissue into a 50 mL Falcon tube containing pre-warm Papain (0.5 mg/mL), Dispase (2 mg/mL), DNase (0.02 mg/mL) (PDD) enzyme solution and incubate at 37 °C under shaking (150 rpm). Use 1 mL of enzyme solution for each animal.
Incubate in an enzyme solution as follows: for young adult rats, 10–15 min incubation time is enough. Longer times lead to greater debris and lower cell survival. For aged rats, an incubation time of 15–20 min works efficiently. After 10 min, transfer the tube under the laminar hood and slowly pipette up and down 5–7 times with a P1000 pipette. Every 5 min, observe the Falcon tube; if the tissues get soft and the solution milky, stop digestion.
After enzyme incubation, monitor the single-cell suspension. If cells are suspended, the solution will appear milky. If further large tissues remain, it is better to remove the upper liquid and continue onto the next steps. To dissociate large particles, add more enzymes to the Falcon tube and incubate for 5 min.
To stop enzyme activity, add basic DMEM/F12 medium at three times the volume of the enzymatic solution. Pipette suspension up and down 6–7 times, then centrifuge at 95× g for 5 min.
Discard the supernatant, re-suspend the cell pellet in growth medium (1 mL), and gently pipette up and down 3–4 times. Avoid bubble formation and rapid pipetting; it reduces cell yield and produces more debris. Optional: It is possible to add more medium up to 10 mL and filter through a 70 µm cell strainer. Place cell strainer on a 50 mL Falcon tube and strain the solution through it; this removes un-dissociated tissue clumps, white matter, and connective tissue. However, it might be at the expense of losing some NSPCs.
Cell counting and plating
Perform cell counting on obtained single cells.
In some culture conditions, the isolation procedure fails, and no single cell is obtained. The bright round cells are not visible, and the culture is full of undigested tissue and dead cells. Therefore, this culture is not appropriate for plating. However, the presence of debris and dead cells is inevitable in primary culture. The experimenter must recognize single cells that are small, bright cells. Cell debris have uneven shapes with one layer observable under the microscope, but live cells have three layers and round shapes. Debris appears as a dot-like spot in the culture. Also, it is possible to use DAPI staining and trypan blue staining to distinguish debris from isolated neural stem cells.
Dilute 10 μL of the cell suspension with 10 μL of 0.4% trypan blue in a microtube.
Transfer 10 μL from the mixture onto hemocytometer chambers.
Count four individual squares (WBC cell count squares).
Divide the total number by 4, multiply by 2 (dilution factor) and 104.
The area of each square is 1 mm × 1 mm. The space between the coverslip and slide is 0.1 mm. Therefore, the volume of a large square is 1 mm3 or 0.0001 mL. According to the properties of the hemocytometer chamber, to measure cell count per milliliter, the average number of cells is multiplied by the dilution factor and 104.
By this calculation, the total number of cells in 1 mL of cell suspension is measured.
Measure and optimize cell densities of your interest in specific cell culture vessels (see Note 2). 2 × 105 cells for a T-25 flask, 1.5 × 105 cells per well for 6-well plates, 1 × 104 per well for 12-well plates. It depends on the number of animals used for each condition. (Maintenance and growth of NSPCs in DMEM/F12 require plating in high density. Complete media prepared by basic DMEM/F12 medium, not neurobasal A.)
For aged rats, plate cells as following densities: 2.5 × 105 cells for the T-25 flask, 1.5 × 105 cells per well for 6-well plates, and 1 × 105 cells per well for 12-well plates. Use adherent flasks and well plates. For neurosphere culture use 1.5 × 105 cells per well in non-adherent 6-well plates.
Generation of adherent monolayer culture (Figures 5, 6)
Figure 5. Culture of neural stem/progenitor cells (NSPCs) isolated from the dentate gyrus of young adult rats as a monolayer at different days in vitro. A. Day 2: neural stem/progenitor cells are visible as bright small round cells; these cells start attaching to the surface by day 3, and by day 5 most of the proliferating cells developed processes with small cell bodies. B. Day 18: represents proliferating NSPCs, which were growing continuously to occupy the culture vessel. C. By day 28, NSPCs display a substantial increase in numbers and were inclined to make connections. D. On day 38, rosette-like structures appeared. All scale bars are 100 μm with a 10× objective lens.
Figure 6. Culture of neural progenitor cells from dentate gyrus of adult aged rats as monolayer on different days in vitro. A. Day 12: single cells broaden their processes and grow rapidly. B. Day 19: proliferating neural progenitor cells as spindle-like shape are shown and proliferated to confluent vessel. C. Day 21: proliferated neural progenitor cells at the 80% confluency are visible. D. Day 51: most of the cells present smaller processes. All scale bars are 100 µm with objective lens 10×.
Plate isolated cells onto a PLO/laminin-coated surface of a T-25 flask, which provides an adequate substrate for the maintenance and growth of NSPCs.
Allow cells to expand without disturbance. Leave cells in an incubator for up to 4 days for young adult rats and 7 days for aged rats. Monitor the culture 3–4 days after plating for young adult rats and 7 days for aged rats. Once cells adhere to the surface, remove all the medium above cells using a P1000 pipette and empty it into a 15 mL Falcon tube.
After the initial days since attached cells start expanding, add growth factors (20 ng/mL) and heparin (2 µg/mL) into the culture every 2 days.
After 7 days for young adult rats and 10 days for aged rats, remove 2.5 mL of medium from the T25 flask and add 2.5 mL of fresh medium.
Check the confluency.
At 90% confluency, tap the flask slowly with the palm of your hand to dissociate these cells for biochemical assays.
It takes 10–15 days to reach 80% confluency in the T25 flask for NSPCs obtained from young adult rats. For aged NSPCs, it takes 20–25 days to reach 80% confluency (see Figures 5 and 6).
If there are non-adherent cells or floating spheres in the medium, carry out the following steps:
Remove medium above cells using a P1000 pipette and empty it into a 15 mL Falcon tube.
Slowly pipette up and down 5–6 times with a P1000 pipette to separate the neurospheres and non-adhered cells from debris attached to them. For some cases, you can centrifuge at 95× g for 5 min and get a pellet and then add 300 μL of accutase for 3 min at room temperature.
Centrifuge at 95× g for 5 min at room temperature.
Remove the supernatant and add the same volume of complete serum-free medium to the pellet. Pipette up and down 3–4 times, then return them to their wells (see Note 3).
Subculture of adherent monolayer cultures
When cells reach approximately 80% confluency, they must be subcultured to avoid cell detachment and overgrowth, as well as to maintain cells in a healthy condition.
Perform the following steps for subculturing adherent monolayer cultures.
Remove old medium completely, then add 2 mL of PBS (1×) into the T25 flask. Slowly rinse the flask by slightly leaning it from side to side. Remove the PBS (1×) after 30 s.
Add 1 mL of accutase to the T25 flask; incubate at 37 °C for 2–3 min while observing the T25 flask under the microscope, and check for detachment of cells. If they are not detached, gently tap each side and bottom of the flask to assist in the detachment of cells. Critical: It is efficient to detach cells mechanically with a minimal volume of an enzyme.
Longer incubation time in enzyme solution causes the low attachment of single cells to the surface after subculture. Severe triturating contributes to low cell yield. Accutase is an enzyme solution of proteolytic and collagenolytic enzymes. It can be used as a direct replacement of trypsin.
To inactivate enzymatic activity, add three times the volume of the enzyme solution as DMEM/F12 medium into the T25 flask.
Pipette up and down several times, then collect the whole medium, which now contains single cells, into a 15 mL Falcon Tube.
Centrifuge at 95× g for 5 min at room temperature
Re-suspend the pellet in 1 mL of complete serum-free medium very gently. Try not to form bubbles. Immerse pipette tips in the solution thoroughly to the bottom of the Falcon tube and touching the wall of the tube, then start pipetting slowly. Seed the 100,000–150,000 cells in a 6-well plate (Table 1).
Seed the cells with high density on the coated well plate. At first passage, you can use either 100,000 cells per well of 6-well plates or 50,000 cells per well of 24-well plates. In the next passages, it is better to increase cell density.
Table 1. Recommended cell densities
Cell density Cell culture plate
100,000–150,000 Per well of 6-well plates
50,000–70,000 Per well of 24-well plates
10,000–20,000 Per well of 96-well plates
Neurosphere culture (Figures 7, 8)
Figure 7. Culture of neural progenitor cells from the dentate gyrus of young adult rats as neurospheres on different days in vitro. A. Day 4. B. Day 8: clusters of cells were formed. C. Day 11: neurospheres show round morphology. D. Day 22: size of spheres increased. E. Day 25: spheres grew larger, and their centers appeared dark. F. Day 27: most of the neurospheres with dark centers were apparent. All scale bars are 50 μm with a 20× objective lens.
Figure 8. Culture of neural progenitor cells from dentate gyrus of adult aged rats as neurospheres on different days in vitro. A. Day 25: neurospheres are formed as their spherical shapes take form and their size increases. B. Day 32: neural stem/progenitor cells (NSPCs) proliferated, and diameter of spheres increased sharply. C. Day 36: neurospheres proliferated rapidly as their centers get a dark appearance. D. Day 41: large and dark neurospheres appeared. E. Day 48: neurospheres begin to attach to the surface. All scale bars are 50 μm with a 20× objective lens.
Seed the obtained single cell suspension (2.8–3.5 × 105 cells per milliliter) from the digestion of DG tissue for neurosphere culture as follows:
Re-suspend the cell pellet in 1 mL of serum-free growth medium, then count the cells and seed them per recommended numbers (5 × 105 cells for aged rats and 3–4 × 105 cells for young adult rats) in a non-adherent T25 flask. Then, incubate at 37 °C with 5% CO2 (see Note 4). Critical: Seeding the cells at a higher density than recommended leads to cell aggregation.
It is necessary not to disturb the cells after plating until 4–5 days for the young adult group and 7 days for the aged group. Let cells adapt to culture medium for 4–5 days and start formation of spheres. Avoid any disturbances or movement of the flask (see Note 5).
After this initial incubation period, re-plate cell culture as follows: collect all the medium inside the T25 flask (which contains cells and small cell clusters starting the formation of neurospheres) into a 15 mL Falcon tube.
Add 1–1.5 mL of PBS (1×) to the T25 flask, then pipette the entire surface of the flask, and discard into a 15 mL Falcon tube.
Centrifuge at 95× g for 5 min at room temperature.
Re-suspend the cell pellet in 1 mL of neurosphere medium and pipette up and down 2–3 times slowly.
Transfer to one well of a 6-well plate.
Add growth factors and heparin every 3–4 days; growth factors (20 ng/mL) and heparin (2 μg/mL).
After 14–16 days, spheres are formed for young adult rats. Neurosphere of aged rats begin to grow from 10 days post-plating onward. After 28–30 days aged spheres are fully formed (see Note 3 as well as Figures 7 and 8). If the medium becomes yellow, add 1 mL of serum-free medium after 3 days from beginning of culture.
Quantification of the number of neurospheres
The number of neurospheres serves as an index of NSPCs proliferation. At the stage (optimal time) that the neurospheres reached maximum growth without a dark appearance in the center, estimate the number of spheres as follows:
Place the cell culture well plate under an inverted microscope, objective lens 20×.
Count spheres field-by-field, starting from a corner and going along the well diameter up to the next corner.
Adjust the focus to prevent the detection of cell aggregates instead of healthy spheres.
Quantify the size of the neurospheres
The diameter of neurospheres is used as an index of neurosphere growth.
With a 20× objective lens, take non-overlapped random images from an entire well (8–9 fields), as most of the spheres are included in the field. Use Infinity software to measure diameters of neurospheres. Average of two measured diameters is representative of neurosphere diameter. Then, express different diameters for each image. For example, if there are seven neurospheres in an image, calculate the diameter of each one. Now you have seven neurospheres with different diameters as there are three neurospheres with a diameter of 58 ± 9 micrometers, two neurospheres with a diameter of 89 ± 13 micrometers, and two neurospheres with a diameter of 120 ± 9 micrometers. This provides valuable quantification data to indirectly evaluate the proliferation rate of NSPCs on different days of in-vitro culture.
Use Infinity software as following steps:
Open Infinity software > select file > open image > adjust > micrometer > select a ruler from tool bar > place in the length of scale bar and right click > open a page > write 1 in the magnification box and write the length of scale bar in the length box > draw a horizontal line and a vertical line, and right click > length is shown (Figures 9, 10, and 11).
Figure 9. Quantification of neurosphere size by Infinity software. Open the image.
Figure 10. Open the file, adjust the micrometer, write scale bar length in the length box, and write 1 in the magnification box
Figure 11. Draw two lines and calculate the average two numbers to acquire diameter of each sphere
Passage of neurospheres
Remove the content of the plate and transfer it to a 15 mL Falcon tube.
Tapping the plate and pipetting with DMEM/F12 medium to dislodge the attached spheres.
Centrifuge at 95× g for 5 min.
Remove supernatant and add 300–500 μL of accutase accurately onto the pellet (neurospheres).
Incubate at 25 °C for 4–8 min, based on the size and number of neurospheres. Critical: During enzyme incubation, it is possible to pipette up and down slowly 4–5 times without generating bubbles. Observe under light to see whether neurospheres are dissociated.
Add complete serum-free medium two times the volume of the enzyme and mix gently.
Centrifuge at 95× g for 5 min.
Re-suspend the pellet in 1 mL of neurosphere medium.
Count the cells and plate at the same density of 5 × 105 cells/mL. Critical: The number of neurosphere passages is important to determine cell density. As in our culture system the proliferation rate decreases with increasing passage numbers, it is essential to plate cells at higher density in stages following the first passage.
Immunostaining (Figure 12)
Seed 1–2 × 104 cells per well into a 96-well plate (see Note 6).
Aspirate medium from wells and wash with 100 μL PBS (1×).
Aspirate PBS completely and add 100 μL of 4% paraformaldehyde (PFA) in PBS (PH 7.4), then incubate for 20 min at room temperature (it is advisable not to use cold PFA).
Aspirate PFA and wash each well with 100 μL of PBS (1×) (three times).
Incubate for 10 min with 70–100 μL of 0.2% Triton X-100 to make cell membranes permeable.
To block non-specific antigens, add 100 μL of blocking solution (blocking solution is composed of 10% normal goat serum, 1% BSA, 0.1% Tween in PBS) (see Recipes) and incubate for 60 min.
Aspirate blocking solution.
Wash with 100 μL of PBS (1×) (three times).
Dilute primary antibodies in blocking solution as follows:
To prepare 350 μL of 1:200 Anti-Nestin solution, add 1.75 μL of Nestin antibody to 348 μL of blocking solution. Prepare before use.
To prepare 350 μL of 1:100 Sox2 solution, add 3.5 μL of Sox2 antibody to 346.5 μL of blocking solution. Prepare before use.
To prepare 250 μL of 1:250 GFAP solution, add 1 μL of GFAP antibody to 249 μL of blocking solution. Prepare before use.
Add 70 μL of primary antibodies to each well, seal the edges of the plate with parafilm to prevent evaporation of antibody solution, and incubate for 3 h at room temperature or overnight at 4 °C, preferentially inside a humid chamber.
Aspirate primary antibodies and wash with 100 μL of PBS (1×) (three times) every 5 min.
Dilute secondary antibodies with PBS (1×) (1:1,000 for both secondary antibodies). To prepare 2 mL secondary antibody solution, add 2 µL of secondary antibody into 1,998 µL of PBS (1×).
Incubate cells with 70 μL of secondary antibodies for 2 h at room temperature in the dark.
Add 100 μL of PBS (1×) into each well and incubate for 5 min, remove PBS, repeat the procedure three times, then cover the entire 96-well plate with aluminum foil to prevent light penetration.
Add 70 μL of DAPI solution per well and incubate for 1 min in the dark at room temperature.
Remove the DAPI solution and wash each well with 100 μL of PBS (1×) twice.
Capture images immediately using an inverted fluorescence microscope. For negative control, primary antibodies were not added (see Note 7).
Count and report the percentage of immunopositive cells (Figure 12).
Figure 12. Immunocytochemistry to detect antigenic markers of neural progenitor cells cultured as a monolayer. Neural progenitor cells expressed (A) Sox2 antigen, (B) Nestin antigen, and (C) GFAP (marker of astroglial cells). All scale bars are 100 μm with a 10× objective lens.
Data analysis
For monolayer culture, multiple wells (4-well plates or 6-well plates) containing cells derived from a primary culture or a passage serve as technical replicates to balance variations.
Independent repeats of different experimental settings or passages serve as biological replicates. Each experimental condition and cells derived from it are generally regarded as a biological replication. Four to six biological replicates were used for each condition to reduce false positive data and make data credible. The number of N conditions is defined as biological replicates and the number of wells for each N to repeat the same condition is defined as a technical replicate. Numbers of N for sphere numbers and their sizes in two young adult and aged rats in days in vitro are represented in the following tables (Tables 2–5).
Table 2. Size of sphere (µm) in young group in days in vitro for each N
Young group/sphere size D5 D10 D12 D14 D15 D17 D18 D20
N1 37 54 49 87 153 127 114 183
N2 49 47 50 49.54 104 108.50 129 137
N3 39 64 101 105 190
N4 30 109
Table 3. Size of sphere (µm) in aged group in days in vitro for each N
Aged group/sphere size D5 D10 D15 D17 D18 D31 D39
N1 0 39 45 42 37 68 77
N2 45 42 39 73 60 79
N3 33 32 29 63 99
N4 55 78 152
Table 4. Number of spheres in young group per passage for each N
Young group/sphere number P1 P2
N1 70 79
N2 178 86
N3 145 129
N4 89 28
N5 59 137
Table 5. Number of spheres in aged group per passage for each N
Aged group/sphere number P1 P2
N1 56 79
N2 98 65
N3 78 87
N4 25 12
Number of spheres and size of spheres are presented in the following figures (Figures 13, 14, and 15).
Figure 13. Comparison of size of spheres in days in vitro between young adult and aged rats. Size of spheres were greater in young adult group (not significant). Data was presented as mean ± SEM.
Figure 14. Comparison of number of spheres per passage between young adult and aged rats
Figure 15. Percentage of positive cells for neural stem/progenitor cells (NPCs) markers per dentate gyrus (DG). Approximately 80% of total cells are SOX2 positive. Data presented as mean ± SEM.
Through this method, DGs from four to six adult rats yield 90% confluency in a monolayer system after 15 days. For aged rats, DGs from six rats produced appropriate confluency in monolayer culture after 25 days.
Cells isolated through this approach represent the identifiable morphology of adult NSPCs. These cells also represent typical markers of NSPCs. Sox2 was expressed in 80% of cells. Nestin was expressed in most of the cells, and GFAP was positive in a few cells. For each well, 6–8 non-overlapping microscopic fields were captured with a camera. The percentage of immunopositive cells was quantified as the following equation:
Number of immunopositive cells/number of DAPI-labeled nuclei × 100 = percentage of immunopositive cells.
Percentage of SOX2 and GFAP positive cells for each N was measured (Table 6). Calculated Sox2 and GFAP positive cells are shown in following bar graph (Figure 15).
In this protocol, we expanded cells as a neurosphere culture, which is a well-known culture method for NSPCs with high purity. Actually, according to this protocol, neurospheres that are cultured on the coated plates proliferate as monolayers and are mostly Sox2 positive.
Mature neurons were present in the culture, recognized by their long processes. Therefore, the presence of antigenic markers on the obtained cells, which are specific for NSPCs, was confirmed. This is an efficient and reliable protocol for the culture and expansion of aged rat NSPCs (20–24 months), which was previously reported to be challenging and difficult.
Table 6. Percentage of SOX2 and GFAP positive cells for each N
Sample Percentage of Sox2 positive cells Percentage of GFAP positive cells
N1 82.30452675(200 of 243) 12.23684 (93 of 760)
N2 68.02721088 (300 of 441) 16.94444 (61 of 360)
N3 91.18329466 (393 of 431) 8.333333 (42 of 504)
N4 20.13889(58 of 288)
Mean 80.50501077 14.41338
Standard deviation 11.68245468 5.192688
Validation of protocol
According to the data obtained from monolayer and neurosphere cultures and their immunostaining, we introduced a reliable and reproducible protocol for isolating and expanding adult rat neural progenitor cells derived from the DG of young adult and aged rats. These cells carry specific markers of neural stem cells and reliably reproduce across all experiments and assays. Through the neurosphere cell culture conditions, the stem cells from young adult rats produced free-floating neurospheres up to four passages, and proliferation slowed down at passages 4–5. Neurosphere formation from stem cells of aged rats proceeded until passage 2 under our culture conditions, and there were no identifiable neurospheres at passage 3. In some studies, NPCs of aged rats were expanded through more passages. This study will add more information about the derivation and expansion of adult rat NSPCs of the DG.
Notes
Skulls of aged rats are thicker than those of young adult rats, so their removal is harder and takes longer.
Count cells as quickly as possible, especially in aged rat NSPCs, which are more sensitive.
It requires special attention and effort to handle rat NSPCs gently and slowly while exchanging media. Any vigorous manipulation leads to the detachment of NSPCs.
If there were large enough tissue particles on the first day of culture as to prevent precise counting, count and plate cells at this step (according to our experiments).
These cells are very sensitive, and factors such as quick pipetting, alterations of supplement concentrations, and severe movement of culture vessels prevent cells from attaching. Try not to form bubbles.
Three replicates were used for each marker.
Do not delay taking photos as it may result in fading fluorescence.
Necessary precautions:
Read safety data sheets for all materials and pieces of equipment.
Trypan blue is a carcinogenic agent. Use personal protective clothing and avoid contact with your hands, skin, and eyes. Do not breathe it.
Manipulate all products in an aseptic condition. Use gloves, masks, and protective clothing.
Acknowledgments
This work was supported by a grant (No. 3/54633) from the Ferdowsi University of Mashhad. The authors are very grateful to the Shefa Neuroscience Research Center, Khatam Alanbia Hospital, Sepideh Ghasemi, and Dr. Hadi Aligholi for their technical and scientific support. This study was a part of the Ph.D. thesis of Mina Afhami at Ferdowsi University of Mashhad.
Competing interests
The authors have no conflicts of interest.
References
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Aligholi, H., Hassanzadeh, G., Azari, H., Rezayat, S. M., Mehr, S. E., Akbari, M., Attari, F., Khaksarian, M. and Gorji, A. (2014). A new and safe method for stereotactically harvesting neural stem/progenitor cells from the adult rat subventricular zone. J. Neurosci. Methods 225: 81–89.
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Neuroscience > Cellular mechanisms > Cell isolation and culture
Cell Biology > Cell isolation and culture > Monolayer culture
Stem Cell > Adult stem cell > Neural stem cell
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This is a correction notice. See the corrected protocol.
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Correction Notice: Establishment of Human PD-1/PD-L1 Blockade Assay Based on Surface Plasmon Resonance (SPR) Biosensor
TP Tess Puopolo *
HL Huifang Li *
JG Justin Gutkowski
AC Ang Cai
NS Navindra P Seeram
HM Hang Ma
CL Chang Liu §
(*contributed equally to this work, § Technical contact)
Published: Aug 20, 2023
DOI: 10.21769/BioProtoc.4844 Views: 227
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In the Background section of “Establishment of Human PD-1/PD-L1 Blockade Assay Based on Surface Plasmon Resonance (SPR) Biosensor" (https://bio-protocol.org/e4765), “Alsaab et al., 2018” is incorrectly referenced. The correct citation is “Alsaab et al., 2017”. The sentence should read as follows: “Clinical data have demonstrated that the blockade of PD-1 or PD-L1 can boost T cell–mediated antitumor responses, generates durable clinical responses, and prolongs patient survival time (Ohaegbulam et al., 2015; Alsaab et al., 2017).”
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Epicutaneous Application of Mannan Induces Psoriasis-like Inflammation in an Inbred Mouse Strain
HW Huimei Wu
KN Kutty Selva Nandakumar
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4845 Views: 564
Reviewed by: Komuraiah MyakalaKrishna NakuluriMichael David Schultz
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Original Research Article:
The authors used this protocol in Clinical and Experimental Immunology Mar 2023
Abstract
Mannan from yeast induces psoriasis-like inflammation in the skin of inbred mouse strains. Limitations of available models led us to develop a new psoriasis model with a rapid disease onset, severe disease course, short duration, and a simple and easy-to-induce protocol with much more practically convenient features and cost-benefits. Mannan-induced skin inflammation (MISI) is more severe than the classical imiquimod (IMQ)-induced skin inflammation (IISI), with characteristic features resembling human plaque psoriasis but with relatively fewer toxicity issues. Epicutaneous application of mannan (5 mg) in incomplete Freund’s adjuvant or Vaseline induces severe psoriasis in BALB/c female mice. Psoriasis area and severity index (PASI) and histological evaluation of the skin could help assess the disease development. MISI mimics natural environmental factors affecting the skin relatively more closely than IISI. This disease model can be used to dissect inflammatory pathways in the skin, identify genetic and environmental factors affecting psoriasis, and test potential pharmacological agents or new combinations of available drugs for treatment before designing clinical trials.
Key features
• S. cerevisiae mannan induces psoriasis-like skin inflammation(MISI) when applied on the skin of inbred mice.
• The MISI model has a rapid onset, severe disease, short duration, and simple and easy-to-induce protocol.
• MISI is more severe than imiquimod-induced skin inflammation (IISI).
• Female mice had a more severe disease than males in the MISI model, thereby allowing the study of sex-dependent disease mechanisms.
• The MISI model identifies skin inflammatory pathways and genetic/environmental factors affecting psoriasis.
• The MISI model can be used as a drug testing platform for potential pharmaceuticals to develop new therapeutics for psoriasis patients.
• The MISI model can be used to explore the relative contribution of different pattern recognition receptors in the development and severity of psoriasis.
Graphical overview
Keywords: Mannan Psoriasis Mouse model Imiquimod Inflammation
Background
Psoriasis is a skin inflammatory disease affecting 3% of the population (Girolomoni et al., 2017; Zhong et al., 2018). Psoriasis etiology is still unclear, but a strong association with human leukocyte antigens (HLA) was documented (Knight et al., 2012; Tsoi et al., 2012). Environmental factors like sex, stress, and smoking affect the clinical manifestations of psoriasis (Fry and Baker, 2007). Keratinocyte hyperproliferation, increased blood vessel formation, and infiltration of immune cells are common in psoriasis skin (Van der Fits et al., 2009). Innate and adaptive immune responses are indispensable for disease development (Villanova et al., 2014). Proinflammatory cytokines and the immune cells secreting them activate the effector cells present in the skin. For example, T cell–derived cytokines like IL-17A and IL-17F activate keratinocytes, which produce anti-microbial peptides and more cytokines and chemokines. These inflammatory factors shape psoriasis skin inflammation (Ando et al., 2015).
Mannan is present in nearly all fungi species. Saccharomyces cerevisiae and Candida albicans mannans are pathogenic factors in psoriasis (Picciani et al., 2013). An intraperitoneal injection of mannan extracted from Saccharomyces cerevisiae cell wall was shown to induce psoriatic arthritis and psoriasis symptoms in reactive oxygen species (ROS)-deficient, Ncf1-gene-mutated mice that are available only in specific labs (Khmaladze et al., 2014). On the other hand, an epicutaneous application of imiquimod (IMQ) for 5–7 days leads to the development of an acute psoriasis phenotype. However, the disease developed in IMQ-induced psoriasis is often accompanied by severe weight loss in BALB/c mice (Swindell et al., 2017). Herein, we describe an induction protocol for plaque psoriasis-like inflammation in an inbred mouse strain (BALB/c) by epicutaneous application of mannan (Figure 1). A local (epicutaneous) instead of the intraperitoneal application of mannan could mimic a more natural way of skin exposure to environmental triggers for skin disease induction. Induced skin inflammation resembles human plaque psoriasis. Redness covered with white scales and skin thickness in the lesion area in mannan-exposed skin is more straightforward, making this model simpler for visually observing most manifestations of the disease.
Figure 1. Clinical scoring for psoriasis inflammation. A representative psoriasis inflammation scoring method is given. PASI scores for the above skin conditions are as follows: redness: 4; scales: 3; thickness: 3. Each group contains 10 mice. BALB/cfemale mice were used for experiments. Left column pictures were reproduced from Figure 1A of the published paper (Wu et al., 2023).
Moreover, the proliferation of keratinocytes and innate immune cells, the infiltration of T lymphocytes in the skin, and an increased expression of proinflammatory cytokines were induced after epicutaneous mannan application (Wu et al., 2023). The activation of TLR7 and TLR8, mainly expressed in the immune cells, mediates the effects of IMQ (Bong et al., 2002). In contrast, mannan is a ligand for mannose receptors and is the primary trigger of IL-17 pathways in the host (Jiang et al., 2017). In addition, a higher-level expression of TLR2, TLR4, CD206, Dectin-2, Mincle, and DC-SIGN receptors was found in mannan-induced skin inflammation (MISI) (Wu et al., 2023). Thus, this disease model differs from IMQ-induced psoriasis-like inflammation, mainly in terms of the health status of the animals. Unlike the IMQ model, the body weight loss in MISI is negligible.
Furthermore, mannan had a shorter exposure time (3 days) to induce the disease than IMQ (5–7 days), demonstrating more cost-effective and convenient features. A response to dexamethasone in MISI has shown its usefulness in testing and validating various anti-psoriatic drugs for future treatments in psoriasis patients. Many autoimmune diseases show sexual dimorphism during disease development, and this plaque psoriasis-like skin inflammation induced by mannan also showed strong female preponderance, demonstrating its use in exploring sex-dependent mechanisms (Wu et al., 2022). Although mannan-induced skin inflammation results in acute and relapsing disease symptoms after repeated exposure, like other induced acute psoriasis mouse models, this model can also not completely mimic the relapsing and chronic characteristic features of human plaque psoriasis.
Materials and reagents
Biological materials
8–12-week-old BALB/c female mice maintained in a pathogen-free animal house were purchased from Southern Medical University and Guangdong Medical Animal Experiment Center. Each group contains 5–12 mice. All animal experiments were performed per the guidelines of the National Institutes of Health (NIH Publication No. 8023) and approved by the ethics committee of Southern Medical University (l2018183). Mice were kept in cages in a climate-controlled environment having 12:12 h light/dark cycles and given food and water ad libitum. Southern Medical University Animal Care and Use Committee, Guangzhou, China, approved all the procedures.
BALB/c is among the most popular inbred mouse strains in many international research laboratories. The IMQ-induced psoriasis mouse model is commonly induced in these mice for drug evaluation. Skin redness can be more easily observed in this white mouse. BALB/c female mice were used in all the experiments described below.
Reagents
S. cerevisiae mannan (Sigma-Aldrich, catalog number: M7504) was dissolved (100 mg/mL solution) in sterile phosphate buffered saline, PBS (Gibco, catalog number: 14190-094).
Incomplete Freund’s adjuvant (IFA) (Sigma-Aldrich, catalog number: F5506-10)
Complete Freund’s adjuvant (CFA) (Sigma-Aldrich, catalog number: F5674)
Isoflurane (Healthcare, catalog number: 676544)
Phenobarbital (Sigma-Aldrich, catalog number: 57-30-7)
Imiquimod cream (AldaraTM, catalog number: 678432)
Hair removal cream (Nair, catalog number: 657789)
Paraffin (Leica, catalog number: 3960l095)
4% Paraformaldehyde (Leagene, catalog number: DF0135)
Hematoxylin solution (Mayer) (Beyotime, catalog number: C0105)
0.2% Eosin solution in PBS (Mayer) (Beyotime, catalog number: C0105)
Xylene solution (Beyotime, catalog number: C0105)
Neutral gum (Solarbio, catalog number: G8590)
Vaseline (Yuanye, catalog number: 8009-03-8)
Liquid Paraffin oil (Yuanye, catalog number: 67552)
Ethanol (absolute) (Mackline, catalog number: E821482)
Plus-loaded slides (Thermo Fisher, catalog number: 6885)
Xylene-based mounting medium (HistoLab, catalog number: 00801)
Solutions
Different concentrations of ethanol (see Recipes)
Mannan-PBS solution (see Recipes)
Phenobarbital-normal saline solution (see Recipes)
Preparation of mixture (see Recipes)
Recipes
Different concentrations of ethanol
Reagent Ethanol (absolute) H2O
95% ethanol 950 mL 50 mL
90% ethanol 900 mL 100 mL
80% ethanol 800 mL 200 mL
70% ethanol 700 mL 300 mL
50% ethanol 500 mL 500 mL
Mannan-PBS solution
Solution 100 mg/mL mannan-PBS solution PBS
50 mg/mL mannan-PBS solution 50 μL 50 μL
Phenobarbital-normal saline solution
Solution Phenobarbital powder Normal saline
80 mg/mL phenobarbital-saline solution (5 mL) 400 mg 5 mL
Preparation of mixture
Mixture Mannan-PBS solution (50 mg/mL) IFA
Mannan + IFA 100 μL 100 μL
Equipment
Small animal weighing device (CHIKO, model: ZK-DST)
Digital vernier caliper (Yasuwang, catalog number: ASO-1-894-01)
Mouse skin shaver (Riward, catalog number: CP-5200)
Tissue processing/embedding cassettes with lid (Sigma-Aldrich, catalog number: Z672122)
Brush (Asone, catalog number: CC-5667-04)
Vacuum tissue processor (Leica, model: ASP300S)
Embedding machine (Tissue-Tek; Sakura)
Microtome (Leica, model: RM2255)
Eclipse upright optical microscope (Nikon, model: Ci-E)
Software and datasets
Image-pro plus 6.0 software (Nikon; Ci-E)
GraphPad Prism 5 (GraphPad Prism; version 5)
Procedure
Basic Protocol 1
Mannan-induced skin inflammation
Epicutaneous application of 50 μL of mannan mixture with incomplete Freund’s adjuvant at the back of the BALB/c female mice for three days induces psoriasis-like skin inflammation. Monitoring skin redness, scales, and thickness daily for 7–9 days or until the inflammation subsides using the scoring protocol described in Support Protocol 1 is essential. Before starting any animal experiment, apply for and get valid ethical permits from ethical review boards of respective institutions/regions.
Protocol steps—step annotations
Preparation of mannan stock solution
Dissolve the mannan powder (5 g/vial) by gently adding 2 mL of sterile PBS and then transfer all the contents into a 50 mL centrifuge tube, adding another 48 mL of PBS to get a 100 mg/mL concentration.
Note: Avoid foam formation, though occasional shaking of the solution is required to dissolve the powder completely.
Aliquot and label the mannan solution in sterile 10 mL tubes and freeze them at -20 °C until used.
Note: Mannan solution at 100 mg/mL concentration is stable for over 12 months if stored at -20 °C.
Epicutaneous application of mannan for psoriasis induction
To get consistent results, 8–12-week-old BALB/c female mice (5–10 mice/group) from specific pathogen-free breeding centers are required. Acclimatize the mice for at least one week before starting the experiment.
Use at least five mice per cage in a climate-controlled environment with 12:12 h light/dark cycles; provide food and water ad libitum.
Use mice epicutaneously treated with PBS on the back, similarly to the mannan application (minimum five mice), for control.
Thaw 1 mL of mannan-PBS solution, previously stored in the freezer, at room temperature for 15 min.
Note: Avoid frequent freeze-thaw cycles. The volume of mannan thawed depends on the number of mice used in an experiment.
Remove the hair from the mouse skin at the back (2.0 cm × 3.5 cm area) first by using a small mouse shaver, which has a 2.4 cm wide blade that can cut as close as 0.1 mm to the skin.
Note: Avoid making wounds on the skin surface.
Use the hair removal cream to remove the remaining hair from the specified area of the mouse skin. Take the hair removal cream on a smooth plastic plate and then smear the cream on the surface of the skin. Wait for 20 min and finally remove the cream from the hair using a paper towel.
Accurately weigh the mice and give an intraperitoneal injection of 80 mg/mL phenobarbital-normal saline solution to anesthetize mice.
Note: The weight of mice is measured in grams and accurate to one decimal place, and intraperitoneal injection of phenobarbital-normal saline solution should be no more than 200 μL in volume.
Mix 100 μL of mannan-PBS solution (50 mg/mL) with IFA (100 μL) and apply mannan + IFA mixture for each mouse. Place mannan-PBS solution at the bottom of a 1.5 mL centrifuge tube, and then add IFA gently; use a pipette to mix mannan-PBS and IFA until the mixture becomes an emulsion. Treat at least five mice with the prepared mixture and use another five mice treated with PBS as a control.
Note: Mannan and IFA mixture need to be prepared before each experiment, and homogeneous mixture preparation is required before application on the back of mice to get reproducible results.
Smear the mixture on the back of the mouse uniformly using a cotton swab pre-soaked in PBS.
Note: Apply a thin layer of the mixture on the skin and avoid applying the mixture too much to any one place. Use a fine brush (natural or synthetic, but not foam) for application. Applying a higher concentration (10 mg) of mannan leads to acute inflammatory reactions, and the scales will peel off quickly, hence not optimal for psoriasis induction.
Repeat steps B8 and B9 for three consecutive days.
Note: Apply mannan-IFA mixture at 9:00–10:00 am for three days. Remember to apply mixture at 24-hour intervals.
Score the psoriasis lesions daily by following Support Protocol 1.
Note: The disease progression and scoring start 48 h after mannan treatment for the first time and should be monitored every 24 h for 9 days. Observe redness, scales, and thickness from day 2 onwards. Redness and scales are typically apparent after days 3 and 4, respectively. These psoriasis symptoms reach maximum severity on day 5, and the mice gradually start recovering from day 7 onwards. In most inbred mouse strains (BALB/c, C57Bl/6J, C57Bl/6NQ, KM, DBA/1, ICR, and NIH), signs of inflammation could be observed in naïve mice exposed to mannan under experimental conditions described herein (Wu et al., 2022 and 2023).
Support Protocol 1
Psoriasis Area and Severity Index (PASI) scoring protocol
After mannan application, mice should be inspected daily for disease symptoms, as shown in Figures 1–5. PASI involves scores for redness (0–4), scales (0–4), and thickness (0–4). The standard of PASI scoring was reported earlier (Van der Fits et al., 2009). The total score of PASI (maximum disease severity) per mouse is 12, as shown in the psoriasis inflammation scoring protocol (Table 1). Score redness and scales visually and measure skin thickness using a digital vernier caliper.
Table 1. Scoring standards of PASI for psoriasis inflammation in mice
Scores of psoriasis 0 1 2 3 4
Redness No Mild Moderate Severe Very severe
Scales No Mild Moderate Severe Very severe
Thickness No Mild Moderate Severe Very severe
Protocol steps—step annotations
Gently take each mouse from its cage and observe the back of the mice for psoriasis progression.
Note: The critical components of the PASI scoring are outlined in Table 1.
Evaluate the redness using a 0–4 scale for mannan and PBS-applied mice during disease. Example images are given below for redness scores (Figure 2).
Figure 2. Scores for redness (0 and 3) are given by observing the back skin of the mice. An example is shown in the above figure (n = 10/group). BALB/c female mice were used for experiments. Right column images with 3 points were reproduced from Figure 1A of the published paper (Wu et al., 2022).
Evaluate the scales using a 0–4 scale for mannan and PBS-applied mice during disease. Example images are given below for scoring the scales (Figure 3).
Figure 3. An example was given for scale scores (0 and 3) by observing the back skin of the BALB/c female mice in mannan-induced skin inflammation (MISI) (n = 10/group). The right column picture was reproduced from Figure 1A of the published paper (Wu et al., 2022).
Measure the skin thickness with a caliper daily.
Choose the same place and angle on the skin for measuring thickness, using a digital vernier caliper (0.1 mm = 1 score), above that of naïve mouse skin, scored as 0 (Figure 4).
Figure 4. Representative example for skin thickness scores (0 and 4) in BALB/c female mice after PBS or mannan treatment (n = 10/group). Skin thickness is measured using a vernier caliper.
Perform scoring procedures daily for psoriasis development (refer to Figure 5 and Table 2 ).
Note: The incidence of psoriasis is generally very high (100%) and consistent from experiment to experiment. However, disease severity is strain dependent. The most disease-prone strains are BALB/c and C57Bl/6 mice.
Figure 5. Development of psoriasis in mannan-induced psoriasis from days 1 to 9 in BALB/c female mice (n = 10 mice/group). The image of day 6 was reproduced from Figure 1A of the published paper (Wu et al., 2022).
Table 2. An example of individual and total PASI scores in BALB/c female mice from days 1 to 9 in MISI
Day Redness Scales Thickness PASI
1 0.5 0 0 0.5
2 1.2 1.0 0.8 3
3 2.0 2.0 1.9 5.9
4 2.5 2.8 2.5 7.8
5 3.5 4 3.5 11.0
6 2.5 2.8 2.8 7.1
7 1.5 1.8 1.9 5.2
8 1.0 1.5 1.0 3.5
9 1.0 0.5 0.5 2.0
Skin starts to show redness from day 1 and scales from day 2 after the mannan-IFA mixture application, with a maximum psoriasis severity at days 4 or 5. Symptoms of psoriasis disappear entirely around day 9. A second cycle of exposing the mice to mannan on days 10–12 induced even more severe psoriasis symptoms than the first cycle of exposure (Wu et al., 2023).
Basic Protocol 2
Imiquimod-induced skin inflammation (IISI)
Imiquimod (IMQ) treats keratosis, basal cell carcinoma, and external genital/anal warts. However, topical treatment with IMQ induced typical psoriasis-like inflammation, including increased epidermal thickness, erythema, and skin thickness (Cai et al., 2011). IMQ is a toll-like receptor-7 and -8 (TLR7 and 8) ligand and can exacerbate psoriasis development in patients, possibly acting via the IL-17/IL-23 axis.
Protocol steps—step annotations
Divide IMQ cream from a 250 mg pack into five portions.
Note: Preparing equal portions of Aldara cream may sometimes be challenging to achieve. Hence, careful attention is required to have a constant amount of IMQ smeared on the back of mice.
Thirty-six mice were randomly divided into three groups containing PBS, mannan, and IMQ groups. Remove the fur using a mouse shaver and hair removal cream from a 2 cm × 3.5 cm area of the back of the mouse.
Note: Avoid making wounds on the skin surface.
Accurately weigh the mice and give an intraperitoneal injection of 80 mg/mL phenobarbital-natural saline solution to anesthetize mice.
Note: The weight of mice is measured in grams and accurate to one decimal place, and intraperitoneal injection of phenobarbital-natural saline solution should be no more than 200μL in volume.
Apply 50 mg of IMQ cream on the back of the mouse from 9:00 to 10:00 am daily for five days consecutively.
Note: Apply the cream at 24 h intervals for five days. Apply a thin coat of IMQ cream on the skin and avoid applying too much cream to any one place. Use a fine brush (natural or synthetic, but not foam) to apply the IMQ cream.
Score mice’s redness, scales, and thickness before applying IMQ using Support Protocol 1.
Notes:
A similar disease course was observed in MISI and IISI, as shown in Figure 6. Individual scores and total PASI in MISI (day 5) and IISI (day 5) were given in Table 3.
There is negligible weight loss after mannan application, while in the IMQ-induced psoriasis model, 12%–15% loss of initial weight was observed (Wu et al., 2022).
Figure 6. Comparison of psoriasis area and severity index (PASI) scores and clinical disease of mannan with imiquimod (IMQ)-induced psoriasis. PASI and psoriasis disease in female BALB/c mice. (A) PASI scores of PBS- mannan and IMQ-induced psoriasis-like skin inflammation (n = 12/group). (B) Representative images of PBS-, mannan-, and IMQ-applied mice at the peak of psoriasis (day 5, day 7 for IMQ) were shown. The data represent mean ± SEM. **, p < 0.01.
Table 3. Individual and total PASI scores in BALB/c female mice after mannan or IMQ application
Scores PBS Mannan IMQ
Redness 0.5 2.5 3.2
Scales 0 3 2.5
Thickness 0 3 2.5
PASI 0.5 8.5 8.2
Basic Protocol 3
Histological staining of psoriasis skin
Use hematoxylin and eosin staining to assess the skin morphology from psoriasis and naïve mice. Hematoxylin has a negative charge, and it is alkaline, which gives it a blue color after binding with the nucleus. Eosin binds with protein cations so that the cytoplasm is stained red. Hematoxylin and Eosin (H&E) staining can show the epidermis’ hyper-proliferation and immune cell infiltration in the epidermis and dermis. Determine the epidermal thickness by measuring the average interfollicular distance under the microscope in a blinded manner. Histopathological evaluation of diseased skin sections shows hyperkeratosis and acanthosis, elongated “rete-like” ridges, and infiltration of immune cells in the diseased skin.
Protocol steps—step annotations
Cut a specified area (2 cm × 3.5 cm) of the mouse skin and prepare for the histology.
Fix the skin by submerging it in 4% paraformaldehyde for 24 h at 4 °C. Then, cut the skin into 1 cm × 1 cm pieces using scissors and tweezers.
Note: Adequate fixing of the tissues is essential to avoid stiff and brittle specimens because of dehydration and tissue processing steps. Several factors (quality of buffer, penetration into tissues, volume, temperature, concentration, and time) can affect this fixation step.
Dehydrate the skin tissues in different concentrations of alcohol for 12 h using Support Protocol 2.
Embed the dehydrated skin in paraffin using 1 cm × 1 cm skin pieces in a plastic paraffin-embedding cassette.
Cut the tissue into 8 mm sections using a tissue microtome at room temperature.
Note: Use longitudinal instead of cross-sectional paraffin sections and then store them at -20 °C before staining.
Stain the skin sections with H&E by using Support Protocol 3.
Note: Use plus-loaded slides (for example, FisherbrandTM SuperfrostTM Plus microscope slides) for electrostatic adherence of tissue sections to the glass without adhesives or protein coatings. This step will avoid the loss of tissue sections from the slides during the staining process.
Support Protocol 2
Dehydrate the skin tissues through graded ethanol baths. Next, skin samples are made transparent in xylene solution, immersed in 75 °C paraffin oil separately, and embedded in the paraffin-embedding cassettes, which are stable for many years. Paraffin wax is a mixture of n-alkanes with a carbon chain length between 20 and 40 that is solid at room temperature but melts at temperatures between 65 °C and 70 °C.
Protocol steps—step annotations
Dehydrate the skin samples as follows:
Dehydrate in 50% ethanol for 60 min.
Dehydrate in 70% ethanol for 120 min.
Dehydrate in 80% ethanol for 120 min.
Dehydrate in 90% ethanol for 90 min.
Dehydrate in 95% ethanol for 90 min.
Dehydrate in 95% ethanol for 30 min.
Dehydrate in 100% ethanol for 40 min.
Dehydrate in 100% ethanol for 30 min.
Rehydrate the skin sections in xylene solution with two changes for 10 min each.
Immerse the skin sections in 75 °C paraffin oil with two changes for two hours each.
Support Protocol 3
H&E staining of skin samples
The H&E stain is one of the primary medical diagnostic stains performed with formalin-fixed skin samples. This stain gives a candid picture of the different cells and structures in the skin. Hematoxylin (a complex of aluminum ions and oxidized hematoxylin) stains nuclei and keratohyalin granules in blue. The nuclear membrane will stain dark blue. The counterstain, eosin, gives a pink/orange color. Eosin stains red blood cells as a dark shade, muscles, and connective tissues as light orange, and the cytoplasm as a faint pink. Hematoxylin and eosin may be diluted in water and ethanol, respectively, as required.
Protocol steps—step annotations
Submerge the skin samples in each solution as described below:
Deparaffinize the skin sections in xylene solution with two changes for 10 min each.
Rehydrate them in absolute ethanol for 5 min.
Rehydrate them in 95% ethanol for 5 min.
Rehydrate them in 85% ethanol for 5 min.
Rehydrate them in 75% ethanol for 5 min.
Wash the slides in flowing water for 3 min.
Stain the slides with Mayer’s hematoxylin solution for 8 min.
Wash the slides in running tap water for 5 min.
Note: Do not stain the sections with excessive hematoxylin; if it happens, then wash the slides with 1% H2O2 in 70% ethanol.
Counterstain the slides using 0.2% eosin in PBS for 2 min.
Dehydrate the slides in 75% ethanol for 5 min.
Dehydrate the slides in absolute ethanol for 5 min.
Treat the slides with xylene solution, two changes for 5 min each.
Mount the slides with a xylene-based mounting medium.
Note: Xylene is toxic (Kandyala et al., 2010), and commercially available alternatives can be used (Falkeholm et al., 2001). Representative examples of H&E-stained skin sections are shown in Figure 7a –7c.
Figure 7. Histological staining of skin and measurement of epidermal thickness. (A) Representative pictures of histology staining of skin from mice applied with PBS, mannan, or imiquimod (IMQ) (n = 5/group). (B) Measured five different places of the epidermis to calculate mean thickness values. BALB/c female mice were used for experiments. Red arrows were used for marked epidermis thickness. Scale bar = 200 μm. The data represent mean ± SEM. ***, p < 0.001.
Support Protocol 4
Measurement of epidermal thickness
Measure the epidermal thickness by light microscopy, which is a reliable and reproducible method. However, sample collection methods and different sample processing procedures for histology, like fixation, dehydration, embedding, and staining, may alter the tissues, leading to inaccurate thickness measurements. Hence, control and diseased skin samples collected and processed in the same way should be measured simultaneously to minimize such issues. Measure the thickness of the epidermis with a computerized light microscope system (Figure 8), which performs systematic area measurements in sections. The images thus obtained can be analyzed using an image analysis software like Image-pro plus.
Figure 8. Measurement of epidermal thickness in mice using a microscope and image analysis software. Example for measuring epidermis thickness of skin in histological images in (A) PBS- or (B) mannan-exposed mice. The red line was used for marking and measuring epidermis thickness. Left column pictures were captured at 20× magnification using an eclipse upright optical microscope. BALB/c female mice were used for experiments. Skin samples were harvested on day 4 from PBS- and mannan-treated groups.
Protocol steps—step annotations
Measure the thickness of the epidermis under an eclipse upright optical microscope using Image-pro plus (Nikon, Japan) in at least five different tissue areas to measure the thickness accurately. Show mean thickness values from an experiment as shown in Figure 7B.
For measuring epidermal thickness, open Image-pro plus 6.0 software and select the line tool.
Draw a line of unit length on the ruler.
Select Analyze → Set Scale. Enter the length of the line drawn in the known distance, the unit of the length in input unit, check the Global checkbox (use this standard format for all pictures) and click OK.
Select the length to measure based on the line in the picture. Choose at least three images from three different mice for mannan-induced psoriasis skin. Choose 10 places to estimate the size of the epidermis. Use similar protocols for PBS and IMQ groups as well. Obtain at least 30 measurements for the epidermis from each group. Analyze the difference between groups using an unpaired t-test.
Press Ctrl + M (Measurement) to display the results in the results window.
Use GraphPad Prism, version 5, to analyze the data.
Use at least five sections from one mouse, and five mice per group, for measurements.
Measure the thickness of skin sections from psoriasis and naïve mice skin sections in parallel.
Blindly take the measurements to avoid experimenter bias, and the thickness values can be expressed as mean ± SEM.
Data analysis
The data were analyzed with GraphPad Prism 5 and are presented as mean ± SEM. Two-tailed unpaired Student’s t-test was used for comparing PASI between mannan- and IMQ-exposed mice. One-way analysis of variance (ANOVA) with Bonferroni or Newman–Keuls correction was used to compare epidermis thickness between PBS, mannan, and IMQ groups. Probability values < 0.05 were considered significant at a 95% confidence interval.
Validation of protocol
The severity of mannan-induced skin inflammation was much higher than imiquimod-induced psoriasis-like skin inflammation (Wu et al., 2023), and female mice developed more severe psoriasis symptoms than males (Wu et al., 2022). Neutrophil cytosol factor (Ncf1), which encodes p47phox, is critically involved in NADPH oxidase 2 (NOX2) complex formation. The NOX2 complex is responsible for the induction of reactive oxygen species (ROS) and is an essential regulator of several autoimmune diseases. Epicutaneous exposure to mannan induced a severe, aggravated disease in mice having either spontaneous Ncf1m1J mutation or Ncf90H mice having a major human single nucleotide variant on the Ncf1 gene, causing an amino acid replacement from arginine to histidine at position 90 (Li et al., 2023).
General notes and troubleshooting
General notes
The usage of a digital vernier caliper for measuring mouse skin thickness of psoriasis skin:
Gently pinch a small piece of skin with your left hand, hold the electronic vernier caliper with your right hand, and press the skin vertically downwards into the outer measuring claw. Gently slide the movable end of the outer measuring claw to make it tightly grip the skin and avoid damaging it. Ensure that each mouse’s skin thickness is measured and calculated similarly. The skin thickness of the PBS group can be used as a negative control.
The usage of isoflurane in anesthetizing experimental mice:
Place three cotton balls into a 50 mL centrifuge tube and soak the cotton balls with isoflurane. Place the mouse head into the centrifuge tube for 30 s when anesthetizing. Moreover, note that the time should not exceed 60 s. Alternatively, a commercially available isoflurane anesthetizing machine can be used.
Note: Phenobarbital, a barbituric acid derivative, is a long-acting sedative and an anticonvulsant (Suddock et al., 2023). It acts as a non-selective central nervous system depressant. It promotes binding to inhibitory gamma-aminobutyric acid (GABA) subtype receptors and modulates chloride currents through receptor channels. It also inhibits glutamate-induced depolarizations. Whereas isoflurane is an inhaled, short-acting general anesthetic used in surgery (Miller et al., 2023). It is used for induction and maintenance of general anesthesia. It induces muscle relaxation and reduces pain sensitivity by altering tissue excitability. Isoflurane decreases the extent of gap junction-mediated cell-cell coupling and alters the activity of the channels that underlie the action potential. It also activates calcium-dependent ATPase in the sarcoplasmic reticulum by increasing the fluidity of the lipid membrane. Isoflurane binds to the GABA receptor, the large conductance Ca2+-activated potassium channel, and the glutamate and glycine receptors.
Skin paraffin-embedding method for H&E staining.
1.0 cm × 1.0 cm skin is vertically embedded in a paraffin-embedding cassette after dehydration. Pour liquid paraffin at 70 °C into the cassette and place it on ice for cooling. Note that the skin must be placed vertically in the embedding cassette.
Troubleshooting (Table 4)
Table 4. Troubleshooting
Possible questions Solutions
Psoriasis does not develop after mannan application. • Make sure that 100 mg/mL mannan-PBS solutions are stored in a -20 °C freezer.
• Deeply anesthetize mice with phenobarbital for 40 min before mannan-IFA mixture applications.
• Make sure that all mice for experiments are at least 8 weeks old.
Mannan-PBS solution and IFA are unable to mix evenly. • Take mannan solution into 1.5 mL tubes at first, and then add IFA gently. Use 1 mL syringes to draw mannan solution and mix it with IFA for at least 20 times.
Mice dead after IMQ application.
• Make sure that all mice for experiments are at least 8 weeks old.
• Weigh the IMQ cream accurately and no more than 62.5 mg/mice per day should be applied.
Severe weight loss of mice after IMQ application. • Replenish water and food at regular intervals, and provide edible nuts, if necessary, on bedding material in the cages to avoid excess stress to the animals.
Wounds in the skin due to shaving before the experiments. • Use a small, serrated shaver available commercially.
Fighting between animals during experimental period.
• The number of mice kept per cage must follow ethical principles and should not exceed the maximum allowed based on cage space available per mouse.
• Generally, do not mix mice from different breeding cages after weaning them. So, the number of mice per cage must be calculated and housed before the weaning stage.
• Do not place male mice in cages used for female mice, even while changing the mice to new cages during experiments or the scoring phase. Male mice get stressed if exposed to materials used for placing female mice and will start fighting, creating wounds in the skin and tail areas.
• Fighting between female mice might be due to puberty-related issues and maneuvering for dominance. Since mice can live in groups a pecking order is normally established. For the most part female mice usually live peacefully. Lack of enrichment, such as enough places to hide, exercise wheels, and toys can be aggravating factors.
• If female mice are fighting, transfer them to a new cage together because the aggression is, in general, based on the familiar smells the mice have that remind them of their territory and where they fit within the group.
Acknowledgments
We thank Dr. Ia Khmaladze (second Ph.D. student of KSN who graduated from Karolinska Institute, Stockholm, Sweden) and worked with mannan induced psoriasis model earlier for active scientific discussions. We acknowledge Southern Medical University for high-level talent introduction project grants (C1051004, C1034211) given to KSN.
Competing interests
The authors declare no competing financial interests.
Ethical considerations
All animal experiments were performed following the guidelines of the National Institutes of Health (NIH Publication No. 8023) and approved by the ethics committee of Southern Medical University (l2018183). Mice were placed in cages in a climate-controlled environment having 12 h light/dark cycles. Southern Medical University Animal Care and Use Committee, Guangzhou, China, approved all the procedures.
References
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Doxycycline-inducible Expression of Proteins at Near-endogenous Levels in Mammalian Cells Using the Sleeping Beauty Transposon System
KZ Karolina Zak
CA Costin N. Antonescu
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4846 Views: 923
Reviewed by: Gal HaimovichKeisuke Tabata Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Mar 2022
Abstract
The function of a protein within a cell critically depends on its interaction with other proteins as well as its subcellular localization. The expression of mutants of a particular protein that have selective perturbation of specific protein interaction motifs is a very useful strategy for resolving a protein’s mechanism of action in a cellular process. In addition, expression of fluorescent protein fusions is a key strategy for determining the subcellular localization of a protein. These strategies require tight regulation to avoid potential alterations in protein interactions or localizations that can result from protein overexpression. Previous work led to the development of a Sleeping Beauty transposon system that allows doxycycline-inducible expression of protein mutants or fusions; titration of doxycycline allows expression of protein fusions or mutants at near endogenous levels. When used in combination with siRNA gene silencing, this strategy allows for knockdown-rescue experiments to assess the function of specific protein mutants. In this protocol, we describe the use of this Sleeping Beauty strategy for expression of eGFP fusion or mutant proteins in ARPE-19 and MDA-MB-231 cells. This includes design of expression plasmids, transfection, and selection to obtain stable engineered cells, as well as doxycycline treatment for controlled induction of protein expression, either alone or in combination with siRNA silencing for knockdown-rescue experiments. This strategy is advantageous as it allows rapid generation of stable cells for controlled protein expression, suitable for functional studies that require knockdown-rescue as well as various forms of live cell fluorescence imaging.
Key features
• Highly versatile doxycycline-inducible expression system that can be used in various mammalian cell lines.
• Stable integration of transgene allows for sustained and stable expression.
• Titration of doxycycline levels allows expression of transgene at near endogenous levels.
Keywords: Stable cells Doxycycline inducible Mammalian cell lines Sleeping beauty transposon system Fluorescent protein fusions
Background
Cell biology studies often require the expression of exogenous or modified proteins (through the use of transgenes) in cells in culture or other model systems, including expression of specific mutants that each selectively perturb a single aspect of that protein’s function. In addition, the study of protein localization often requires expression of the protein of interest fused to a fluorescent protein such as eGFP (Snapp, 2005) or to an enzyme suitable for proximity biotinylation (Gingras et al., 2019). Hence, optimal strategies for expression of transgenes are essential for a wide range of cell biology approaches.
Transient transfection to introduce plasmids encoding these various transgenes into mammalian cells in culture are commonly used for this purpose. However, this strategy can lead to artificial localization or function of the expressed proteins, as the level of expression of the exogenous proteins using this approach often dramatically exceeds that of the endogenous. Furthermore, transient transfection often leads to expression restricted to a minor fraction of cells (5%–20%), which limits the use of this method for cell population–based assays. In contrast, retroviral and lentiviral systems allow for efficient integration of a cassette for expression of a transgene into the genome. When combined with a selection stage, this approach allows for expression of the transgene in a large majority of cells in a population, as well as relatively stable expression levels of the transgene over time. However, the utility of lentiviral or retroviral strategies may be limited by size constraints for the transgene imposed by the packaging process, the time required to generate viral particles, as well as the need for specific biosafety protocols for work with these virus-like particles.
The modification of endogenous genes with CRISPR/Cas9 has also emerged as a compelling strategy to allow expression of specific mutants or fusions of a specific gene from its endogenous promoter, thus avoiding artifacts of protein overexpression (Bukhari and Müller, 2019). While powerful, this strategy also has potential limitations. Generation of loss-of-function mutants of a particular gene using CRISPR/Cas9 involves passaging of these modified cells and may lead to cellular adaptation that may limit the study of the direct effects of the protein being studied. Furthermore, many genes are present in multiple copies in many cultured cell lines, in particular aneuploid cell lines such as those derived from tumors (e.g., the breast cancer cell line MDA-MB-231 used here). Hence, it can be challenging to use CRISPR/Cas9 to modify all copies of a gene within the genome of these cells in culture, limiting the use of this approach for expression of loss-of-function mutants.
The use of transposons for stable integration of transgenes into the genome of cultured cells is an effective strategy that parallels the use of viral transduction strategies, without the need for packaging of transgene material into viral particles. The Sleeping Beauty transposon method enables stable integration of a transgene found within a vector containing inverted terminal repeats (ITRs) when co-transfected into cells with a vector encoding a transposase enzyme. This approach can be used in combination with selection to achieve integration of the transgene into a large majority of cells in a population (Kowarz et al., 2015).
While the Sleeping Beauty system has been used extensively, a re-engineered transposase enzyme with enhanced activity—SB100X (Mátés et al., 2009)—is ideally suited for this approach, and achieves highly efficient and random integration into TA dinucleotide sites within the target cell genome (30,000 such sites in the human genome) (Kowarz et al., 2015). Kowarz & col. developed a suite of plasmids for use of the Sleeping Beauty transposon system that includes plasmids to allow either inducible (pSBtet) or constitutive (e.g., pSBbi) expression and a range of fluorescent protein and antibiotic selection markers and demonstrated the use of these strategies for stable gene expression in HeLa and HEK293T cells (Kowarz et al., 2015). The pSBtet series of plasmids developed by Kowarz & col. allows doxycycline-inducible expression of a transgene alongside constitutive expression of one of several selection markers (puromycin, hygromycin, neomycin, or blasticidin) and a fluorescent protein (BFP, eGFP, or RFP). We focus here on the use of pSBtet-BP (constitutive expression of BFP and puromycin selection, Figure 1A). Notably, the selection marker, fluorescent protein, and reverse tetracycline transactivator (rtTA) in the pSBtet plasmids are expressed as a self-cleaving polyprotein (Figure 1A).
Figure 1. pSBtet-BP plasmid and cloning strategy. (A) Map of the pSBtet-BP plasmid as obtained from Addgene, plasmid number 60496 and developed by Kowarz & col. (Kowarz et al., 2015). Shown are the Tetracycline responsive element (TRE) promoter, the open reading frame encoding luciferase (which is replaced by the transgene of interest during the cloning stage, Procedure A), as well as the expression of a polyprotein from the constitutive synthetic RPBSA promoter; this polyprotein undergoes cleavage to generate blue fluorescent protein (BFP), reverse tetracycline transactivator (rtTA), and puromycin resistance protein. Also shown is the expression of ampicillin resistance for bacterial selection. Also shown (dashed line boxes) are the regions relevant to subcloning of the transgene, as described in Procedure A. (B) Shown is the cloning strategy for excision of the luciferase-encoding sequence and subcloning of the transgene of interest using NcoI and ClaI sites within the pSBtet-BP plasmid. NcoI and ClaI cut sites are shown in red text; the Kozak sequence is shown in yellow and the open reading frame (ORF) of the luciferase is underlined.
We recently used the Sleeping Beauty transposon strategy developed by Kowarz & col. (Kowarz et al., 2015) in ARPE-19, MDA-MB-231, and SUM149-PT cells for generation of stable cells that allow doxycycline-inducible expression of protein fusions or mutants (Cabral-Dias et al., 2022; Rahmani et al., 2023; Sugiyama et al., 2023) or short hairpin RNA (shRNA) (Lo et al., 2023). Here, we first describe subcloning of a transgene of interest into the pSBtet plasmid (Procedure A). We also note that the use of these Sleeping Beauty plasmids requires an initial transfection and selection process that is unique to some cell lines. We thus report the protocol optimized for the generation of stable cells in ARPE-19 and MDA-MB-231 (Procedure B). Finally, the use of doxycycline for expression of a transgene by the Sleeping Beauty strategy can allow for control of expression to approximate the endogenous level of expression of the protein. This is achieved through titration of doxycycline, allowing determination of an optimal range of doxycycline for each stable cell line generated. This approach allows expression of the transgene at near-endogenous levels, either alone or in combination with siRNA gene silencing of the endogenous protein for knockdown-rescue approaches (Cabral-Dias et al., 2022; Rahmani et al., 2023; Sugiyama et al., 2023). Thus, we also describe the use of doxycycline to induce expression of the protein of interest, either alone to determine optimal doxycycline condition for expression of the transgene (Procedure C), or in combination with siRNA gene silencing of the endogenous protein for knockdown-rescue approaches (Procedure D).
Materials and reagents
Restriction enzymes, e.g., NcoI and ClaI (New England Biolabs, catalog numbers: R0193L and R0197L, respectively)
Competent DH5α E. coli (Thermo Fisher, catalog number: 18265017)
D-MEM/F-12 (1×), liquid 1:1, with L-glutamine and HEPES buffer, 500 mL (Gibco, catalog number: 11330032)
RPMI-1640 medium, liquid; with L-glutamine and sodium bicarbonate, 500 mL (Sigma-Aldrich, catalog number: R8758-500ML)
Tissue culture flasks, 75 cm sq (250 mL, vented, polystyrene) (Sarstedt, catalog number: 83.3911.002)
6-well tissue culture plates, sterile, polystyrene (Sarstedt, catalog number: 86.1254.001)
Fetal bovine serum (Canada), 500 mL (Thermo Fisher Scientific, catalog number: 12483020)
Tetracycline-Free FBS (Wisent, catalog number: 081-150)
Penicillin-Streptomycin liquid 100 mL (Gibco, catalog number: 15070063)
Trypsin-EDTA (0.25% Trypsin with EDTA 1×, 500 mL (Gibco, catalog number: 25200072)
Dulbecco’s phosphate-buffered saline (dPBS), without calcium chloride and magnesium chloride, liquid, 500 mL (Sigma-Aldrich, catalog number: D8537-500ML)
Multiple PCR microtube, 0.5 mL (thin wall, neutral) (Sarstedt, catalog number: 72.735.002)
Ampicillin, sodium salt, 25 g (BioShop, catalog number: AMP201.25)
Chloramphenicol, 25 g (BioShop, catalog number: CLR201.25)
LB Broth (Miller), liquid microbial growth medium (Sigma-Aldrich, catalog number: L2542-500ML)
NucleoBond Xtra Midi EF, Midi kit for endotoxin-free plasmid DNA (Macherey-Nagel, catalog number: 740420.10)
FuGENE HD Transfection Reagent, 1 mL (Promega, catalog number: E2311)
Opti-MEM reduced serum medium (Thermo Fisher Scientific, catalog number: 31985070)
Puromycin dihydrochloride (Sigma-Aldrich, catalog number: P7255-25MG)
pCMV(CAT)T7-SB100 plasmid (Addgene, Plasmid #34879)
pSBtet-BP plasmid (Addgene, Plasmid #60496)
Note: Various other plasmids described in Kowarz et al. (2015) allow alternatives for generation of stable cells to express the transgene using the Sleeping Beauty transposon system.
siRNA in RNAse-free water reconstituted at 20 μM
Lipofectamine RNAiMAX Transfection Reagent (Thermo Fisher Scientific, catalog number: 13778150)
MDA-MB-231 cells (ATCC, catalog number: HTB-26)
ARPE-19 cells (ATCC, catalog number: CRL-2302)
ARPE-19 growth medium (see Recipes)
Tet-free ARPE-19 growth medium (see Recipes)
MDA-MB-231 growth medium (see Recipes)
Tet-free MDA-MB-231 growth medium (see Recipes)
Equipment
Shaking 37 °C platform incubator (Thermo Fisher Scientific, model: SHKE6000)
Cell incubator set to 37 °C and 5% CO2 (Thermo Fisher Scientific, model: HERAcell 150i)
Centrifuge (unrefrigerated) for 50 mL Falcon tubes (Thermo Fisher Scientific, model: Sorval ST16R)
Procedure
Plasmid design and isolation
Subclone transgene of interest into the pSBtet-BP plasmid, using standard molecular biology techniques. We suggest synthesizing the gene of interest using commercial services for subcloning into NcoI (base pair 684) and ClaI (base pair 2361) within the pSBtet-BP plasmid (see Figure 1B). This strategy will result in excision of the open reading frame for luciferase, so that it can be replaced with the transgene of interest. Please note that it is important to maintain the integrity of the Kozak sequence within the final plasmid (Figure 1B, sequence with yellow shading). It is also important to ensure that the synthesized transgene does not contain internal NcoI and ClaI cleavage sequences; these should be removed by silent point mutations if present. Alternatively, SfiI sites can be used to subclone a gene of interest within the pSBtet-BP plasmid (Kowarz et al., 2015).
Isolate plasmid using midi-prep or maxi-prep procedure.
Inoculate 100 mL of LB broth culture supplemented with 100 μg/mL of ampicillin with a single clone of competent DH5α E. coli transformed with the pSBtet-BP transgene plasmid.
Inoculate 100 mL of LB broth culture supplemented with 25 μg/mL of chloramphenicol with a single clone of competent DH5α E. coli transformed with the pCMV(CAT)T7-SB100 plasmid.
Grow bacterial liquid cultures in a shaking 37 °C incubator overnight.
Perform plasmid isolation using a commercial kit available for this purpose, ensuring that these are certified to be endotoxin-free. We suggest using NucleoBond Xtra Midi EF, and several other alternatives are available. This procedure is described as per the manufacturer’s instructions.
Transfection of Sleeping Beauty plasmids and puromycin selection
Thaw fresh batch of ARPE-19 cells and passage twice (see Note 1). The protocol for growth and one passage of cells (see Note 3) is as follows:
Grow cells in a T-75 flask in ARPE-19 growth medium at 37 °C and 5% CO2 (see Recipes below for medium composition).
When cells reach 60%–80% confluence, remove medium and wash with dPBS.
Incubate cells in 1.5 mL of trypsin solution for 3–5 min at 37 °C or until cells detach from the surface of the T-75 flask.
Add 8.5 mL of ARPE-19 growth medium and use a pipette to wash medium solution over the surface of the flask to ensure that all cells are detached.
Centrifuge cells at 200× g for 5 min and discard the supernatant.
Carefully resuspend the pellet into 10 mL of fresh ARPE-19 growth medium.
Transfer 2–3 mL of this cell suspension to a fresh T-75 flask, to which add an additional 7–8 mL of ARPE-19 growth medium.
The remaining 7–8 mL of cell suspension from the previous step can be used for seeding (e.g., step B2 below) or discarded.
Seed ARPE-19 cells into wells of a 6-well plate. Seeding should be done at approximately 5.0 × 105 cells per well (in 2 mL of medium per well), leading to ~20%–30% confluence at time of seeding and 40%–50% confluence on the day of transfection (step B3).
Transfect cells with plasmids using FuGENE HD transfection reagent. Each plasmid transfection condition should be performed in triplicate (3 wells of the 6-well plate per condition).
Prepare the following transfection mixture in a sterile microfuge tube per well of a 6-well plate to be transfected (see Note 4), in the following order:
i. Opti-MEM medium for a total volume of 150 μL
ii. 3 μg of pSBtet-BP transgene plasmid
iii. 0.3 μg of the pCMV(CAT)T7-SB100 plasmid
iv. 9 μL of FuGENE transfection reagent
Vortex for 30 s and incubate this transfection mixture for 15 min at room temperature.
Remove growth medium from cells and wash once with 2 mL of dPBS. After removal of dPBS, add 1.9 mL of Opti-MEM medium to each well. Then, add 100 μL of transfection mixture (from step B3) to each well, dropwise.
Incubate cells with transfection mixture in Opti-MEM for 24 h. Then, remove this medium, wash cells once in dPBS, and replace medium with ARPE-19 growth medium. Incubate cells for another 24 h at 37 °C with 5% CO2.
Replace medium with ARPE-19 growth medium and incubate for 24 h at 37 °C with 5% CO2. This step is essential to allow cells to recover and maintain viability following transfection.
After 24 h, replace regular growth medium with ARPE-19 growth medium supplemented with 2 μg/mL of puromycin and incubate cells for 2–3 days at 37 °C with 5% CO2. Some cell death can be observed during this period.
When cells in the 6-well plate reach ~50%–60% confluence, passage cells into a T-25 flask, as follows:
Remove medium from cells and wash 2× in dPBS.
Add 0.3 mL of Trypsin to each cell and incubate for 5 min at 37 °C and 5% CO2.
When cells begin to detach, add 2 mL of ARPE-19 growth medium, ensuring that cells are detached from the bottom of the well.
Combine the cell suspension mixture from each well of triplicate transfection condition into a single tube.
Centrifuge cells at 200× g for 5 min and discard the supernatant.
Carefully resuspend the pellet in 5 mL of fresh ARPE-19 growth medium.
Transfer cell suspension into a single T-25 flask and incubate for 24 h at 37 °C with 5% CO2.
Replace medium with ARPE-19 growth medium supplemented with 2 μg/mL of puromycin and incubate at 37 °C with 5% CO2, while changing medium supplemented with puromycin every 2–3 days.
When cells reach confluence, passage cells as in step B1 and transfer into a T-75 flask.
When cells reach confluence again, passage cells again as in step B1. Ensure that cells have been passaged approximately 3–4 times over the course of 2–3 weeks with puromycin selection medium. If expression of multiple transgenes using this strategy is required, please see Note 5.
Doxycycline induction of transgene expression—transgene expression alone
Culture ARPE-19 cells stably transfected with pSBtet-BP transgene (produced following Procedure B, called ARPE-19-transgene cells henceforth) in tet-free ARPE-19 growth medium. This should require 2–3 passages (approximately 7–8 days) of culture in this medium, as per step B1 above. This stage ensures that there is minimal expression of the transgene prior to silencing the expression of the endogenous protein.
Seed ARPE-19-transgene cells into wells of a 6-well plate. Seeding should be done at approximately 5 × 105 cells per well (in 2 mL of medium per well), leading to ~20%–30% confluence at time of seeding. For initial experiments, seeding ~6 wells worth of cells is recommended, as this will allow testing of a range of doxycycline concentrations.
At 24 h post-seeding, replace medium with tet-free ARPE-19 growth medium supplemented with 0–2,000 ng/mL of doxycycline (e.g., 0, 50, 100, 250, 500, and 2,000 ng/mL doxycycline) and incubate for 24 h at 37 °C with 5% CO2 before proceeding to downstream experimental applications such as fluorescence microscopy or western blotting experiments. Examples of this approach, with detection of fluorescent protein fusion expression by western blotting, are seen in Figure 2A, and by fluorescence microscopy in Figure 2B. Detection of transgene expression by western blotting allows comparison of the expression of the transgene to that of the endogenous protein (Figure 2A, compare expression of NCK1-eGFP to that of endogenous NCK1), while detection of transgene expression by microscopy allows estimation of the proportion of cells that harbor the transgene. Additional examples of detection by western blotting are found in Cabral-Dias et al. (2022), Figure S2B and S2C (see Note 2).
Figure 2. Detection of NCK1/2 transgenes in ARPE-19 cells stably engineered with pSBtet-BP-transgene plasmids. ARPE-19 cells were subjected to transfection to generate cells that stably carry the transgene for inducible expression of eGFP-tagged NCK1 or 2, as in Procedure B. Then, induction of NCK1-eGFP (A) or NCK2 -eGFP (B) was performed as in Procedure C, using doxycycline concentrations (ng/mL) as indicated for 24 h. (A) Whole-cell lysates of NCK1-eGFP stable cells were resolved by SDS-PAGE followed by transfer to PVDF membranes and then immunoblotted using antibodies as indicated. Also shown are the approximate molecular weights (kDa) for each blot panel. (B) Cells were subjected to PFA fixation, and DAPI labeling using standard fluorescence microscopy sample preparation, and then imaged using a Quorum Diskovery instrument comprised of a Leica DMi8 microscope. Imaging was done using a 40× objective with 405- and 488-nm laser illumination, 450/55 and 525/50 emission filters, and acquired using a Zyla 4.2Plus sCMOS camera (Andor). Scale bars: 40 μm.
Doxycycline induction of the transgene for knockdown-rescue experiments
Culture ARPE-19 cells stably transfected with pSBtet-BP transgene (as per Procedure B, called ARPE-19-transgene cells henceforth) in tet-free ARPE-19 growth medium. This should require 2–3 passages (approximately 7–8 days) of culture in this medium, as per step B1 above. This stage ensures that there is minimal expression of the transgene prior to silencing the expression of the endogenous protein. Please see Note 6 when designing plasmids for knockdown-rescue approaches.
Seed ARPE-19-transgene cells into wells of a 6-well plate. Seeding should be done at approximately 2.5 × 105 cells per well (in 2 mL of medium per well), leading to ~10%–15% confluence at time of seeding and 20%–30% confluence on the day of transfection (first transfection is 24 h after seeding).
Transfect cells with target siRNA using Lipofectamine RNAiMAX reagent (transfection is performed twice, separated by 24 h). Time (in hours) is relative to the time of seeding at t = 0 h, with transfections performed at t = 24 h and 48 h, as follows:
Prepare the following transfection mixture in a sterile microfuge tube for each well of a 6-well plate to be transfected:
i. 100 μL of Opti-MEM medium
ii. 3.25 μL of Lipofectamine RNAiMAX reagent
iii. 2.75 μL of 20 μM siRNA solution
Vortex for 10 s and incubate this transfection mixture for 15 min.
Wash ARPE-19 cells in a 6-well plate 2–3 times with dPBS and remove medium.
Add 900 μL of Opti-MEM to each well. Then, add 100 μL of transfection mixture to each well in a dropwise motion.
Let cells incubate in the incubator for 3 h at 37 °C with 5% CO2.
Wash cells with PBS after 3 h and replace with tet-free ARPE-19 growth medium.
Allow the cells to rest while incubating in tet-free ARPE-19 growth medium for 24 h at 37 °C with 5% CO2 and repeat this protocol (steps D3a–D3f) the next day, followed by another 24 h rest (incubation in tet-free ARPE-19 growth medium at 37 °C with 5% CO2) before proceeding to doxycycline induction.
At 72 h post-seeding (24 h after the last siRNA transfection), replace medium with tet-free ARPE-19 growth medium supplemented with 200 ng/mL of doxycycline to each well (or another optimal, experimentally determined doxycycline concentration) and incubate for 24 h at 37 °C with 5% CO2 before proceeding to downstream experimental applications such as immunofluorescence microscopy or western blotting experiments. Examples of this approach for immunofluorescence microscopy are found in (Cabral-Dias et al., 2022), Figures 4 and 5.
Notes
Ensure use of freshly thawed, low passage number cells when making stable cell lines (Procedure B).
The determination of optimal doxycycline conditions for induction will be specific to each experiment and protein being studied. Expression within ~2–3 fold of that of the endogenous protein, for example as determined by western blotting, is a reasonable initial condition to test for downstream applications; however, the appropriate expression level should be tested for each protein.
The protocol described here is well suited for ARPE-19 cells. The same protocol is also effective for other cell lines such as MDA-MB-231 cells, while noting differences in MDA-MB-231 growth medium and tet-free MDA-MB-231 growth medium (see Recipes).
The ratio of pSBtet-BP plasmids containing the transgene of interest and the transposase enzyme (pCMV(CAT)T7-SB100) used for transfection (step B3) may require optimization for each plasmid. We report transfection of a 10:1 ratio of pSBtet-BP transgene plasmid to pCMV(CAT)T7-SB100 integrase plasmid; other proportions may also be considered if stable transfection is not initially successful. The 10:1 ratio of pSBtet-BP transgene plasmid to the SB100 integrase plasmid is optimized to enhance the likelihood of a single integration event per cell; however, further reduction of the amount of pCMV(CAT)T7-SB100 integrase plasmid may be considered should further optimization be required to achieve a single integration event per cell (Grabundzija et al., 2010; Kowarz et al., 2015).
It is expected that the pCMV(CAT)T7-SB100 integrase plasmid is not retained following the selection process, as the latter takes two weeks or longer, and this integrase-expressing plasmid is not itself integrated into the host cell genome. Should an experimental design require expression of a second transgene using an additional pSBtet or pSBbi plasmid (e.g., using a different selection marker to express a different transgene), this will again require co-transfection of the pCMV(CAT)T7-SB100 alongside the appropriate pSBtet/pSBbi plasmid.
To allow for knockdown-rescue experiments, it is essential that the transgene used to generate the pSBtet plasmid also has a silent mutation in the region of the gene corresponding to the siRNA sequence.
Recipes
ARPE-19 growth medium
DMEM/F12 medium (500 mL)
10% FBS (50 mL)
Penicillin-Streptomycin (5,000 U/mL) (5 mL)
Tet-free ARPE-19 growth medium
DMEM/F12 medium (500 mL)
10% tetracycline-free FBS (50 mL)
Penicillin-Streptomycin (5,000 U/mL) (5 mL)
MDA-MB-231 growth medium
RPMI-1640 medium, liquid; with L-glutamine and sodium bicarbonate (500 mL)
10% FBS (50 mL)
Penicillin-Streptomycin (5,000 U/mL) (5 mL)
Tet-free MDA-MB-231 growth medium
RPMI-1640 medium, liquid; with L-glutamine and sodium bicarbonate (500 mL)
10% tetracycline-free FBS (50 mL)
Penicillin-Streptomycin (5,000 U/mL) (5 mL)
Acknowledgments
This work was supported by a Project Grant from the Canadian Institutes of Health Research (PJT156355) to C.N.A. We thank Laura A. Orofiamma for the critical reading of this manuscript. The Sleeping Beauty plasmids described here were developed by Kowarz & colleagues (Kowarz et al., 2015). We have previously used this method to develop stable cell lines in ARPE-19, MDA-MB-231, and SUM149-PT cells (Cabral-Dias et al., 2022; Lo et al., 2023; Rahmani et al., 2023; Sugiyama et al., 2023).
References
Bukhari, H. and Müller, T. (2019). Endogenous Fluorescence Tagging by CRISPR. Trends Cell Biol. 29(11): 912–928.
Cabral-Dias, R., Lucarelli, S., Zak, K., Rahmani, S., Judge, G., Abousawan, J., DiGiovanni, L. F., Vural, D., Anderson, K. E., Sugiyama, M. G., et al. (2022). Fyn and TOM1L1 are recruited to clathrin-coated pits and regulate Akt signaling. J. Cell Biol. 221(4): e201808181.
Gingras, A. C., Abe, K. T. and Raught, B. (2019). Getting to know the neighborhood: using proximity-dependent biotinylation to characterize protein complexes and map organelles. Curr. Opin. Chem. Biol. 48: 44–54.
Grabundzija, I., Irgang, M., Mátés, L., Belay, E., Matrai, J., Gogol-Döring, A., Kawakami, K., Chen, W., Ruiz, P., Chuah, M. K., et al. (2010). Comparative Analysis of Transposable Element Vector Systems in Human Cells. Mol. Ther. 18(6): 1200–1209.
Kowarz, E., Löscher, D. and Marschalek, R. (2015). Optimized Sleeping Beauty transposons rapidly generate stable transgenic cell lines. Biotechnol. J. 10(4): 647–653.
Lo, L., Uchenunu, O., Botelho, R. J., Antonescu, C. N. and Karshafian, R. (2023). AMPK is required for recovery from metabolic stress induced by ultrasound microbubble treatment. iScience 26(2): 105883.
Mátés, L., Chuah, M. K. L., Belay, E., Jerchow, B., Manoj, N., Acosta-Sanchez, A., Grzela, D. P., Schmitt, A., Becker, K., Matrai, J., et al. (2009). Molecular evolution of a novel hyperactive Sleeping Beauty transposase enables robust stable gene transfer in vertebrates. Nat. Genet. 41(6): 753–761.
Rahmani, S., Ahmed, H., Ibazebo, O., Fussner-Dupas, E., Wakarchuk, W. W. and Antonescu, C. N. (2023). O-GlcNAc transferase modulates the cellular endocytosis machinery by controlling the formation of clathrin-coated pits. J. Biol. Chem. 299(3): 102963.
Snapp, E. (2005). Design and Use of Fluorescent Fusion Proteins in Cell Biology. Curr. Protoc. Cell Biol. 27(1): ecb2104s27.
Sugiyama, M. G., Brown, A. I., Vega-Lugo, J., Borges, J. P., Scott, A. M., Jaqaman, K., Fairn, G. D. and Antonescu, C. N. (2023). Confinement of unliganded EGFR by tetraspanin nanodomains gates EGFR ligand binding and signaling. Nat. Commun. 14(1): 2681.
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4,847 | https://bio-protocol.org/en/bpdetail?id=4847&type=0 | # Bio-Protocol Content
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A Protocol to Retrieve and Curate Spatial and Climatic Data from Online Biodiversity Databases Using R
MC Marina Coca-de-la-Iglesia
VV Virginia Valcárcel
NM Nagore G. Medina
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4847 Views: 765
Reviewed by: Prashanth N SuravajhalaJeff W Bizzaro Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in American Journal of Botany Aug 2022
Abstract
Ecological and evolutionary studies often require high quality biodiversity data. This information is readily available through the many online databases that have compiled biodiversity data from herbaria, museums, and human observations. However, the process of preparing this information for analysis is complex and time consuming. In this study, we have developed a protocol in R language to process spatial data (download, merge, clean, and correct) and extract climatic data, using some genera of the ginseng family (Araliaceae) as an example. The protocol provides an automated way to process spatial and climatic data for numerous taxa independently and from multiple online databases. The script uses GBIF, BIEN, and WorldClim as the online data sources, but can be easily adapted to include other online databases. The script also uses genera as the sampling unit but provides a way to use species as the target. The cleaning process includes a filter to remove occurrences outside the natural range of the taxa, gardens, and other human environments, as well as erroneous locations and aspatial correction for misplaced occurrences (i.e., occurrences within a distance buffer from the coastal boundary). Additionally, each step of the protocol can be run independently. Thus, the protocol can begin with data cleaning, if the database has already been compiled, or with climatic data extraction, if the database has already been parsed. Each line of the R script is commented so that it can also be run by users with little knowledge of R.
Keywords: Ecology Climate Biodiversity Database Data cleaning R language GBIF BIEN WorldClim
Background
Our knowledge of species distributions is central to biogeographers but also to phylogeneticists and ecologists. Indeed, species distributions are needed to perform phylogenetic climate reconstructions, species niche characterizations, or species distribution models, and to address many evolutionary questions. However, obtaining accurate spatial information on species distributions requires occurrence databases of good quality, with high geographical coverage, which are difficult to obtain.
The main sources of geographical information are field inventories and biodiversity collections (museums and herbaria), for which accessibility has been a serious limitation until recently. The digitization efforts of recent decades have facilitated access to vast amounts of biodiversity data previously scattered in different institutions around the world, through online databases such as the Global Biodiversity Information Facility (GBIF; GBIF.org, 2021). As a result, we now have an unprecedented opportunity to benefit from centuries worth of naturalist observations from all over the world. However, the use of this valuable information is limited by persistent knowledge gaps and technical limitations. On the one hand, our knowledge of species distributions is still poor, biased, or imprecise (Hortal et al., 2007), and this is reflected in the information collected in biodiversity databases, which is not consistent across lineages or across regions. These biases result in some groups of organisms and regions of the world having scarce information while others have large amounts of data (Hortal and Lobo, 2005). On the other hand, the complexity of the process of parsing and preparing online data for analysis is high. For example, it is common for online repositories to contain records with imprecise or erroneous spatial information (such as terrestrial organisms with records in the sea) or with outdated taxonomic nomenclature (Soberón and Peterson, 2004). Therefore, every study based on online data requires a first step of cleaning and parsing to remove or minimize the impact of these sources of uncertainty (persistent knowledge gaps and technical limitations) on further analysis (Hortal et al., 2007).
In parallel with the international digitization efforts of the last decades, several methods and pipelines have been designed to deal with these sources of uncertainty and to simplify the different steps of working with online biodiversity data. Some of the most relevant protocols have been developed in R (R Core Team, 2018) and include geographic, taxonomic, or temporal data cleaning (see for example: Biogeo, Robertson et al., 2016; SpeciesGeoCorder, Töpel et al., 2016; CoordinateCleaner, Zizka et al., 2019; bRacatus, Arlé et al., 2021; BDcleaner, Jin and Yang, 2020; plantR, Lima et al., 2021). However, none of them address the uncertainty introduced by both the spatial knowledge gaps and the technical limitations. Moreover, most of them focus on one or a few steps of the process. Thus, to complete the process (from the initial download of occurrences to the climatic data extraction of the cleaned and parsed spatial database) users have to deal with different protocols, some of which require programming skills or a deep R background.
The R protocol that we present here is designed to produce reliable databases of species occurrences and climate data from online repositories. It provides an automatic procedure for dealing with the most common sources of spatial uncertainty in online biodiversity databases. It also includes an automatic script to run each sample (species, genus, family, etc.) separately, allowing for an easy and fast way to process hierarchical databases. The script also includes a post-processing code to run after the spatial pipeline and extract the climatic data. The protocol describes a step-by-step guide on how to download, parse, clean, and merge spatial and climatic data from three online databases (Figure 1; WorldClim, Fick and Hijmans, 2017; BIEN, Maitner, 2020; GBIF, GBIF.org, 2021). Moreover, the protocol can be easily adapted to include any other online biodiversity database that may be of interest. The cleaning steps include how to automatically update nomenclatural information, identify, and remove records outside the natural distribution of taxa, records from gardens and other human environments, or geographically inaccurate records. To explain the protocol, we used the AsianPalmate Group (AsPG) of Araliaceae as a case study, using genera as a sample unit. To speed up the implementation, we selected 16 of the AsPG genera. The selection of genera was made to exhibit uneven spatial information across genera and across areas of the world. This approach aimed to address the issue derived from knowledge gaps as a source of spatial uncertainty. Additionally, the chosen genera are largely affected by erroneous and misplaced records, serving to tackle the issue arising from technical limitations as a source of spatial uncertainty.
Figure 1. Workflow for the entire protocol pipeline. It includes all the steps and alternatives described in the protocol. Each red arrow represents the steps of the protocol that we can start with, depending on the data available.
In summary, the main advantages of this protocol are that it: (1) can be applied to all groups of organisms (as long as they have information available in GBIF or BIEN databases) and at any taxonomic rank, not only at the species level; (2) provides an automatic way to handle hierarchical databases, which is very helpful when studying highly diversified groups (genera with a high number of species, families with a high number of genera, etc.); (3) provides a complete pipeline from spatial data download (including merging multiple databases) to climatic data extraction; (4) deals with uncertainties arising from technical limitations (such as incorrect records), but also with the uncertainties arising from persistent knowledge gaps (such as spatial biases in different parts of the world and across lineages); (5) provides an easy way to filter out records outside the natural range of the taxa; (6) applies a spatial correction for erroneous occurrences outside the coastal boundary; (7) includes independent steps for each part of the process that can be run separately; and (8) can be easily used and modified by any types of users regardless of their skills, knowledge, or background on R, because it is accompanied by instructions to guide the user.
Equipment
Computer with Microsoft® Windows® 10 education, KUBUNTU 22.04, or Mac® OS X® 12.6 operating systems and versions.
Software
R version 4.3.1 (https://r-project.org/)
Packages: "BIEN", "countrycode", "data.table", "devtools", "dpyr", "plyr", "raster", "readr", "rgbif", "rgdal", "spocc", "spThin", "SEEG-Oxford/seegSDM" and "tidyr".
RStudio version 2023.06.0+421 (https://rstudio.com/products/rstudio/)
The use of RStudio is optional. RStudio is an interface that improves the use of R.
Databases
POWO (https://powo.science.kew.org)
GBIF (https://www.gbif.org/)
BIEN (https://bien.nceas.ucsb.edu/bien/)
WorldClim (https://www.worldclim.org/)
Procedure
The R script can be freely downloaded from GitHub (https://github.com/NiDEvA/R-protocols; Note 1). The pipeline of the procedure coincides with the steps of the R script of this protocol (Figure 1). First, you need to create two working folders, one called “input” (it contains the information needed to run the R script) and another one called "output" (it will contain the resulting files after running the R script). In this protocol, we used the genus rank as the operational taxonomic unit, but the script also contains commented lines (those preceded with "#") with the functions needed if you want to use species as the sample unit. Besides, it can also be easily modified to use family or any other higher taxonomic level as the operational unit if needed. We have also cleaned the data by removing records outside the natural distribution of genera, from gardens and other human environments, or those that are geographically inaccurate, but it can also be readily adjusted to meet specific data cleaning requirements as needed.
Prepare a checklist of the native range of the taxa
It is necessary to know the countries where the taxa are native. For plants, this information can be found in the World Checklist of Selected Plant Families (WCSP, Govaerts et al., 2008). WCSP is a database that compiles checklists of 200 seed plant families, and it is available in Plants of the World Online (POWO, 2023; http://www.plantsoftheworldonline.org/). The database is frequently updated, and each new name published in the International Plant Name Index (IPNI; International Plant Names Index, 2020) is reviewed and added to POWO. Other sources of information on the natural range of other organisms are available in ASM Mammal Diversity Database (https://www.mammaldiversity.org/index.html, Mammal Diversity Database, 2020), Avibase - The World Bird Database (https://avibase.bsc-eoc.org/avibase.jsp, Lepage et al., 2014), Catalogue of Life ( https://www.catalogueoflife.org/, Bánki et al., 2022), Checklist of Ferns and Lycophytes of the World (Hassler, 2022a), Global Assessment of Reptile Distributions (http://www.gardinitiative.org/, GARD, 2022), Reptile Database (http://www.reptile-database.org/, Uetz et al., 2021), USDA Plants Database (https://plants.usda.gov, USDA, NRCS, 2022), and World Plants (https://www.worldplants.de , Hassler, 2022b). For plants not included in POWO or for animals, go to steps A1–A4 to manually create the list of countries where the taxa are native. For plants included in POWO database, go to step A5 to automatically create the list of native countries.
Create a text file in a text editor (e.g., Notepad++, BBedit, or Notepadqq in Linux), with the names of all taxa separated by "Enter" in a plain text editor. Save it as "Natural_Distribution_Checklist_TDWG.txt".
Go to https://wcsp.science.kew.org/home.do (or the corresponding webpage, see above) and enter the taxon name in the search engine. WCSP uses two ways to describe the distribution of taxa, one in narrative form and the other one through international codes (Figure 2). The international code used in WCSP is the third level of geographical codes of the Taxonomic Databases Working Group (TDWG, Brummitt, 2001) (Note 2).
Figure 2. Example of Brassaiopsis genus in WCSP. WCSP provides status, distribution, family, and original compiler information of each taxon.
Copy each code of three capital letters (just the codes, not the numbers that appear at the beginning of the country code line) and paste in "Natural_Distribution_Checklist_TDWG.txt" in the same line right after the corresponding taxa name separated by ";". In some cases, symbols ["?", "(?)", "+", "†"] or lowercase letters may appear in distribution. According to TDWG, "?" is used when the presence of a taxon in a given area is not certain. If this symbol is used within brackets, it is because there is no exact location known within a country. When a taxon is extinct or may be extinct in an area, the symbol "†" is placed after the country code. When the country code is not known, "+" is used. Lowercase letters for the country code indicate naturalization. For this protocol, we have only used the codes with three capital letters that do not have any symbols. For more information, consult the "about checklist" section on the WCSP website.
Repeat steps A2 and A3 until all taxa are completed and save the document in the "input" folder. The format of the resulting text file should look as in Figure 3. It is advisable to sort the taxa alphabetically in the text file.
Figure 3. Format for the text file containing the natural distribution countries of the taxa. The first element of each row is the name of an AsPG genus followed by the Level-3 TDWG code of the countries where the taxa occur naturally, separated by a semicolon.
Additionally, steps A1–A4 can be automated in R for plants included in POWO. We have provided a working example in the code (see point 5 in the code). However, it is important to note that many families are not included in the POWO database, so in most cases it will be necessary to generate the list manually as indicated in steps A1–A4.
Create an account in GBIF database
Visit the website https://www.gbif.org/. Click on Login located in the upper-right corner of the website, and then on REGISTER (Figure 4).
Figure 4. Home of the GBIF Database. To create an account, it is necessary to fill the required information in the section Register; remember the information included, because it will be necessary later in the R protocol.
Fill the COUNTRY, EMAIL, USERNAME, and PASSWORD fields, click on next, and follow the instructions to create the account. It is also possible to create the account through Google, Facebook, or GitHub. Important to remember: save the information filled in the email, username, and password, because it will be used later in the R script.
Initial preparation in R
Open RStudio (Note 3).
Create the paths in R for the input and output data. These paths correspond to the "input" and "output" folders. Replace the example path with the path of the “input” folder.
Install and load the packages needed to run the R script. This packages are "BIEN" (Maitner et al., 2018), "countrycode" (Arel-Bundock et al., 2018), "data.table" (Dowle et al., 2019), "devtools" (Wickham et al., 2021), "dplyr" (Wickham et al., 2020), "plyr" (Wickham, 2020), "raster" (Hijmans et al., 2020), "readr" (Wickham et al., 2018), "rgbif" (Chamberlain et al., 2020a), "rgdal" (Bivand et al., 2020), "seegSDM" (Golding and Shearer, 2021), "spocc" (Chamberlain et al., 2020b), "spThin" (Aiello-Lammens et al., 2019), and "tidyr" (Wickham et al., 2022). The packages can be downloaded directly into RStudio with the "install.packages" function, except for the "seegSDM" package, which is installed through the "devtools" package as it is specified in L63 of the R script. In this protocol we use BIEN (https://www.biendata.org/) and GBIF (https://www.gbif.org/) as the online biodiversity databases for obtaining the spatial data; if you want to include any other online database to obtain records for your case study, you will need to look for the correspondent R package and install them at this step of the protocol.
Create a vector with the names of the taxa (sample unit). The vector can be created in R (see "Alternative 1" in the R script) or it can be imported from a file (.csv, .txt, .xls; see "Alternative 2" in the R script). For this protocol, we read the file on the GitHub page “Alternative 3”, as we already have the information in GitHub for this example. Be aware that the names and order of sample units must be exactly the same as in the "Natural_Distribution_Checklist_TDWG.txt" file (see above step A1).
Read the natural distribution text file built in procedure A named "Natural_Distribution_Checklist_TDWG.txt". First, load in R the text file that contains the natural distribution of sample units as a vector. Then, convert the vector into a list. Each item in the list corresponds to a genus (or the correspondent sample unit: species, family, etc.) followed by its corresponding Level-3 TDWG country codes, separated by ";".
Download the occurrence data from online databases (GBIF and BIEN, or the desired database)
Use the package "rgbif" to download the records from the GBIF database.
Indicate the username, email, and password created in step B2. Replace the "XX" with your credentials (important to not remove the "characters").
Search the taxon keys of each taxon. GBIF has a key to identify each taxon in the database. In case you use species or families instead of genera as the sample unit, replace rank = "genus" in the name_backbone function with rank = "species" or rank = "family", respectively.
Prepare the download request and download data.
i. Create the path to a folder that will contain the files downloaded from GBIF and create a folder inside the "input" folder, naming it as "download_GBIF". This folder will contain the files downloaded from GBIF. Indicate the number of taxa included in the text file with the checklist of taxa native range.
ii. Prepare and download the occurrences. The download request is executed with the "occ_download" function, which allows you to establish criteria for filtering the data downloaded. In this protocol, as we wanted to keep only native records with coordinates, we used the arguments explained in Table 1. The "occ_download_wait" function indicates the status of the download, and the code continues running until the status changes to “succeeded”. The download is performed with the "occ_download_get" function and will result in as many zip files inside the "download_GBIF" folder as taxa included in your request. The "occ_download_import" function imports into R the data set downloaded from the "download_GBIF" folder (Note 4).
Table 1. Necessary arguments of the occ_download function from “rgbif” package
Argument Description To download
pred Downloads only the occurrences equal to unique condition Select "taxon" for "taxonKey" and TRUE for "hasCoordinate"
pred_not Downloads only the occurrences not equal to the condition Select "INTRODUCED", "INVASIVE", "MANAGED", and "NATURALISED" for "establishmentMeans"
pred_in Downloads the occurrences equal to multiple conditions Select "taxon.keys" for "taxonKey"
Use the BIEN_occurrence_genus function from R package "BIEN" to download the records from the BIEN database version 4.1.1 (Note 5). It is necessary to indicate some arguments to start the download (Table 2). We will refer to the resulting data set as “raw.BIEN.dataset” onwards. If there are no records in “raw.BIEN.dataset”, go directly to step E3 to replace the column names and see Note 6.
Table 2. Necessary arguments of the BIEN_occurrence_genus function from “BIEN” package. If you use species as the sample unit, then you will need to use the BIEN_occurrence_species function; replace the argument "genus" with "species" with the remaining arguments staying the same.
Argument Description For download
genus Name of genus This argument corresponds to the vector of taxa names created in R (taxa.names)
cultivated If TRUE, it also returns cultivated occurrences Select FALSE (it is selected by default)
all.taxonomy If TRUE, it returns all taxonomic information Select TRUE (FALSE is selected by default)
collection.info If TRUE, it returns additional information about collection and identification Select TRUE (FALSE is selected by default)
observation.type If TRUE, it returns information on type of observation Select TRUE (FALSE is selected by default)
political.boundaries If TRUE, it returns information on political boundaries Select TRUE (FALSE is selected by default)
natives.only If TRUE, it returns only native species Select TRUE (is selected by default)
Save the R workspace with the downloaded data as "1_Workspace_Download.RData". It is very useful to save the objects created in the downloaded data. If there is a problem in later steps, this workspace can be loaded and thus avoid making another download.
Unify the format of the downloaded databases and simplify the database by removing unnecessary columns
In order to join the information from the two databases, the number of columns and their names have to be identical in "raw.GBIF.list" and "raw.BIEN.dataset". Note that some columns from GBIF and BIEN have different names and yet contain the same information. In those cases, it is necessary to rename the columns (see below). Columns with information that will not be used in further analysis can be removed in this step too.
Simplify raw GBIF data set "raw.GBIF.list".
Create new columns for future merging between GBIF and BIEN raw data sets.
i. Add the "countryName" column to include full country names using the "countrycode" R package and the "countryCode" column. The "countryCode" column includes a 2-letter ISO 3166-1 standard for country codes and their subdivisions. This standard is used by GBIF to indicate the country in which the occurrence was recorded, while BIEN uses the full name. The “countrycode” function transforms the "countryCode" column in country names (Note 7).
ii. Add “dataOrigin” column filled with GBIF. This column indicates if the record belongs to GBIF or BIEN; in this case, it will be filled with “GBIF” for all records.
Select useful columns to simplify the data set before merging with BIEN. The number of columns of "raw.dataset" is approximately 50; it is advisable to reduce this number. For our purpose, the useful information is inside the following columns: "gbifID", "dataOrigin", "basisOfRecord", "genus", "species", "scientificName", "decimalLongitude", "decimalLatitude", "elevation", "countryName", "countryCode", "locality", "eventDate", "institutionCode", "collectionCode", and "catalogNumber". Therefore, we will keep only these columns. We will refer to the resulting data set as "simple.GBIF.dataset" onwards.
Export "simple.GBIF.dataset" as a csv file.
Simplify raw BIEN data set "raw.BIEN.dataset".
Remove duplicated columns. In the download, the "data_collected" column is duplicated. This is because the "collection.info=TRUE" argument in the download adds the column "data_collected" in addition to other variables. However, if we select "collection.info=FALSE", we would lose other variables related to collection information and, thus, it is necessary to use “TRUE” (Note 8).
Remove records that are also from the GBIF database to avoid replicated data. BIEN data source may contain occurrences that are also in GBIF, this information is available in the "datasource" column of "raw.BIEN.dataset".
Create necessary columns for merging with GBIF. Add the country code and elevation variables.
i. Add the country codes assigned by the 2-letter ISO 3166-1 standard from the name of the country available in "country" column of “simple.BIEN.dataset”.
ii. Add an empty elevation column with "NA" (Not Available information). The elevation is not included in the downloaded information from BIEN, but it is included in GBIF, and we do not want to lose this information when merging the two databases.
iii. Add an empty "ID_Origin" column filled with "NA". This information is not available in the BIEN database, but it is included in GBIF, and we do not want to lose this information.
iv. Add "dataOrigin" column filled with BIEN. This column indicates if the record belongs to GBIF or BIEN; in this case, it will be filled with BIEN for all records.
v. If your sample unit is species and you have used "BIEN_occurrence_species" to download the data, then you will not have any column indicating the name of the genus. Since this column may be of interest, then it is desirable to run this function to add a column with the name of the genera, named "scrubbed_genus". This line is commented on the script and, thus, it is not performed in the protocol unless you uncomment it (Note 9).
Select useful columns from all available variables. For our purpose, the useful information is inside the following columns: "ID_Origin", "dataOrigin", "observation_type", "scrubbed_genus", "scrubbed_species_binomial", "verbatim_scientific_name", "longitude", "latitude", "country", "country_ISOcode", "locality", "date_collected", "datasource", "collection_code", and "catalog_number". Therefore, we will keep only these columns. We will refer to the resulting data set as "simple.BIEN.dataset" onwards. Note that the R object "ID_Origin" contains the code of the record in the original database but transformed as exponential. To look for the exact original code, go to that field in the exported ".csv".
Export "simple.BIEN.dataset" as a csv file.
Match columns between "simple.GBIF.dataset" and "simple.BIEN.dataset". To merge the two data sets you need to rename columns in both objects to match the following extract names ("ID_Originin", "Data_Origin", "Basis_of_Record", "Genus", "Spp", "Scientific_name", "Longitude", "Latitude", "Elevation", "Country_Name", "Country_ISOcode", "Locality", "Date", "Institution_code", "Collection_code", and "Catalog_number") and following the equivalences indicated in Table 3. If there are no records in “raw.BIEN.dataset”, it is important that you see Note 10.
Table 3. Equivalences between information of GBIF and BIEN simple data sets needed for merging data sets. Names of selected columns of "simple.GBIF.dataset" and "simple.BIEN.dataset", and their corresponding name in the merged data set.
GBIF BIEN Merged
ID_Originin (new) ID_Originin ID_Originin
Data_Origin (new) Data_Origin Data_Origin
genus scrubbed_genus Genus
species scrubbed_species_binomial Spp
scientificName verbatim_scientific_name Scientific_name
decimalLatitude latitude Longitude
decimalLongitude longitude Latitude
elevation Not available (Later created as "elevation") Elevation
countryName (new) country Country_Name
countryCode Not available (Later created as "country_code") Country_code
locality locality Locality
eventDate date_collected Date
institutionCode datasource Institution_code
collectionCode collection_code Collection_code
catalogNumber catalog_number Catalog_number
basisOfRecord observation_type Basis_of_Record
Merge "simple.GBIF.dataset" and "simple.BIEN.dataset". Build the new data set from the equivalence information of the "simple.GBIF.dataset" and "simple.BIEN.dataset" objects specified in Table 3. The resulting table has the information of both data sets, and it has 16 variables. We will refer to the resulting data set as "merged dataset" onwards. If there were no records in "raw.BIEN.dataset", you still need to rename your "simple.GBIF.dataset" to "merged.dataset", because it is the name used onwards in the R script (Note 10).
Save "merged.dataset" as a csv file named "2_merged_dataset.csv". This file contains all the simplified GBIF and BIEN information. Or only GBIF data, in case no record was downloaded from BIEN.
Add occurrences from other sources
This procedure is only necessary if the data from GBIF and BIEN is incomplete (that is, they do not completely reflect the distribution range of the study case) and the author deems it necessary to include other data sources (such as additional online databases, herbarium specimens, or citations in the literature) to complete taxa ranges. If this is not the case, skip this procedure and go to G.
For additional online databases. In this protocol, we will use only GBIF and BIEN occurrences downloaded using "rgbif" and "BIEN" R packages. This step is commented on the R script. However, we provide an optional example using package "spooc" in R. This package can download occurrences from a diverse set of data sources, including Global Biodiversity Information Facility (GBIF; GBIF.org, 2021), USGS Biodiversity Information Serving Our Nation (BISON, 2021), iNaturalist (2021), Berkeley Ecoinformatics Engine (2021), eBird (Sullivan et al., 2009), Integrated Digitized Biocollections (iDigBio, 2021), VertNet (2021), Ocean Biogeographic Information System (OBIS; Grassle, 2000), and Atlas of Living Australia (ALA, 2021). The procedure with this package is very similar to the one already done in this protocol with BIEN and GBIF. The result of the download is a list with as many elements as taxa in the "taxa.names" vector. Each of these elements contains information on the given taxon in the eight databases. Once the "spooc" R package has been run, you will need to reformat and simplify the downloaded databases so that they match the column names and structure specified in step E3. To do this, first go to step E2b and adapt the script onwards to remove GBIF replicates (E2b), create a match column to merge the databases (E2c), adapt the script in E2d to identify the columns of interest, and in E2e export the results in a file named "simple.spooc.dataset". Then adapt E3 to rename columns in "simple.spooc.dataset" as already done for GBIF and BIEN. Finally, go to E4 and adapt it to merge the three databases ("simple.spooc.dataset", "simple.GBIF.dataset", and "simple.BIEN.dataset").
For georeferenced herbarium specimens and literature occurrences. The exported csv file "merged_dataset.csv" is used to manually add the records obtained from herbaria and literature.
Open Microsoft Excel.
Import "merged dataset" csv file exported in step E5.
i. Select "Data > Get External Data > From text" and open de "merged_dataset.csv" file. A new window, called "Text Import Wizard", will appear.
ii. Select "Delimited" in "Original Data Type" and click on "Next".
iii. Select "Comma" in "Delimiters" and click on "Next".
iv. It is advisable to select "Text" in "Column data format" for all columns of the file. Click on "Next". This is done to avoid problems with cell format in Excel that occur in columns that have special characters and symbols and, also (and most importantly), in the columns that include the coordinates.
Add new occurrences filling all the fields of the variables of the table. When no information is available, complete the field with NA. This format will make the importation of the database to R easier (Note 11).
Export the table as csv.
i. Select "File > Save as".
ii. A new window will appear. Choose the "input" folder as the destination folder for the file. Introduce "merged_dataset_Version2" as the file name and select "CSV (Comma delimited) (*.csv)" as the file type. Click on "Save". If you are using Excel for MacOS, then export as "MS-DOS Comma Separated (.csv)" and check that the separators are commas. If that is the case, then run line 260. If separators are semicolons, run the line 259 in R script.
iii. A new window will appear to remind you that only the active spreadsheet will be saved. Select "OK". Select "OK" in the next window that appears.
Check the “merged.dataset” object
It is necessary to check that the data set has the correct format.
If you come from step F2, return to RStudio and import the csv file obtained in step F2diii, "merged_dataset_Version2.csv". This step is commented in the R script because it has not been done in this protocol. If you come from procedure E or step F1, go straight to step G2.
Visualize "merged.dataset". Check that the number of columns in the data set is 16. The structure must be a data.frame. "Latitude", "Longitude", and "Elevation" must be in numeric format, and the rest of the columns must be in character format (Note 12).
Data cleaning
This procedure is focused on cleaning the most common errors.
Remove the records that lack decimal coordinates. This was already done for GBIF records during the import in step D1 but is necessary for records imported from the BIEN database (or any other database if you run F1, Note 12).
Remove the records with invalid coordinate values (that is, Longitude or Latitude = 0). This is common in records from GBIF.
Remove the records in which coordinates have low precision (for the geographical scale and our purposes, we have removed the coordinates that had less than 1 decimals, this can be modified in the script to meet the precision you need as indicated in the R script).
Remove replicated records among databases. Delete the rows that have the same information for the fields: "Genus", "Spp", "Date", "Locality", "Longitude", "Latitude", "Elevation", "basisOfRecord", "catalogNumber", and "CountryCode". Sometimes, the same record has been uploaded in different databases; avoiding redundant data is desirable to reduce processing time. Note that this filter is tied to the number of decimals.
Remove records outside natural distribution of the taxa. Each record of the database after running steps H1–H4 (that is, R object “filtered.dataset”) is compared with a filtered version ("checklist.filtered") of the checklist obtained in step A5 ("checklist"), and those outside the natural distribution will be removed.
Add level-3 TDWG code to "filtered data" for future comparison with the checklist created in step A4.
i. Load the world shapefile with the level-3 TDWG code used by WCSP available in GitHub https://github.com/tdwg/wgsrpd.
ii. Select the WGS84 projection to the shapefile using the "crs" function from "raster" package.
iii. Extract level-3 TDWG code for the occurrences from "filtered.data" (Note 13).
iv. Remove records outside the limits of the level-3 TDWG shapefile. These records correspond to the ones that have "NA" values in the "LEVEL3_COD" column and, also, to those in which coordinates place the occurrence in the sea (because our study case is terrestrial). Remove the "optional" column that is created by default when using the "extract" function (Note 14).
Divide your data set ("filtered.data.TDWG") in sections according to your sample unit. Each section is a table with the records of one genus. Be aware that the sections of this table must correspond to the sample unit of your study (family, genus, species, etc.). Check that the taxa names appearing in "Name" column of the "checklist.filtered" that is cleaned in line 445 of the R script are the same as those indicated at the beginning of the "value" column of the same object.
Filter your data set ("filtered.data.TDWG") to retain only natural records. Compare each genus (or the correspondent sample unit) with the countries of its natural distribution. The "Country_TDWGcode" column of a given sample unit is compared with the corresponding element of "filtered.checklist" created in step H5b from the original checklist created in step A5. The resulting object is a list that contains all the filtered occurrences, and that is converted into a dataframe (“filtered.dataset.WCSP” onwards). Although GBIF database has a column that indicates if the record comes from cultivation, it is highly advisable to run this part of the script to remove naturalized records as well.
Export the "filtered.dataset.WCSP" object as "3_Cleaning_dataset_WCSP.csv" and save the workspace as "2_Workspace_Cleaning.RData".
Distribution maps
This procedure is focused on visualizing the global distribution of all taxa together and individual maps of each sample unit after data cleaning.
Create a global distribution map with all sample units as a PDF named "4_global_distribution_map.pdf". The occurrences of all sample units are colored in red and the surface of the world in grey.
Create distribution maps for each sample unit as a PDF named "4_distribution_maps.pdf". Convert "filtered.dataset.WCSP" dataframe into a list in which each element of the list belongs to the occurrences of a single sample unit. Each page of the PDF contains a map of each sample unit (expanding the geographic area in which it appears) titled with the name of the taxon. The occurrences of each sample unit are colored in red and the surface of the world in grey.
Data thinning
The “spThin” package chooses an occurrence and removes nearby occurrences according to the indicated distance in the buffer. This procedure is intended to remove the bias when spatial data is unevenly distributed across your data set, and there are certain areas for all or a few sample units that are disproportionately sampled. To identify this sampling bias, visually inspect the maps created in procedure I. If your sampling bias affects most or all of your sample units, then proceed with the thinning in step J1a; if the sampling bias affects only one or two sample units, then proceed with the thinning in step J1b. If you detect sampling bias, the thinning is crucial to minimize errors in further spatial-based analyses, such as avoiding overestimation in the bioclimatic data and oversampled areas. If there is no bias in your data set, then you can skip this procedure and go to procedure K.
Remove occurrences randomly with a given distance buffer (50 kilometers in this case) from the "Latitude" and "Longitude" columns. You may need to modify this buffer based on the geographical scale of your case study. This is done in the "thin.par" argument of the "thin" function in the R script. It is very important to put the distance in kilometers. The random elimination of occurrences is done by sample unit. In our case study, the sample unit is indicated in the "Genus" column, but you can modify it in the script if needed.
Perform this step if you want to apply the thinning to all your sample units. The output of this step is a csv file for each sample unit that will be directly exported to the chosen directory (Note 15). If you want to apply the thinning to just one sample unit, then go to step J1b.
Perform this step if you want to run the thinning just in one sample. In this step you will only apply the thinning to the sample unit that is affected by sampling bias. If you have not run step J1a, then select the proper buffer based on your geographical scale (Case 1). If you perform this step because a given sample unit is still affected by sampling bias after running step J1a, you must increase the buffer with respect to the one used in step J1a (Case 2). This step has not been done in the protocol, but the procedure is included and commented on the R script (Note 16).
Import csv files after data thinning using R package "readr" from "thin" folder.
Import csv files for all the taxa obtained in step J1 if you only performed J1a or if you performed J1a and J1b (Note 17). Continue with step J3a.
Perform this step only if the thinning has been performed on only one sample unit (that is, if you run step I1b directly without running step I1a). Import csv file obtained in I1b for the sample unit thinned. This step is commented on the R script because it has not been done in this protocol. Continue with step I3b.
After data thinning, the exported files have only three columns: "Genus", "Longitude", and "Latitude". Therefore, if you run procedure I, add all the remaining 13 columns that contain the additional information of each record from the "filtered.dataset.WCSP" object to the data set obtained after the thinning ("thin.occ", in case you run step I2a; or "thin.taxon", in case you run step I2b).
If you come from step I2a, join columns of the "filtered.dataset.WCSP" to all thinned taxa imported in. Check that "thin.occ" has the same number of rows as "joined.dataset" (Note 18). Check that "joined.dataset" contains all the columns in "filtered.dataset.WCSP".
If you come from step I2b, join columns of the "filtered.dataset.WCSP" to one thinned taxon and add to the rest of the taxa of the "filtered.dataset.WCSP" object. This step is commented on the R script because it has not been done in this protocol (Note 18).
Export the joined file "joined.dataset" as a csv named "5_joined_dataset.csv" and save the workspace with thinned files as "3_Workspace_Thinning.RData".
Load bioclimatic variables from WorldClim version 2
This online climatic database contains 19 variables with the average values of 19 parameters that represent precipitation and temperature for the years between 1970 and 2000. There are two ways to obtain these bioclimatic variables. Alternative 1 is shown in step J1a, and it is available for all the resolutions available in WorldClim (10, 5, 2.5 min, and 30 s). Alternative 2 is shown in step J1b, and it is only available for resolutions of 10, 5, and 2.5 min.
Alternative 1. Download from the WorldClim website and import in R. We use the 30 s resolution, (only available on the website) because it is the highest resolution available and matches the minimum threshold for coordinate precision set in step H3 (two decimals in our case study). For this protocol, we used Alternative 2. If you use Alternative 1, uncomment lines in R code.
Go to the following link: https://www.worldclim.org/data/worldclim21.html. Click in bio 30s, unzip the zip file downloaded named "wc2.1_30s_bio.zip" (Figure 5), and rename the folder as "Bioclimatic _variables_WC2". Relocate this folder inside the "input" folder.
Figure 5. Download bioclimatic variables on the WorldClim website
To avoid sorting problems in R, rename bioclimatic variables as follows: replace original names ("wc2.1_30s_bio_1.tif") by adding a zero before the number or variable ("wc2.1_30s_bio_01.tif"). This is only done for variables 1 to 9.
Import bioclimatic variables to R. Remove "_" and ".tif" characters in column names.
Alternative 2. Download the standard WorldClim Bioclimatic variables directly from R using R package "raster". This alternative is only for the resolution of 10, 5, and 2.5 min. In the argument "res" of "getData" function, indicate 10, 5, or 2.5 for the resolution selected. Thus, if you chose for a minimum threshold of two decimals in step H3 and select this alternative, be aware that you will be losing precision for your climatic analysis. For this protocol, we used this alternative.
Spatial correction for terrestrial organisms
Despite the removal of all occurrences with inaccurate coordinates, it may happen that some of the occurrences may fall outside the limits of the earth's surface according to the limit of the cartographic base used as template. For terrestrial organisms, these occurrences may be wrong (if the distance to the coastal limit is huge) or simply misplaced (if the distance to the coastal limit is small). Because our ultimate goal is to extract climatic data (see procedure L), we do not want to include wrong occurrences, but it is desirable that we do not lose misplaced records. Thus, we need to check for occurrences out of the Earth’s limit to remove wrong occurrences and apply a spatial correction for misplaced occurrences. If there are no occurrences outside Earth’s limits in your database, go to procedure M and proceed with climatic data extraction. If there are occurrences outside the limits, then identify the occurrences that are between the coastal limit and 5 km from the coastal limit (misplaced occurrences) as established by bioclimatic variable 1 of WorldClim version 2 (same limit for the 19 available bioclimatic variables) and recalculate new coordinates so that the occurrence falls in the nearest climatic cell of the template. Occurrences located more than 5 km from the coastal limit (wrong occurrences) are eliminated (Note 19).
Visualize the distribution of all the filtered occurrences of the case study available in the "joined.dataset" object, using bioclimatic variable 1 as template. The map will be automatically exported to a PDF file. If you have not performed the thinning process in procedure J, uncomment the line 563 of the R script only.
Convert "joined.dataset" into the spatial object "joined.dataset.spatial".
Check if all "joined.dataset.spatial" points are within the boundaries of the bioclimatic variable layer. Occurrences located outside the layer boundaries are shown in red, located in the "outside_pts" object. The maps will be exported in a PDF file. If there are no occurrences in red, go to step M1 (Note 20).
Coordinate correction. Extract climatic data from bioclimatic variable 1 for all occurrences of the case study and identify which points have no climatic data (NAs, Note 19). The "nearestLand" function of the "seegSDM" package recalculates the coordinates of the occurrences that lie between the distance set in "dist" and the coastal limit to place it in the nearest climatic cell, and thus obtain climatic data. To check coordinate correction, maps with corrected points in green and uncorrected points in red appear in the viewer in the lower right corner of the RStudio screen. The process is repeated by increasing the distance by 1 km until a 5 km distance from the coastal limit is reached. Occurrences outside the 5 km are removed.
Extract climatic data from bioclimatic variable layers
Convert “joined.dataset” to spatial object and select WGS84 projection (Note 20).
Extract bioclimatic data. We use the "bilinear" method because this method returns values that are interpolated from the values of the four nearest raster cells. The "simple" method returns values in which the point falls. Using the "bilinear" method, we assume that the coordinates of the data set have a certain precision error, whereas if we use the “simple” method this error is ignored. Be aware that this step can be time consuming (Note 21).
Join the bioclimatic values to "joined.dataset" in a new dataframe named "climatic.data" and remove "optional" column created during the extraction of climatic data.
Visualize and export the final data set as “6_Final_dataset.csv”
Save the final workspace as "4_Workspace_Final_Data.RData".
Notes
Any text line preceded by the character "#" in the R script available from GitHub ("commented lines") is only intended to guide the user or to provide alternative code to run under specific cases and will not be run when executed in the R console. If the specific case applies to you, and you need to run the functions in the commented line, you need to uncomment the line (remove "#") and execute it. To comment on a section of R script press "Ctrl + Shift + C", and it will not run.
In case the WCSP geographical information for a given taxon is incomplete or not available, complete the natural distribution of the sample unit in other sources such as literature or other checklists and include it in the text file using the level-3 TDWG code (available in Brummitt, 2001), as mentioned in steps A3 and A4.
To run the lines of the script in RStudio, place the mouse cursor at the beginning of the line or selected regions of the code and press "Ctrl + Enter". If the line is uncommented (that is, not preceded by "#"), it will be run.
Check in the "download_GBIF" folder that there is a zip file with the reference of the "gbif.download.key" object. The results are also available on your downloads user page of GBIF: https://www.gbif.org/user/download.
Replace the function "BIEN_occurrence_genus" with "BIEN_occurrence_species" when sample unit is species and the argument "genus" by "species" inside the function.
If there are no records in "raw.BIEN.dataset" only the object "raw.GBIF.dataset" will be further used in the R script. If this is the case, you must rename the columns in step E3 and replace simple.GBIF.dataset. with merged.dataset in step E4 and continue with the rest of the protocol.
A warning may appear when creating the "countryName" column ("Some values were not matched unambiguously: ZZ"). This is because there are records with an ISO code equal to "ZZ", which corresponds to an unknown country. The space before ZZ corresponds with empty values. The records with ZZ and empty values in ISO code will have NAs in the "countryName" column.
If sample unit is species, replace raw.BIEN.dataset[,-28] by raw.BIEN.dataset[,-27].
If sample unit is species, detach "scrubbed_species_binomial" column into "genus" and "temp.spp" to obtain the name of the genus in a separate column.
If there are no records in "raw.BIEN.dataset" change only the column names of "raw.GBIF.dataset", continue to step E4, and replace merged.dataset <- rbind(simple.GBIF.dataset,simple.BIEN.dataset) by merged.dataset <- simple.GBIF.dataset.
When the coordinate columns are filled, the separator for latitude and longitude columns must be a period. Avoid using ";" in fields since Excel exports csv separated by ";".
If you have added new occurrences in step F2, replace the object "merged.dataset" with "merged.dataset.2" in the steps G2 and H1 of the R script.
A warning may appear when all values of shape.level3TDWG are extracted ("In sp::proj4string(x): CRS object has comment, which is lost in output"). It does not affect the result.
Every time the function "extract()" is used, a new unnecessary column is created called "optional" and it is advisable to remove it.
Depending on the number of occurrences of each sample unit (genus in this case), this step can be time consuming. Note that the number of csv files created must be the same as the number of sample units.
If you run step I1b after running step J1a, it is highly important that you delete the csv file of that given sample unit obtained with the previous buffer (step J1a) to keep just one csv file per sample unit (i.e., the last csv file created for that sample unit in step J1b). The number of csv files in the "thin" folder must be equal to the number of sample units in "taxa.names".
When creating "thin.occ" the following warning appears: "There were "number" warnings (use warnings() to see them)". This warning means: "In bind_rows (x, .id): binding character and factor vector, coercing into character vector". Although this warning does not affect the results, check the object "thin.occ" to confirm that it contains all the information (three columns: "Genus" or "Spp", "Longitude", and "Latitude") exported in csv files in step J2, and that the unzip files are located in the "unzip_GBIF_files" path.
In case the number of rows in the "joined.dataset" is not the same as in "thin.occ", it is possible that joining the two tables may generate duplicates. So, execute line 591: joined.dataset<-joined.dataset[!duplicated(joined.dataset[,c("Genus","Longitude","Latitude")]),]. This is done to remove duplicates and check the number of observations between "joined.dataset" and "thin.occ" again.
We set a limit of 5 km to correct the coordinates, but this value can be modified according to the needs in "success <- ifelse(dist > 5000, TRUE, success)" in step J5. Replace "5000" by the new distance. It is important that the new distance value is expressed in meters.
If the message "Plot occurrences outside layer limits (NA values)" appears when executing step L4, and occurrences outside limits are not obtained, go to step M1 and uncomment the line 762.
Depending on the number of occurrences of each sample unit, this step can be time consuming. If the case study has many occurrences and many sample units, it is advisable to use a computer with a large RAM memory. If the extraction of climatic data of all occurrences takes a long time, replace method = "bilinear" by method = "simple".
Acknowledgments
This protocol was derived for the publication in Coca-de-la-Iglesia et al. (2022), currently available in American Journal of Botany. We acknowledge the reviewers for their comments and suggestions on the manuscript and the code. We are also indebted to the people who are part of the Writing Workshop developed by the Biology and Ecology Departments of the Universidad Autónoma de Madrid, for all the comments and discussions that have helped to realize this work, especially to I. Ramos for helping us correct errors in the code. This study was supported by the Spanish Ministry of Economy, Industry and Competitiveness 607 [CGL2017-87198-P] and the Spanish Ministry of Science and Innovation [PID2019-106840GA-608 C22]. M. Coca de la Iglesia was supported by the Youth Employment Initiative of European 609 Social Fund and Community of Madrid [PEJ-2017-AI-AMB-6636 and CAM_2020_PEJD-610 2019-11 PRE/AMB-15871].
Competing interests
We declare no competing interests.
References
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4,848 | https://bio-protocol.org/en/bpdetail?id=4848&type=0 | # Bio-Protocol Content
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Cell Cycle Analysis of Candida albicans by Flow Cytometry
SP Shraddheya Kumar Patel *
SS Satya Ranjan Sahu *
NA Narottam Acahrya
(*contributed equally to this work)
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4848 Views: 647
Reviewed by: Alba Blesa Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Gut Microbes Jan 2023
Abstract
The cell cycle is a vital process of cell division that is required to sustain life. Since faithful cell division is critical for the proper growth and development of an organism, the study of the cell cycle becomes a fundamental research objective. Saccharomyces cerevisiae has been an excellent unicellular system for unraveling the secrets of cell division, and the process of synchronization in budding yeast has been standardized. Cell synchronization is a crucial step of cell cycle analysis, where cells in a culture at different stages of the cell cycle are arrested to the same phase and, upon release, they progress synchronously. The cellular synchronization of S. cerevisiae is easily achieved by a pheromone or other chemicals like hydroxyurea treatment; however, such methodologies seem to be ineffective in synchronizing cells of multimorphic fungi such as Candida albicans. C. albicans is a human pathogen that can grow in yeast, pseudohyphal, and hyphal forms; these forms differ in morphology as well as cell cycle progression. More importantly, upon subjecting to DNA replication inhibitors for synchronization, C. albicans develops hyphal structures and grows asynchronously. Therefore, here we describe a simple and easy method to synchronize C. albicans cells in the G1 phase and the subsequent analysis of cell cycle progression by using flow cytometry.
Keywords: Cell synchronization Cell cycle Hydroxyurea Filamentation Propidium Iodide SYTOX green Flow cytometry
Background
Cell cycle is a multievent cellular process that plays an essential role in maintaining genome stability by coordinating processes like DNA replication, DNA damage repair, and chromosome segregation during cell division. The whole cycle is completed in four phases, where DNA replication is confined to the S phase, G1 is the gap between the M phase and S phase, G2 is the gap between the S phase and M phase, and the division of the nucleus followed by the cytoplasm is restricted to the M phase (Figure 1). G1 and G2 phases are known for cell growth, metabolic activity, and for preparing for the next phase of the cell cycle (Hartwell et al., 1974). When the cells are proliferating, they can be at any phase of the cell cycle; therefore, to properly explore the mechanism of cell cycle progression, synchronization of a heterogeneous population of cells becomes critical. In human cells, serum starvation and double thymidine block are commonly used approaches for synchronizing a cell population (Ma and Poon, 2017). Saccharomyces cerevisiae cells can be synchronized in the G1 phase of the cell cycle with the mating pheromone, α-factor. Similarly, hydroxyurea and nocodazole synchronize budding yeast cells in the early S phase and G2/M phase, respectively (Rosebrock, 2017).
Figure 1. Overview of the cell cycle phases and some routinely used reagents for cell synchronization at the mentioned phases of the cell cycle. For example: thymidine, methotrexate, hydroxyurea, aphidicolin, dedoxy uridine, and mimosine are used to synchronize the cells at G1 or early S phase.
Upon releasing these synchronized cells into a fresh medium, cells undergo similar cell cycle progression that can be monitored after collecting cell populations at different time points. DNA content measurement of a cell population by using DNA-binding fluorescent dyes and a flow cytometer is one of the reliable techniques for getting insights into the cell cycle dynamics of an organism (Haase and Reed, 2002). The existing methodologies of cell synchronization do not work effectively in the case of fungi like Candida albicans. C. albicans is a gut pathogen that survives in at least three morphological forms: yeast, pseudohyphae, and hyphae. Interestingly, these cell types differ even in the rate and order of cell cycle events (Berman, 2006). In addition, most of these reagents used to synchronize cells also induce filamentation in C. albicans (Manohar et al., 2018; Kumari et al., 2023; Patel et al., 2023). Therefore, cell synchronization is one of the major challenges for exploring the cell cycle in C. albicans. Additionally, the use of external chemical reagents for cell synchronization may affect the native cellular physiology by inducing stress in the cells, which will never be a true estimation of the cell cycle and end up in erroneous cell cycle analyses. Therefore, here we describe and demonstrate a chemical method/approach where a time-point-based incubation results in > 90% cell synchronization at the G1 phase so that the cell cycle analysis can be executed without hampering the native physiology of the cells (Figure 2 and Supplementary Figure 1).
Figure 2. Schematic stepwise demonstration of the cell cycle analysis of C. albicans. Synchronized cells will be allowed to grow for 150 min and will be harvested at every 30 min to monitor cell cycle progression. Cells will be fixed with ethanol, stained with SYTOX green, and analyzed by flow cytometry to complete the cell cycle analysis.
One of the key advantages of using flow cytometry for cell cycle analysis in yeast is its ability to analyze a large number of cells at a single-cell level rapidly. This technique employs a laser-based system to measure the fluorescence emitted upon a fluorochrome (in this case, a nucleic acid staining dye) binding to cellular DNA while cells pass through a flow cell. Different nucleic acid staining dyes including 4′,6-diamidino-2-phenylindole (DAPI), propidium iodide (PI), and 7-amino actinomycin D (7-AAD) have been used by researchers to access the cell cycle based on the cell types to be analyzed, although PI is the preferred one. In S. cerevisiae, PI staining has been proven effective, and the same was initially adopted by us for cell cycle analysis of C. albicans (Manohar et al., 2018). However, a major concern associated with PI is the relatively high coefficient of variation (CV) and low signal amplification rate of PI-stained cells, which limit the accuracy and reproducibility of the cell cycle (Haase and Reed, 2002). To overcome this problem, we now use SYTOX green as it exhibits low CV, high signal amplification with low signal-to-noise ratio, and low photobleaching rate, thus being an ideal fluorochrome for accurate and reproducible DNA content measurement (Thakur et al., 2015; Patel et al., 2023). In summary, this protocol outlines detailed steps involved in the cell cycle analysis of a wild-type strain of C. albicans, SC5314, using SYTOX green with a simple cell synchronization strategy providing the precise measurement of DNA content in each phase of the cell cycle by flow cytometry (Figure 3). By employing this protocol, researchers can gain valuable insights into the cell cycle dynamics, which will be helpful in understanding the diverse physiological processes of C. albicans. The same protocol can be adopted for other fungal species as well.
Figure 3. Demonstration of cell cycle progression of wild-type C. albicans cells. Using the described protocol, a synchronized population of C. albicans cells (SC5314 strain) was allowed to progress through the cell cycle and, at mentioned time points (0–1 h), cells were harvested, fixed, stained with SYTOX green, and analyzed by flow cytometry.
Materials and reagents
C. albicans SC5314 or any other strain of interest
Autoclavable round bottom glass culture tubes (Borosil, catalog number: 9900006)
Yeast extract peptone dextrose broth (YPD) (Himedia, catalog number: M1363)
Ethanol (Fisher Chemical, catalog number: 2051537)
Sterile 1.5 mL microcentrifuge tube (Axygen, catalog number: MCT150LC)
FACS tube (Tarson, catalog number: 850010)
Cuvettes (Eppendorf, catalog number: 0030106300)
Proteinase K (Puregene, catalog number: PG-6070)
RNase A (SRL, catalog number: 98915)
Sodium citrate (Mp Biomedical, catalog number: 194817)
SYTOX green (Thermo Fisher Scientific, catalog number: S7020)
Dimethyl sulfoxide (DMSO) (Mp Biomedical, catalog number: 196055)
Acetic acid glacial 99%–100% (Merck, catalog number: CE1C710249)
Tris (Mp Biomedical, catalog number: 194855)
CaCl2 (Sigma-Aldrich, catalog number: C4901-500G)
Glycerol (Himedia, catalog number: MB060-1L)
NaCl (Mp Biomedical, catalog number: 152575)
Solutions
Sodium citrate buffer (see Recipes)
Proteinase K solution (see Recipes)
RNase A solution (see Recipes)
Recipes
Sodium citrate buffer
Reagent Final concentration Quantity
Sodium citrate 50 mM 14.7 g
H2O n/a Make up the volume
Total n/a 1,000 mL
Adjust the pH to 7.4 with glacial acetic acid.
Proteinase K solution
Reagent Final concentration Quantity
Proteinase K 20 mg/mL 100 mg
Tris-HCl (1 M, pH 8.0) 10 mM 50 μL
CaCl2 (100 mM) 1 mM 50 μL
Glycerol 50% 3.125 mL
H2O n/a 1.775 mL
Total n/a 5 mL
Store the solution in 1 mL aliquots in 1.5 mL microcentrifuge tubes at -20 °C.
RNase A solution
Reagent Final concentration Quantity
RNase A 10 mg/mL 200 mg
Tris-HCl (1 M, pH 7.5) 10 mM 200 μL
NaCl (5 M) 15 mM 60 μL
H2O n/a 19.54 mL
Total n/a 20 mL
Boil the solution at 95 °C for 15 min and allow it to cool slowly at room temperature. Store the solution in 1 mL aliquots in 1.5 mL microcentrifuge tubes at -20 °C.
Equipment
Laminar airflow (Thermo Scientific Biological Safety Cabinets, catalog number: 41346502)
Sterile pipette sets (Gilson, catalog number: F123606-1mL, F123605-200µL, and F123604-20µL)
Spectrophotometer (Eppendorf Bio Photometer plus, catalog number: 6132)
Shaker incubator 30 °C (Scigenic Biotech, catalog number: LE-4676-AA)
Incubator 30 °C (Scigenic Biotech, catalog number: C-1NC-100-1)
Centrifuge (Eppendorf, catalog number: 5810R with SL086 rotor)
Flow cytometer (BD LSRFortessa Cell Analyzer, catalog number: 647177H6)
-20 °C freezer (Vestfrost, catalog number: BFS 345)
Heat block (Eppendorf Thermo Mixer C, catalog number: 5382000023)
Software
BD FACSDiva Software
FlowJo v8.2.0
GraphPad prism v8.0
Procedure
Cell synchronization
Freshly streak C. albicans strain on a sterile YPD agar plate and incubate at 30 °C for 24–36 h. Inoculate an isolated colony in 5 mL of sterile YPD broth and grow at 30 °C in a shaker incubator for 16 h at 200 rpm. This growth condition has been repeatedly found to cause effective spontaneous synchronization of C. albicans cells (> 90% cell synchronization will be achieved). Critical: Always pick a single isolated colony and inoculate it in 5 mL of YPD broth for all the strains to be analyzed.
Dilute the synchronized culture with YPD broth to obtain an OD600 of 0.2 in a total volume of 10 mL and allow them to grow for up to 150 min at 30 °C and 200 rpm for timewise cell collection.
Cell fixation by ethanol
Collect 1 mL of C. albicans every 30 min in a 1.5 mL microcentrifuge tube and centrifuge at 10,621× g for 1 min at room temperature. Remove the supernatant and wash the pellet twice with 1 mL of sterile distilled water. Resuspend the pellet in 1 mL of sterile distilled water and immediately place it on ice to arrest cell progression.
Take 10 μL of culture for cell number counting by using a hemocytometer. Collect approximately 1 × 107 cells from the stock for ethanol fixation. Critical: When working with multiple strains simultaneously, determine the cell number of any one of the strains (not necessary to count the cell number of every strain). Accordingly, dilute all the strains to get the desired cell number for fixation. As the methodology is standardized for a wide range of cell populations (1 × 106–1 × 108 cells), there will be no issue with minor differences in cell populations between different samples.
Resuspend the counted (~1 × 107 cells) cells in 300 μL of sterile distilled water and add 700 μL of 100% cold ethanol dropwise while vortexing. Vortex gently in between the drops. Caution: While adding fixative, vortex slowly to ensure good fixative penetration and reduce cell clumping. As the fixation process is an exothermic reaction and heat liberated could damage the sample, fix the cells with cold ethanol only.
Incubate the ethanol-resuspended cells at -20 °C for 16–20 h. Critical: C. albicans strains with pseudohyphal and hyphal cells need to be incubated for 24 h.
Pause point: After ethanol fixation, cells can be stored for several months at -20 °C by sealing properly with parafilm.
Caution: Cells can only be stored in ethanol as it will make the cells porous, which helps efficient staining by SYTOX green at later stages. Storing in ethanol also prevents microbial growth during long-term storage. Unlike fixed animal cells that can be stored at 4 °C, fungal cells, due to thick cell walls, must be stored at -20 °C for efficient pore formation.
Rehydration of fixed cells
Centrifuge the ethanol-fixed cells at 10,621× g for 2 min at room temperature and pipette out the supernatant carefully without disturbing the pellet. Caution: Cells will form a loose pellet after centrifugation; if the pellet is disrupted during aspiration, repeat centrifugation for 1 min at the same g force. Remove the ethanol completely as ethanol presence will hamper downstream RNase A and Proteinase K enzymatic digestion. One should open the cap of the tube carefully for rehydration as ethanol spillage may erase the sample labeling.
Resuspend the pellet in 1 mL of 50 mM sodium citrate buffer (pH 7.2), mix by vortexing, and incubate for 10 min at room temperature for rehydration. Centrifuge the cells at 10,621× g for 2 min at room temperature and pipette out the supernatant.
Repeat step C2 for complete rehydration and finally resuspend the rehydrated cells in 1 mL of 50 mM sodium citrate buffer (pH 7.4).
RNase A and Proteinase K treatment
Incubate the rehydrated cells in 25 μL of 10 mg/mL RNase A at 37 °C for 12 h for RNA degradation. Critical: Since SYTOX green can bind to both DNA and RNA, make sure that RNase A is active to cause complete digestion of RNA.
After incubation, directly add 25 μL of 20 mg/mL Proteinase K into the samples without centrifugation, vortex briefly, and incubate at 50 °C for 1 h. Critical: Increase the incubation time for an additional hour when working with pseudohyphal and hyphal C. albicans cells. Proteinase K treatment breaks down cellular proteins to produce uniform optical scattering and inactivates nucleases to prevent DNA degradation in the samples.
Centrifuge the enzymatically treated cells at 12,745× g for 1 min at room temperature and carefully remove the supernatant. Resuspend the cells in 500 μL of 50 mM sodium citrate buffer (pH 7.4).
SYTOX green staining
Take 200 μL of SYTOX green from a 5 mM stock and make up the volume to 1 mL with DMSO to make a 1 mM working solution. Add 1 μL of 1 mM SYTOX green working solution to each sample and incubate them at 4 °C for 2 h in the dark. Caution: Prepare the SYTOX green working solution in an amber color microcentrifuge tube, divide the working solution into 100 μL aliquots, and store at -20 °C. Prevent direct light exposure at the time of staining by covering the FACS tubes with aluminum foil during incubation.
Wash the cells with 500 μL of 50 mM sodium citrate buffer (pH 7.4) and transfer it into properly labeled FACS tubes for acquisition. Pause point: Stained cells can be stored overnight for next day acquisition as the SYTOX green is a relatively more stable dye than PI.
Experimental setup and cell acquisition
Perform the routine cleaning of the flow cytometer to avoid contamination and switch on the laser light prior to acquisition. Caution: Before starting the acquisition, fill the sheath tank and empty the discard tank for uninterrupted acquisition. Switch ON the laser 10 min before starting acquisition to stabilize the laser, which is important for efficient excitation of the fluorochrome.
Login to the respective FACS software (BD FACSDiva software), open the workspace, and set up a new experiment. Select the forward-scatter area and height (FSC-A & H) and the side-scatter area and height (SSC-A & H) along with SYTOX green in the Cytometer window and ensure the connectivity status. Critical: SYTOX green is excited by Blue laser (488 nm) with an emission maximum of 523 nm, which is similar to that of fluorescent isothiocyanate (FITC) and Alexa fluor 488; so, one can select FITC or Alexa Fluor 488 if SYTOX green is not listed in the fluorochrome panel of the Cytometer window as cytometer configuration varies from machine to machine.
Set up the dot plot of SSC-A vs. FSC-A to eliminate debris and FSC-H vs. FSC-A to discriminate singlet population, along with the histogram plot SYTOX green-A vs. Count to determine the different stages of the cell cycle in the worksheet window.
Run a test sample at low flow rate to adjust the voltage of all three parameters (FSC, SSC, and SYTOX green) and determine the events per second. Critical: Maintain a low flow rate for all samples throughout the acquisition. Do not change the flow rate to medium or high, and do not acquire samples with different flow rates as that will cause variation in SYTOX green excitation, and the obtained results will be erroneous.
Once the experimental setup and voltage setting are stable, acquire all the samples with 50,000 gated events. After the acquisition, clean the fluidics system according to the user’s manual and switch off the laser. Critical: To avoid cell clumping, place a vortex machine next to the flow cytometer, vortex for a brief period, and then quickly acquire. Maintain 400–500 events/second for all the samples and accordingly dilute the samples with sodium citrate buffer.
Data analysis
Export data of all individual samples in .fcs file format from the FACSDiva software and import into the FlowJo v8.2.0 software for analysis.
Follow a similar gating strategy as that of the FACSDiva software during acquisition. First, discriminate the debris from the cells by plotting SSC-A vs. FSC-A; then, from the cells, gate the singlet population by plotting FSC-H vs. FSC-A; finally, take singlet cells for cell cycle analysis via histogram plot of SYTOX green-A vs. Count (Figure 4).
Figure 4. Demonstration of gating strategy during cell cycle analysis using FACSDiva software. Stained cells were shorted for the singlets by forward and side scattering, and then analyzed for SYTOX green staining. At 0 h, most of the cells were found to be synchronized to the G1 phase. For each time point, a similar gating strategy was applied.
Create a batch analysis of the histogram plot having all samples in the layout editor with proper labeling of each stage of the cell cycle, and finally export the images into PowerPoint.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Patel, SK. et al. (2023). Pol32, an accessory subunit of DNA polymerase delta, plays an essential role in genome stability and pathogenesis of Candida albicans. Gut Microbes (Figure 3, panel d).
Manohar, K. et al. (2018). TLS dependent and independent functions of DNA polymerase eta (Polη/Rad30) from Pathogenic Yeast Candida albicans. Mol. Microbiology (Figure 8).
Notes
When working with fast/slow growing strains of C. albicans, perform a growth curve analysis and compare the growth difference in terms of time with the wild-type strain, and accordingly increase or decrease the time of incubation from 16 h for cell synchronization at 30 °C. For example: if a strain is 30 min slower to reach the stationary phase as compared with the wild-type strain, then incubate that particular strain an additional 30 min for exact synchronization.
RNA digestion is very crucial for cell cycle analysis, as improper digestion leads to indistinct cell stages (absence of prominent G1 and G2 phase). Check for the presence of RNA through fluorescence microscopy; cytosol will give off a bright green fluorescence if RNA is not properly digested, whereas punctate nuclei will be observed with proper RNA digestion without cytosolic fluorescence.
Activity of RNase A and Proteinase K can be checked by setting up digestion with purified RNA and protein, respectively.
Check the optical alignment of the flow cytometer prior to acquisition and perform check-beads run for proper alignment if necessary. Improper alignment may lead to a high CV.
Always keep the XML file with the FCS files while exporting the data from the FACSDiva software, which will help in reimporting the data to the FACSDiva software, if necessary, as an FCS file alone cannot be imported to FACSDiva.
Acknowledgments
This protocol was developed based on previously published results by the authors (Manohar et al., 2018; Patel et al., 2023). This work was supported in part by DBT and SERB grants to NA.
Competing interests
The authors declare no competing interests.
References
Berman, J. (2006). Morphogenesis and cell cycle progression in Candida albicans. Curr. Opin. Microbiol. 9(6): 595–601.
Haase, S. B. and Reed, S. I. (2002). Improved flow cytometric analysis of the budding yeast cell cycle. Cell Cycle 1(2): 132–136.
Hartwell, L. H., Culotti, J., Pringle, J. R. and Reid, B. J. (1974). Genetic control of the cell division cycle in yeast. Science 183(4120): 46–51.
Kumari, P., Sahu, S. R., Utkalaja, B. G., Dutta, A. and Acharya, N. (2023). RAD51–WSS1-dependent genetic pathways are essential for DNA–protein crosslink repair and pathogenesis in Candida albicans. J. Biol. Chem. 299(6): 104728.
Ma, H. T. and Poon, R. Y. C. (2017). Synchronization of HeLa Cells. Methods Mol. Biol. 1524: 189–201.
Manohar, K., Peroumal, D. and Acharya, N. (2018). TLS dependent and independent functions of DNA polymerase eta (Polη/Rad30) from Pathogenic Yeast Candida albicans. Mol. Microbiol. 110(5): 707–727.
Patel, S. K., Sahu, S. R., Utkalaja, B. G., Bose, S. and Acharya, N. (2023). Pol32, an accessory subunit of DNA polymerase delta, plays an essential role in genome stability and pathogenesis of Candida albicans. Gut Microbes 15(1): e2163840.
Rosebrock, A. P. (2017). Methods for Synchronization and Analysis of the Budding Yeast Cell Cycle. Cold Spring Harb. Protoc. 2017(1): pdb.top080630.
Thakur, S., Cattoni, D. I. and Nöllmann, M. (2015). The fluorescence properties and binding mechanism of SYTOX green, a bright, low photo-damage DNA intercalating agent. Eur. Biophys. J. 44(5): 337–348.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Cellular morphology of wild-type C. albicans cells before and after synchronization without and with staining with SYTOX green
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Microbiology > Microbial cell biology
Cell Biology > Cell-based analysis > Flow cytometry
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Serial-section Electron Tomography and Quantitative Analysis of Microtubule Organization in 3D-reconstructed Mitotic Spindles
RK Robert Kiewisz
DB Daniel Baum
TM Thomas Müller-Reichert
GF Gunar Fabig
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4849 Views: 933
Reviewed by: Xiaokang WuDick McIntoshAlexandros C Kokotos
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Original Research Article:
The authors used this protocol in eLIFE Jul 2022
Abstract
For the analysis of cellular architecture during mitosis, nanometer resolution is needed to visualize the organization of microtubules in spindles. Here, we present a detailed protocol that can be used to produce 3D reconstructions of whole mitotic spindles in cells grown in culture. For this, we attach mammalian cells enriched in mitotic stages to sapphire discs. Our protocol further involves cryo-immobilization by high-pressure freezing, freeze-substitution, and resin embedding. We then use fluorescence light microscopy to stage select mitotic cells in the resin-embedded samples. This is followed by large-scale electron tomography to reconstruct the selected and staged mitotic spindles in 3D. The generated and stitched electron tomograms are then used to semi-automatically segment the microtubules for subsequent quantitative analysis of spindle organization. Thus, by providing a detailed correlative light and electron microscopy (CLEM) approach, we give cell biologists a toolset to streamline the 3D visualization and analysis of spindle microtubules (http://kiewisz.shinyapps.io/asga). In addition, we refer to a recently launched platform that allows for an interactive display of the 3D-reconstructed mitotic spindles (https://cfci.shinyapps.io/ASGA_3DViewer/).
Key features
• High-throughput screening of mitotic cells by correlative light and electron microscopy (CLEM).
• Serial-section electron tomography of selected cells.
• Visualization of mitotic spindles in 3D and quantitative analysis of microtubule organization.
Graphical overview
Keywords: Microtubules Mitotic spindle HeLa cells Tissue culture cells Human cells Serial sectioning Electron tomography 3D reconstruction Automatic segmentation of microtubules Automated spatial graph analysis Electron microscopy Correlative light and electron microscopy CLEM
Background
It is becoming more and more important to collect data on the three-dimensional (3D) architecture of cells in different model systems. For example, in the field of mitosis, it is important to understand the organization of the microtubule cytoskeleton in each mitotic stage. Working with 3D data, however, can be challenging due to a number of specific methodological and computational requirements. Here, we apply serial-section electron tomography for quantitative analysis of spindle organization in mammalian cells in culture. Obtaining 3D data by electron tomography is time consuming and this leads to the generation of large data sets that can be difficult to handle. Electron tomography also requires extensive computational resources. Moreover, analysis of the generated 3D spatial graphs involves complex computational and visualization tools due to the high complexity of the spindle ultrastructure (Müller-Reichert et al., 2003; Kiewisz et al., 2022). There is a clear need for dedicated software tools for analyzing 3D spatial graphs. Such software can help researchers overcome the challenges associated with working with 3D data and accelerate research in the field of mitosis. Recently, progress has been achieved by increasing access to resources for handling and analyzing large data sets.
Our protocol gives step-by-step instructions for sample preparation and data acquisition by electron tomography (O’Toole et al., 2020). In addition, the computational tools for the quantification and visualization of the 3D data will also be described in detail. The workflow presented here has been applied for serial-section large-scale electron tomography of entire mitotic spindles in mitotic HeLa cells (O’Toole et al., 2020; Kiewisz et al., 2022). Our workflow, however, can be applied for the visualization and analysis of any other cell type grown on appropriate support. In addition, we introduce a newly developed platform that allows for quantitative analysis of the 3D spatial graphs, including all segmented microtubules of the 3D-reconstructed spindles (https://cfci.shinyapps.io/ASGA_3DViewer/). This platform, termed Automated Spatial Graph Analysis (ASGA) is designed for a flexible online display of spindles with a modular structure that allows for the swift addition of new analysis tools. Using this 3DViewer within ASGA, segmented spindles can be visualized, quantitatively analyzed, and interactively shared.
Materials and reagents
Biological material
HeLa Kyoto cells (Gerlich Lab, IMBA, RRID: CVCL_1922)
Reagents
Acetone (CH3COCH3), glass-distilled, EM-grade (EMS, Electron Microscopy Sciences, catalog number: 10015)
BSA (albumin fraction V) (Carl Roth, catalog number: 8076.2)
Chloroform (CHCl3) (EMS, catalog number: 12550)
Colloidal gold solution (Ø 20 nm) (BBI, British Biocell International Solutions, catalog number: EM.GC20/4)
Disodium hydrogen phosphate (Na2HPO4) (Carl Roth, catalog number: P030.1)
DMEM (Dulbecco’s Modified Eagle Medium, 1×) + GlutaMaxTM-I (Thermo Fisher Scientific, catalog number: 10566016)
DMSO (Thermo Fisher Scientific, catalog number: D4540-100ML)
Ethanol absolute (C2H5OH) (VMR, catalog number: 20821.296)
FBS (fetal bovine serum) (HI), origin: USA (Thermo Fisher Scientific, catalog number: 10082147)
Fibronectin (Sigma-Aldrich, catalog number: F0556-100UL)
Formvar (Science Services, catalog number: E15800)
High-vacuum grease (Thorlabs Elliptec GmbH, catalog number: SG10)
Hydrogen peroxide 30% (Merck KGaA, catalog number: 1072100250)
Lead citrate (C12H10O14Pb3) (Science Services, catalog number: E17810)
Methanol (CH3OH) (VWR, catalog number: 20834.291)
Methylene blue (C16H18ClN3S) (Carl Roth, catalog number: A514.1)
Osmium tetroxide (OsO4) (Science Services, catalog number: E19100)
Penicillin/Streptomycin (Thermo Fisher Scientific, catalog number: 10378016)
Poly-L-lysine (0.01%) (Sigma-Aldrich, catalog number: A-005-C)
Potassium chloride (KCl) (AppliChem GmbH, catalog number: 31494.1210)
Potassium dihydrogen phosphate (KH2PO4) (Carl Roth, catalog number: 3904.1)
Propanol (C3H7OH) (VWR, catalog number: 20858.293)
Sodium chloride (NaCl) (Carl Roth, catalog number: HN00.2)
Sulfuric acid 96% (H2SO4) (AppliChem GmbH, catalog number: 131058.1211)
Trypsin-EDTA 1× (0.25%) (Thermo Fisher Scientific, catalog number: 25200056)
Uranyl acetate [UO2(CH3COO2)] (Polysciences, catalog number: 21447-25)
Solutions
AFS (automatic freeze substitution) cocktail containing methylene blue (see Recipes)
DMEM-WT (standard medium for mammalian cell culture) (see Recipes)
DMEM-HPF (high-pressure freezing) solution (see Recipes)
Epon/Araldite resin (see Recipes)
Fibronectin coating solution (see Recipes)
Formvar solution (see Recipes)
Lead citrate post-staining solution (see Recipes)
PBS 10× buffer (see Recipes)
Uranyl acetate post-staining solution (see Recipes)
Recipes
AFS (automatic freeze substitution) cocktail containing methylene blue
Oversaturate 60 mL of acetone from a freshly opened bottle of glass-distilled, anhydrous acetone with methylene blue by adding 5 g to it and mixing it by shaking it several times at room temperature. After placing the closed bottle at -80 °C overnight, filter the solution with a syringe filter. Then, use the filtered acetone for the preparation of the AFS cocktail as described below. Aliquot 1 mL each in 2 mL cryo-tubes, freeze them immediately in liquid nitrogen, and store them in liquid nitrogen until further use.
Reagent Final concentration Quantity
Osmium tetroxide 1% (w/v) 0.5 g
Uranyl acetate 0.1% (w/v) 0.05 g
Acetone (saturated with methylene blue) n/a 50 mL
Total n/a 50 mL
DMEM-WT (standard medium for mammalian cell culture)
Reagent Final concentration Quantity
FBS 9.01% (v/v) 50 mL
Penicillin/Streptomycin 0.9% (v/v) 5 mL
DMEM n/a 500 mL
Total n/a 555 mL
DMEM-HPF (high-pressure freezing) solution
Reagent Final concentration Quantity
BSA 10% (w/v) 1 g
DMEM-WT n/a 10 mL
Total n/a 10 mL
Epon/Araldite resin
Mix all reagents in a single-use plastic beaker with a magnetic stirrer overnight. The next day, fill up 10 mL syringes with mixed resin, close them with Parafilm, and store them at -20 °C until further use.
Reagent Final concentration Quantity
Embed 812 n/a 25 mL
Araldite 502 n/a 15 mL
DDSA n/a 55 mL
BDMA n/a 2.5 mL
Total n/a 97.5 mL
Fibronectin coating solution
Reagent Final concentration Quantity
Fibronectin n/a 0.1 mL
PBS (1×) n/a 0.9 mL
Total n/a 1 mL
Formvar solution
Reagent Final concentration Quantity
Formvar powder 1% (w/v) 1 g
Chloroform n/a 100 mL
Total n/a 100 mL
Lead citrate post-staining solution
Reagent Final concentration Quantity
Lead citrate 0.4% (w/v) 2 g
Double-distilled water n/a 50 mL
Total n/a 50 mL
PBS (10×) buffer
Reagent Final concentration Quantity
Na2HPO4 100 mM 8.9 g
KH2PO4 18 mM 1.2 g
NaCl 1.37 M 40 g
KCl 27 mM 1 g
Double-distilled water n/a 500 mL
Total n/a 500 mL
Uranyl acetate post-staining solution
Reagent Final concentration Quantity
Uranyl acetate 2% (w/v) 0.2 g
Methanol n/a 10 mL
Total n/a 10 mL
Laboratory supplies
Beaker, single-use plastic (VWR, catalog number: 414004-148)
Compressed air bottle (Falcon Safety Products UK Limited, catalog number: 88004/DPSRX/UK)
Coverslips (24 mm × 60 mm) (Marienfeld, catalog number: 0107242)
Cryo-tube (1.5 mL) (Greiner Bio-One, catalog number: 126261)
Desmotome VT1, straight (Ustomed, catalog number: 83-898-000)
Epon/Araldite (Araldite 502/Embed 812/DDSA/BDMA kit) (Science Services GmbH, catalog number: 13940)
Eppendorf tubes (0.5, 1, and 2 mL) (Eppendorf, catalog number: 0030121589, 0030120094)
Falcon tubes (15 and 50 mL) (Sigma-Aldrich, catalog number: T1943, T2318)
Filter paper (Sigma-Aldrich, catalog number: WHA1540090)
Flow-through rings (Leica, catalog number: 16707157)
Glass slides (76 mm × 26 mm) (Engelbrecht, catalog number: 11101)
Grid staining system (PELCO® Staining slot matrix) (Ted Pella, catalog number: 22510)
Grid staining system (PELCO® Staining vessels) (Ted Pella, catalog number: 22510-2)
Grids (oval slot 2 mm) (EMS, catalog number: G2010-Cu)
NuncTM EasYFlaskTM (T-75 and T-175) (Thermo Fisher Scientific, catalog number: 156499, 159910)
Parafilm (Pechiney, catalog number: HS234526A)
PHA/PLA 3D printer filament (1.75 mm), natural color (ColorFabb, catalog number: 010003)
Planchettes for high-pressure freezing (Ø 3 mm, indentation 40/600 μm) (Wohlwend GmbH, catalog number: Art. 737)
Precision wipes (KimTech) (Kimberly-Clark, catalog number: 75512)
Razor blades (double edge, coated) (EMS, catalog number: 72000)
Razor blades (single edge) (Plano GmbH, catalog number: T586)
Reagent bath (Leica, catalog number: 16707154)
Sapphire discs for high-pressure freezing (Ø 3 mm, thickness 0.16 mm) (Wohlwend GmbH, catalog number: 500)
Syringe (1 mL) (Braun, catalog number: 9166017V-02)
Syringe (10 mL) (Braun, catalog number: 4617100V-02)
Syringe filter (0.2 μm mesh size) (Sarstedt AG, catalog number: 83.1826.001)
Syringe needle (25 G, nr. 18) (Becton Dickinson S.A., catalog number: 305125)
Syringe needle (27 G, nr. 20) (Becton Dickinson S.A., catalog number: 305109)
Tweezers (style 5X) (EMS, catalog number: 78320-5X)
Tweezers (style SS) (EMS, catalog number: 78325-SS6)
Equipment
3D printer (Prusa i3 MK2s) (Prusa Research, catalog number: i3-MK2s)
3D printer nozzle (V6, Stainless-steel nozzle, 1.75 mm) (E3D)
CCD camera (F214), mounted on a TECNAI T12 TEM (TVIPS GmbH)
CCD camera (US1000), mounted on a TECNAI F30 TEM (Gatan)
Cell culture laminar flow cabinet Heraeus (HERASAFE HSP12) (Heraeus)
Centrifuge (5702 R) (Eppendorf, catalog number: 5703000010)
Diamond knife (Ultra 35°) (Diatome, catalog number: DU3515)
Dual-axis TEM holder (Fischione, model: 2040)
Formvar film casting device (EMS, catalog number: 71305-01)
Freezer (MDF U55V, -80 °C) (Sanyo)
High-pressure freezer (Compact 03) (Wohlwend GmbH)
Incubator HERP (Cell 240) (Thermo Fisher Scientific, catalog number: 51026331)
Incubator HPF (TECO 20) (Selutec, catalog number: 600.005)
Leica EM AFS2 (Leica)
Leica SP5 confocal microscope (Leica)
Multitool, Fortiflex (Dremel, catalog number: 9100-21)
Objective Leica (HCX PL APO lambda blue 63× 1.2 water) (Zeiss)
Objective Leica (HCX PL APU CS 20× 0.7 dry) (Zeiss)
Objective Zeiss (A-Plane (44 10 20), 5×/0.12) (Zeiss)
Objective Zeiss (A-Plane (44 10 51), 40×/0.65) (Zeiss)
Oven (UNB200, 60 °C) (Memmert, catalog number: GZ-52200-01)
Scale (BP 110s) (Sartorius)
Stereomicroscope (SMZ-168) (Motic, catalog number: 1100200500454)
Transmission electron microscope (TEM) (TECNAI F12) (Thermo Fisher Scientific)
Transmission electron microscope (TECNAI F30) (Thermo Fisher Scientific)
Ultramicrotome Leica (EM UC6) (Leica)
Warm bath (GFL 14L) (GFL, catalog number: 3340)
Software and datasets
ASGA (v0.37.0, 25.04.2022)
ASGA-3DView (v1.3.1, 03.08.2022)
Automatization scripts for tomogram reconstructions (n/a, 08.11.2020)
EMMenu (4.0.8.16, n/a)
ImageJ/Fiji (v1.52v, 14.04.2020)
IMOD (v4.8.22 - 4.12.40, n/a - 18.03.2023)
Amira (2022.2, 20.11.2022)
AmiraZIBEdition (2023.12, 19.04.2023)
R (v4.00 - 4.2.3, 04.2020 - 03.2023)
SerialEM (v3.0.2 - 3.8.0beta, n/a)
Procedure
Start of the cell culture
Mammalian cells are cultured in DMEM-WT. Cryopreserved cells are used to initiate the culture.
Quickly thaw the cryotube with cells in a warm water bath.
Add thawed cells to a T75 flask with 10 mL of pre-warmed DMEM-WT medium.
Incubate cells for one day at 37 °C with 5% CO2 in a humidified incubator.
The next day, replace the medium with a fresh pre-warmed DMEM-WT medium.
Continue cell passage 2–3 times to ensure the culture is healthy.
The day before the initial experiment, ensure that the cell colony achieves approximately 70% confluency.
High-pressure freezing
3D-printed chambers for high-pressure freezing:
Mitotic cells are highly sensitive and must be handled with care to avoid any interference with the mitotic process. To avoid any difficulties associated with the cryo-preservation of mitotic cells, we developed custom-made chambers using 3D printing technology to plate cells on a solid support. We designed 3D-printed chambers for small sapphire discs (3 mm in diameter). These chambers are designed to provide a stable environment for mitotic cells for a short time period before high-pressure freezing. Due to their small volume, these chambers increase the number of cells attached to the sapphire discs, thus enabling one to collect cells of the desired mitotic stages (Figure 1; Files S1 and S2 for 3D models). The chamber can hold either one (single chamber) or up to four sapphire discs (quadruple chamber). Before use, the 3D-printed chambers are cleaned with ethanol and stored in a humidified incubator at 37 °C with 5% CO2. For a high-throughput approach, we suggest using the quadruple chambers as they allow for an increased yield of collected mitotic cells.
Figure 1. Technical drawing of custom-designed and 3D-printed incubation chambers optimized for attachment of mitotic cells to sapphire discs (Ø 3 mm, red) after shake-off. This chamber design allows for maximizing the yield of attached cells on a small surface area of the sapphire disc. All dimensions are given in millimeters. (A) Culture chamber for a single sapphire disc optimized for small-volume experiments. (B) Culture chamber for four sapphire discs optimized for high-throughput experiments. This figure was reproduced with permission from the book chapter “High-throughput screening of mitotic mammalian cells for electron microscopy using classic histological dyes”, Methods in Cell Biology (162), 151–170 (2021), Copyright Elsevier [Kiewisz et al. (2021)].
Preparation of sapphire disks: Sapphire discs were chosen for cryo-immobilization of cells due to their high thermal conductivity.
Cleanup: clean sapphire disc in pure acetone followed by washing in pure ethanol and extensive washing and storage in ddH2O for further use (Figure 2A).
Figure 2. Attachment of mitotic cells on sapphire discs and assembly of sandwiches for high-pressure freezing. (A) Cleaning steps for the sapphire discs. (B) Collection of mitotic cells by applying the shake-off technique. One or multiple flasks are hit ten times to the surface of the bench. (C) Placement of coated sapphire discs into the 3D-printed incubation chamber designed for four samples. Transparent coated sapphire discs are indicated with black circles. (D) Top view of the opened holder for high-pressure freezing. (E) Placing the aluminum carrier into the sample holder in preparation for freezing. (F) Prefilling the carrier with the freezing medium. (G) Removing a sapphire disc from the quad incubation chamber (lower left corner) with the help of tweezers. This is followed by a quick dipping of the sample into the freezing medium. (H) Placing the sapphire disc onto the aluminum carrier (cells facing down). (I) Closing of the holder to immediately start high-pressure freezing. This figure was reproduced with permission from the book chapter “High-throughput screening of mitotic mammalian cells for electron microscopy using classic histological dyes”, Methods in Cell Biology (162), 151-170 (2021), Copyright Elsevier [Kiewisz et al. (2021)]. See also Video 1.
Video 1. Video tutorial showing the assembly of a sandwich for cryo-immobilization using the Wohlwend Compact 03 high-pressure freezer. This video corresponds to Figures 2 and 3.
Figure 3. Technical drawing of the sandwich as used to cryo-immobilize mitotic cells attached to the sapphire by using the Wohlwend Compact 03 high-pressure freezer. All dimensions are given in millimeter. (A) A sapphire disc (with the attached cells facing down, not shown here) is placed on top of an aluminum carrier with a cavity of 0.04 mm. (B) Cross-section views (A-A as shown in A) of the sample holders. (C) Schematic representation of sandwich assembly and insertion of the sample into the sample holder. The flow of the LN2 during high-pressure freezing is indicated by black arrows. (D) Side view of the sample loading and the sandwich components (sapphire disc and aluminum planchette). This figure was reproduced and modified with permission from the book chapter “High-throughput screening of mitotic mammalian cells for electron microscopy using classic histological dyes”, Methods in Cell Biology (162), 151–170 (2021), Copyright Elsevier [Kiewisz et al. (2021)].
Place all sapphire discs into a glass beaker with the cleaning solution and gently mix it to ensure that all sapphire discs are separated. Let it incubate for approximately 5–10 min.
After the incubation time, remove all sapphire discs with fine tweezers and rinse them in water two times by placing them in three dishes with ddH2O and then once in ethanol (70%, v/v). Store them in an Eppendorf tube in 70% ethanol until further use.
Coating: after the cleanup, coat the sapphire discs with poly-L-lysine (0.1% in ddH2O, w/v) and fibronectin (1:10 dilution in 1× PBS, v/v).
Initially, slowly air dry the sapphire discs by placing them for 1 h in a closed Petri dish to prevent any contamination. Then, coat the discs with poly-L-lysine by applying a single drop on each disc and leaving it for 5 min under a Petri dish.
After that, absorb the excess solution with small pieces of filter paper and then place the sapphire discs on a heating plate set to 60 °C for 2 h.
Remove sapphire discs from the heating plate and leave them to cool down in a closed Petri dish. It helps to place the discs on a slightly sticky rubber pad (e.g., a piece of table tennis rubber or rubber glove). This will prevent the discs from sticking to a pipette tip.
Next, coat the sapphire discs with fibronectin. Prepare a fresh 10% (v/v) fibronectin solution by diluting fibronectin in 1× PBS and gently mixing the solution with a pipette (the solution should not be mixed extensively to prevent fibronectin aggregation).
Finally, apply a drop of fibronectin solution on the sapphire discs and leave them at room temperature for 10 min. After that, remove the droplet with a pipette.
Place the sapphire discs into custom-designed 3D-printed chambers and store them in the cell culture incubator until further use.
Mitotic shake-off: our preferred method for enriching mitotic cells is called a mitotic shake-off (Guizetti et al., 2011; Ma and Poon, 2017). This method takes advantage of the fact that mitotic cells round up during mitosis and can be easily detached. This approach is preferred because it does not require the use of any drugs and reduces temperature stress that may influence mitosis processes (Rao and Engelberg, 1965; Figure 2B).
Warm the culture medium to 37°C and store it open in the incubator to allow for saturation with 5% CO2.
If possible, move all T75 flasks with cell cultures to the incubator located conveniently in the same room as your high-pressure freezer machine one day before the planned experiment. This will limit the number of environmental changes during the mitotic shake-off.
To synchronize the cells, remove the cell culture flask from the incubator and hit the bottom of the flask vigorously on a solid surface (e.g., the bench).
Remove the entire medium from the flask that contains primarily dead and already mitotic cells and wash it by adding 5 mL of pre-warmed medium.
Discard the medium from the flask, add 10 mL of fresh pre-warmed medium, and immediately place it back in the incubator for the next 2 h.
Repeat steps B3c–B3e for multiple flasks at approximately 15–20 min intervals if multiple rounds of cryo-immobilization are needed.
After 2 h of incubation time, repeat step B3c for the first flask.
This time, collect all the medium after the shake-off with mainly mitotic cells by aspirating it with a 15 mL pipette, and place it in a single 15 mL Falcon tube.
Centrifuge collected medium gently at 0.2× g for 5 min at 37 °C. After this step, one should be able to see a small cell pellet at the bottom of the Falcon tube.
Remove the entire supernatant. Be very careful to not remove the cell pellet.
Resuspend the cells with 1 mL of pre-warmed medium, mix it, and immediately add it to the sapphire discs stored in the 3D-printed chamber (Figure 2C).
Immediately move the 3D-printed chamber to a humidified incubator at 37 °C with 5% CO2 for 10–15 min (depending on the mitotic stage that should be analyzed and on the cell type) to allow for the reattachment of the cells to the sapphire discs.
Sandwich assembly for high-pressure freezing and cryo-immobilization:
For optimal cryo-immobilization and ultrastructural preservation of the samples, high-pressure freezing was applied (McDonald and Morphew, 1993). We used the Wohlwend Compact 03 high-pressure freezer (Figure 3; Video 1). However, any other high-pressure freezer could be used for cryo-immobilization of mammalian cells.
Warm up a metal heating block to 37 °C.
Remove the 3D-printed chamber with sapphire discs from the incubator and place it on a prewarmed metal block.
Assemble each sandwich as shown in Figure 2D–2I. For this, place the sample holder under a stereomicroscope (Figure 2D).
Place an aluminum planchette with the 40 μm cavity facing up inside the holder (Figure 2E) and fill the cavity with 1 μL of the pre-warmed DMEM high-pressure freezing solution (Figure 2F).
Take one sapphire disc from the 3D-printed chamber and remove it slowly from the chamber.
Dip the sapphire disc with cells in the pre-warmed DMEM high-pressure freezing solution (Figure 2G).
Flip the sapphire disc so that the attached cells face downward. Then, gently place a sapphire disc on top of an aluminum carrier (Figure 2H). Check that air bubbles are not trapped between the aluminum carrier and the sapphire disc, as this will result in bad cryo-immobilization and eventually a breaking of the sapphire disc during freezing.
Immediately proceed by closing the sample holder (Figure 2I) and freeze the specimen using the high-pressure freezing machine.
Retrieve the sandwich of the aluminum planchette and the sapphire disc with the cryo-immobilized cells by opening the sample holder under liquid nitrogen. Press with a cold tweezer from the back and place the sapphire discs inside a cryo-tube. Be careful not to lift out the specimen from the liquid nitrogen bath during this procedure.
Store all cryo-immobilized specimens under liquid nitrogen until further use.
Freeze-substitution and embedding in Epon/Araldite
After cryo-immobilization, it is necessary to perform freeze-substitution of the cells (McDonald and Müller-Reichert, 2002; Studer et al., 2008). This process allows for a slow exchange of cellular water for acetone and later for epoxy resin (Epon/Araldite). The addition of osmium tetroxide preserves the cellular ultrastructure for subsequent transmission electron microscopy (TEM) (Mollenhauer, 1964). This process also introduces heavy metals (osmium tetroxide and uranyl acetate) to the sample, thus providing contrast for electron microscopy. We also add methylene blue to the freeze substitution cocktail to selectively stain chromosomes for imaging of the resin-embedded samples by light microscopy (Kiewisz et al., 2021). This step allows for the selection and staging of mitotic cells before TEM.
Freeze-substitution of cryo-immobilized specimens:
Prepare saturated methylene blue solution in acetone by adding 5 g of methylene blue powder into 60 mL of anhydrous, glass-distilled acetone at room temperature and mix it well.
Store the solution in a 50 mL Falcon tube and leave it at room temperature for 2 h.
After that time, store the solution at -80 °C overnight.
After removing the solution from the -80 °C freezer, quickly filter the dye-saturated acetone solution with a syringe filter.
Prepare a freeze-solution cocktail by adding 1% (w/v) osmium tetroxide and 0.1% (w/v) uranyl acetate to the dye-saturated acetone. Store 1 mL aliquots of this freeze-substitution solution in 2 mL cryo-vials in liquid nitrogen until further use.
Open the high-pressure frozen sandwiches under liquid nitrogen and transfer the sapphire discs to the cryo-vials containing the AFS cocktail.
Perform freeze-substitution in an automated freeze-substitution machine. Keep samples at -90 °C for 1 h, before warming up to -30 °C with increments of 5 °C/h. Then, keep samples at -30 °C for 5 h, and again warm up to 0 °C in steps of 5 °C/h. At 0 °C, remove the sapphire discs from the cocktail (samples should not remain at 0 °C for more than one hour to avoid overstaining).
Infiltration with Epon/Araldite (resin) and specimen embedding:
Wash the samples one time with pure anhydrous acetone at room temperature inside the cryo-vials (this will warm up a sample to room temperature in one rinse step) and then transfer the sapphire discs to a glass Petri dish filled with anhydrous acetone.
Place the Petri dish under a binocular microscope and arrange each sapphire disc with cells facing up. To determine the side of the disc with the attached cryo-immobilized cells, the sapphire discs are inspected for cellular contents under the dissection microscope while remaining immersed in acetone.
Transfer each sapphire disc to round flow-through rings within a plastic reagent bath (Leica), designed for 3 mm sapphire discs, with the cells facing up. Pre-fill the molds to a low level already with a 1:3 (v/v) resin/acetone mixture to prevent the sapphire discs from drying out. After all sapphire discs have been transferred to the plastic molds, check again under a dissection scope that all are properly placed in the indentations at the bottom of the molds. Then, fill up the mold to at least three-quarters with the acetone/resin mixture.
Infiltrate the samples with Epon/Araldite resin at room temperature with increasing concentrations of resin/acetone (v/v). Start with 1:3, then 1:1, and then 3:1 (1 h for each step). The exchange of the resin solutions should be performed very slowly to prevent the detachment and loss of cells from the surface of the sapphire discs and to avoid movement of the discs.
As a final step, fill the block molds with pure Epon/Araldite resin, and incubate the samples at room temperature overnight. It is important to completely fill the molds to avoid a meniscus of the resin within the individual mold subunits. This will cause a lensing effect of the transmitted light in the light microscope that is used to screen the blocks for cells of interest during the screening step.
Polymerize samples at 60 °C for at least 48 h in an oven.
Release of resin-embedded cells from sapphire discs:
Cut the plastic molds open with a scalpel or a mechanical multitool (Dremel Fortiflex). Break the sample blocks out of the molds using a desmotome VT1 (a tool with a sharp arrow-shaped tip) as a chisel and a hammer. Under an ultramicrotome, carefully remove the resin around and on top of the sapphire discs using a single-edge razor blade. This needs to be done with great care to avoid breakage of the sapphire disc edges (Figure 4).
Figure 4. Removal of a sapphire disc from the Epon/Araldite block after resin polymerization. (A) Resin block with attached sapphire disc locked in self-locking forceps and dipped into liquid nitrogen. Only the sapphire disc should be submerged in liquid nitrogen for a few seconds. (B) Using a heating block to remove the sapphire disc from the cold resin block. After a few seconds, when the ice is starting to form on the surface of the cold sapphire disc, the edge of the sapphire disc is pressed to the heated razor blade of the heating block. This will leave the embedded cells (not shown here) in the resin. This figure was reproduced with permission from the book chapter: “High-throughput screening of mitotic mammalian cells for electron microscopy using classic histological dyes”, Methods in Cell Biology (162), 151–170 (2021), Copyright Elsevier [Kiewisz et al. (2021)].
Lock each sample in the self-locking straight forceps (artery clamp). Submerge only the top part of the resin block containing the sapphire disc into liquid nitrogen for a few seconds (Figure 4A). Avoid submerging the block too deep into the liquid nitrogen as this could break the resin block.
Remove the discs by gently pressing the edge of the discs to a heated razor blade (Figure 4B). The sapphire discs should self-release from the surface of the specimen without using extensive force, due to the different shrinkage and expansion of the resin and the sapphire disc exposure to the extreme temperature. If this method fails after 2–3 attempts of dipping, go back to step C3a and make sure that the resin from the sides of the sapphire discs is removed completely. This should allow a smooth removal of the sapphire disc from the resin block.
Pre-screening of samples and serial-sectioning of a selected specimen
Cells are located at the surface of the resin blocks (Kiewisz et al., 2021). Therefore, it is easy to pre-screen cells for subsequent ultramicrotomy. Although a standard light microscope can be used for this purpose, rare stages like prometaphase or early anaphase are hard to spot with certainty when relying solely on the different refractive indices within the resin. A fluorescence microscopy approach can be applied to obtain high-resolution image stacks that allow for identifying the desired stages. To achieve this, methylene blue or other histological dyes (Kiewisz et al., 2021) are added during freeze-substitution to specifically stain the chromosomes. This approach enables the screening of cells of interest before sectioning the specimens.
Using an upright transmitted light microscope, screen the surface of the resin block to locate cells that are rounded and show any mitotic hallmarks, e.g., condensed chromosomes aligned at the metaphase plate. Image the area of interest with a microscope camera or through the eyepiece with a smartphone. This step is necessary as the transmitted light microscope can only be used to observe cells that are relatively shallow.
For a better relocation of cells of interest, it is advised to image the area at different magnifications (Figure 5B and 5C).
Figure 5. Pre-screening of resin-embedded HeLa cells using methylene blue. (A) Cartoon showing assembly of resin embedded cells on the microscope stage. (B) Overview image of the region of interest (red box) using an upright light microscope. (C) Higher-magnification image of the region of interest is shown in B. The cell of interest is indicated by a white arrow. (D) Low-magnification fluorescence image of resin-embedded HeLa cells stained with methylene blue. The region of interest is indicated by a red box and corresponds to the area of interest as shown in B. (E) Higher-magnification image of the region of interest as shown in D. The cell of interest is indicated by a white arrow. (F) Razor blade-trimmed resin block containing the cell of interest. (G) Electron micrograph of a 300 nm section of the selected cell. (H) Corresponding fluorescence light microscopic image of the cell of interest. (I) Overlay of the fluorescence light and electron microscopic images, as displayed in G and H. This figure was reproduced and modified with permission from the book chapter: “High-throughput screening of mitotic mammalian cells for electron microscopy using classic histological dyes”, Methods in Cell Biology (162), 151–170 (2021), Copyright Elsevier [Kiewisz et al. (2021)].
Position a resin block with the specimen side facing the objective of a confocal microscope (such as the Leica SP5 or Leica SP8 Stellaris). It is advised to use an inverted microscope for this and place the resin block on top of a 24 mm × 60 mm coverslip. The block should be placed on the glass slide with the cells facing down and submerged in a droplet of water (Figure 5A). We found that using a 594 nm excitation laser and detecting the emitted light in the range of 620–730 nm yields the best results for methylene blue as a dye. Make sure to relocate areas of interest by identifying landmarks such as the distribution pattern of the embedded cells or any distinctive features (e.g., dirt).
Collect images of the putative areas of interest and validate if the areas contain the cells of interest (Figure 5D–5E) by investigating the stage of the selected cells. At this point, damaged cells can be excluded from further processing.
Re-localize a region of interest using either an upright brightfield microscope or the microscope attached to the ultramicrotome (Figure 6A–6C).
Trim resin block surface for serial sectioning. We use razor blades for this step. Start with a rather large surface area (Figure 6D; Video 2).
Again, re-localize the region of interest using an upright brightfield microscope (Figure 6E–6G).
Repeat steps D6–D7 until the sample has been trimmed to the appropriate size (Figure 5F and Figure 6H).
On an ultramicrotome, equipped with a diamond knife, overfill the boat of the diamond knife with water.
Add 1–3 droplets of 70% ethanol.
Remove some water from the boat with a syringe until a bright reflection is visible at the edge of the diamond.
Move the trimmed specimen very carefully to the edge of the knife.
Cut 300 nm sections with a speed of 1.6 mm/s. The cutting speed can vary by ± 0.3 mm/s, depending on the room temperature and humidity as well as the polymerization stage of the sample (Figure 6I–6J).
Move ribbons of sections with hair glued to the tip of a toothpick. We prefer to use Dalmatian dog hairs for this procedure. However, any other non-sticky type of hair can be used for this.
Collect ribbons of sections on Formvar-coated copper slot grids (Figure 6K).
Figure 6. Serial sectioning of selected cells in resin-embedded samples. (A) Top view showing a partially trimmed resin block. Trimming of the block is achieved using a razor blade. (B) Top view of the same resin block as shown in A. The image was taken by using a stereomicroscope at 5× magnification. The selected cell is indicated by a white arrow. (C) Zoom-in view of the area of interest at 25× magnification as shown in B. The selected cell is indicated with a white arrow. (D) Rough trimming of the resin block around the area of interest. (E) Top view of the further rough trimmed area around the cell of interest as seen by using a binocular microscope. (F) Top view of the same resin block as shown in E imaged using a stereomicroscope at 5× magnification. The arrow indicates the cell of interest. (G) Zoomed-in view of the area of interest at 25× magnification with the indicated cell of interest white arrow as shown in F. (H) The top view of the fully trimmed trapezoid-shaped block face of the resin sample. (I) Top view onto the boat of a diamond knife showing a ribbon of serial sections attached to the knife edge. (J) Top view onto the boat of a diamond knife showing five ribbons of serial sections. At this stage, it is crucial to keep the correct sequence of the ribbons. (K) Same view as given in J to illustrate how a single ribbon is collected on a Formvar-coated copper slot grid (see also Video 2).
Video 2. Video tutorial showing how to perform serial-section ultramicrotomy. Trimming of a resin block containing a selected cell is achieved by using a razor blade. This is followed by serial sectioning of the trimmed area using a diamond knife. This video showing serial sectioning is speeded up. This video corresponds to Figure 6.
Post-stain sections with uranyl acetate and lead citrate and image sections by using a TEM (Figure 5G).
Add fiducial markers to the sections by immersing them into a droplet of colloidal gold (Ø 20 nm) solution for 1 min. Then, blot excessive fluid with filter paper and air dry the grid for 1 min. Wash grids for 1 min in double-distilled water and let them air dry before storing them in a grid box.
Inspect the serial sections on the grids by using a TEM operated at 80 kV. This process allows to inspect and validate the selected cells by overlaying the electron micrograph with the fluorescence image previously taken (see step D3; Figure 5G–5I).
After this control step, image-select cells with a TEM operated at 300 kV and equipped with a tilting stage (Mastronarde, 1997) with SerialEM software. For a detailed tutorial for acquiring tilt series tomography with this software, we refer to the online tutorial (https://bio3d.colorado.edu/SerialEM/).
Load the grid with the serial sections into a tomography sample holder and insert it into the TEM.
Open SerialEM, create a new map file, and mark the position of a cell of interest on each section on the grid. Make sure to adjust the Eucentric height at all positions as this will speed up the acquisition process.
Using the SerialEM pre-exposure feature, expose each selected position on each section to 2,000 e-/Å2 at a 60° tilt angle for 5 min. This baking process allows the resin section to shrink to a stable level before the image acquisition is started.
Set up serial acquisition in SerialEM to acquire a tilt series for each region at ± 60° with 1° increments at 4,700× magnification (a-axis tilt series).
After all acquisitions are finished, rotate the grid by 90° and repeat steps D19b and D19d to record the b-axis tilt series.
Automatic reconstruction of electron tomogram and microtubule segmentation
Following the tomographic acquisition, each tilt series file has to be reconstructed to obtain an in-silico 3D volume. This process can be time consuming, which is why we chose to fully automate this process. Tomogram processing is carried out with the IMOD software package. The IMOD software package is a set of image processing, modeling, and display programs for tomographic reconstruction and 3D reconstruction of EM serial sections. Automation scripts and instructions for an automatic tomographic reconstruction with IMOD are available on GitHub (https://github.com/RRobert92/Assit_Automatic_Tomogram_Flattening_IMOD). For full documentation and a tutorial on how to use IMOD, we refer the reader to the IMOD tutorial (https://bio3d.colorado.edu/imod/doc/etomoTutorial.html).
Automatic tomogram reconstruction:
To start an automatic tomogram reconstruction, open the etomo package from the IMOD software and follow step-by-step instructions from File S3.
Open the batch processing mode.
Load the automatic reconstruction setting available on GitHub (https://github.com/RRobert92/Assit_Automatic_Tomogram_Flattening_IMOD/blob/main/Kiewisz_Imod_batch_reconstruction.adoc).
Select raw data files (tilt series). Each pair of tomogram files (a- and b-axis) should be stored in a separate folder.
Choose a device (CPU, GPU, or CPU + GPU) on which the tomogram will be processed.
After pressing Run, the tomographic reconstructions will be calculated and stored on the hard drive. If the message shows completed, proceed to step E1g. If the message shows failed, one needs to check the step on which the algorithm aborted the reconstruction and fix it manually.
Post-process reconstructed tomograms by trimming the histogram and rotating the volume around the x-axis.
Run an automatic flattening tool from GitHub (https://github.com/Robert92/Assit_Automatic_Tomogram_Flattening_IMOD/blob/main/AAFT.bat) to flatten each tomogram.
Use the new stack command from IMOD in the terminal to manually remove z-planes from the 3D stack that do not contain image information.
Semi-automatic microtubule segmentation: as previously published (Redemann et al., 2014), microtubules are automatically segmented using the commercial Amira or AmiraZIBEdition software package. The Amira software package is a set of image processing and visualization tools that allows for precise and intuitive image segmentation of big data. A tutorial of this process is shown in Figure 7 and File S4. For a full tutorial on the filament segmentation see https://assets.thermofisher.com/TFS-Assets/MSD/Product-Guides/user-guide-amira-software.pdf.
Figure 7. Semi-automatic segmentation of microtubules in whole spindles obtained from electron tomography of serial sections (300 nm). (A) View of the Amira graphical user interface (GUI) showing the steps to initialize semi-automatic microtubule prediction by computing correlation and orientation fields. To initialize the procedure, select the Cylinder Correlation tool (shown in red box 1, right panel), and select parameters for microtubule template matching (shown in the left panel, red box 2). To start the calculation, press the Apply button indicated with red box 3. (B) GUI showing steps to track microtubules from template matching, generated in A. To initialize microtubule tracing from the generated template, use the Trace Correlation Lines tool (red box 1, right panel) and select parameters for microtubule tracing (shown in the left panel, red box 2). To start tracing, press the Apply button indicated with red box 3. (C) GUI showing the final spatial graph file containing coordinates for the detected microtubule tracks. The red box indicates all traced microtubule filaments stored in .am Amira spatial graph file format.
Load reconstructed tomogram volume into the Amira software (Figure 7).
Select the Cylinder Correlation tool (Figure 7A, red box 1) and select the setting as shown in Figure 7A (red box 2).
Press the green button Apply to start the calculation (Figure 7A, red box 3).
After the calculation is finished, two new files are produced (Figure 7B).
Select the Trace Correlation Lines tool. Then, select the two new files as inputs and use the setting as shown in Figure 7B (red box 1–2).
Press the green button Apply to start the calculation (Figure 7B, red box 3). At the end of the calculation, a new file will be created called a spatial graph containing the predicted microtubule tracks (Figure 7C). Save the file locally on the hard drive.
Be aware that Amira allows the generation of so-called Recipes. Within these Recipes, one could save the configuration of certain modules like Cylinder Correlation and Trace Correlation Lines. Therefore, one has only to apply the recipe to the raw tomogram to obtain the spatial graph as a resulting output, which saves time and prevents human errors in configuring the parameters of the modules. We recommend using the properly configured Recipes when segmenting microtubules if the tomogram quality is not strongly varying.
Correction of microtubule tracks predicted with the Amira software. During the prediction of microtubule tracks from electron tomograms, several errors can occur as false-positive or false-negative microtubule tracks. This correction needs to be done manually and therefore can take up to several hours or a full working day per single tomogram (Figure 8A–8B). Each acquired tomogram needs to be corrected.
Figure 8. Plot indicating the time needed for microtubule segmentation using the Amira software including semi-automatic segmentation and manual correction by experienced users. Segmentation time is shown in full days referring to 24 h time intervals. (A) Bar plot showing the segmentation time per single tomogram (n = 195). (B) Bar plot showing the segmentation time per entire data set (n = 14). Each data set is composed of 8–37 tomograms.
For correction, open the filament tab in the Amira software (Figure 9, red box).
Figure 9. Manual correction of semi-automatically segmented microtubules using the AmiraZIBEdition software. GUI showing an overview of the filament editor tool (red box, top). The orange box indicates the different tools that can be used to correct the segmentation, the blue box shows visualization parameters, and the pink box indicates the stepping tool used to step through all individual microtubules.
In an open new window, one can now see, side-by-side, the tomogram and all predicted microtubule tracks.
On the top of the open window (Figure 9, orange box), one can see all the editing tools. By highlighting with the mouse cursor, one can read the descriptions and the keyboard shortcuts associated with them.
On the top-left side of the open window (Figure 9, blue box), one can see the visualization setting of the spatial graph that controls the thickness of displayed lines.
On the bottom-right side of the open window (Figure 9, magenta box), one can see the editing tool where one can switch between segmented microtubules.
Correct the spatial graph by removing all false-positive segments and adding the missing (false-negative) microtubule tracks.
Remove the false-positive microtubules (Figure 10A) by selecting them in the segment box menu (Figure 9, magenta box) and pressing D (for delete) on the keyboard. To add a new microtubule track, firstly press C and then T, and select the center of the microtubule by clicking on it within the left image viewer. This will create the first node. To extend the newly created microtubule track, press N, and while pressing the control key, continue to select and click in the center of the microtubule to draw a line until reaching the end of it.
Figure 10. Illustration of microtubule tracing correction using the AmiraZIBEdition software package as shown in Figure 9. (A) Example of false-positive filament annotation caused by the application of the semi-automatic segmentation approach. The left panel shows a cross-section view of a tomogram with selected microtubules highlighted in red and white arrow. The right panel shows an overview of all traced microtubules with selected microtubules highlighted in red and white arrow. (B) View from the AmiraZIBEdition software as shown in A. Example of a correctly tracked microtubule. (C) View from the AmiraZIBEdition software as shown in A. Example of manually and fully corrected filament annotation.
Check if the microtubule track is correct and if the ends of the segment are positioned at the ends of the microtubule in the image. One can do that by moving through the Z-planes with the mouse wheel up and down.
If the selected microtubule track looks correct, one can simply skip to the next one (Figure 10B) until all microtubules have been corrected and all false positive segmentations have been deleted (Figure 10C).
Stitching of segmented serial sections. The final step of microtubule segmentation is the stitching of all serial sections together to one tomographic volume with all microtubules stitched together to one spatial graph. For this, we used a research version of Amira called AmiraZIBEdition comprising the SerialSectionStack tool (Lindow et al., 2021). This tool is also available on GitHub (https://github.com/zibamira/SerialSectionAligner or https://www.zib.de/software/serial-section-aligner) as an extension for Amira v2020.3 or higher.
Open Amira with the installed extension or AmiraZIBEdition and create the SerialSectionStack tool (Figure 11A, red box 1).
Figure 11. Stitching of serial sections by using the AmiraZIBEdition SerialSectionStack. (A) The main view of the AmiraZIBEdition software GUI. The red box number 1 indicates the data management window with the open SerialSectionStack tool. The red box number 2 indicates the button used to load all serial section data for stitching. (B) GUI view as shown in A. Loaded datasets in the correct order are indicated in the red box. (C) View of AmiraZIBEdition software GUI showing open SerialSectionAligner tool setting window. The left panel shows the zoomed area as shown in the blue box in the right panel. The numbered red boxes show the sequence of tasks that have to be done to initialize the SerialSectionAligner tool: No. 1, Advanced button; No. 2, Switch to matching mode; No. 3, Switch slice quality to high and the warping quality to the maximum; and No. 4, Compute the alignment of the transformation matrix for the first section by pressing Compute. The green box shows the alignment view window, allowing the user to observe and improve the alignment. (D) To compute the alignment transformation matrix for the next section, the user needs to switch to the next border using the buttons indicated in the red box.
From the project tab (bottom left corner, Figure 11A, red box 2) select add files and select all tomographic volumes with corresponding spatial graph fields. The files should be named with consecutive numbering e.g., Tomogram_1.am, Tomogram_sg_1.am; Tomogram_2.am, Tomogram_sg_2.am (Figure 11B, red box), and they should be all in the same folder.
With the right mouse button, press the SerialSectionStack tool and open the SerialSectionAligner tool. This will update the properties tab view (Figure 11C, right panel, green box).
Initialize this tool by pressing Advanced (Figure 11C, red box 1) and switch to matching mode (Figure 11C, red box 2). Enable a high-quality display for tomographic slices and increase the warping quality to the maximum (Figure 11C, red box 3).
Next, press compute (Figure 11C, red box 4). This will calculate the alignment transformation matrix for the first section based on matching microtubules from the first and the second section. When finished, the computation view window (Figure 11C, right panel, green box) will update to show the aligned two consecutive sections. Repeat this step a few times until the histograms (shown in the bottom right corner) do not change anymore.
Repeat step E4e for each section by switching to the next section interface (Figure 11D, red box) and pressing compute.
When the computation of the alignment matrix for all sections based on microtubule continuity between sections is finished, press create at the bottom of the properties view. One can optionally deselect crop to filament. This option, when enabled, allows the tool to crop tomogram volume to the spatial graph size and produce a smaller tomogram volume (Figure 12A).
At the end of the calculation, a user obtains two files (Figure 12B, red box 1–2): the Amira spatial graph with all stitched microtubules and a 3D volume of the stitched serial sections (Figure 12C–12D).
Figure 12. Correction of stitched microtubules from all serial sections using AmiraZIBEdition SerialSectionStack. (A) After repeating the computation for all borders in between sections (see Figure 11D), the user can compute the stacked volume for all aligned sections using the computed alignment transformation matrix by pressing Create (red box). (B) At the end of the step shown in A, two new files are generated. The stitch tomogram volume (red box number 1) and corresponding file with all stitched microtubules (red box number 2). The file as indicated by the red button is the Amira Spatial Graph file that contains tracks for microtubules stitched from all serial sections. (C) View from AmiraZIBEdition as shown in A, indicating visualization setting. (D) View from AmiraZIBEdition showing a tomographic slice and all stitched microtubules using the setting as shown in C.
As an important final step, check and correct each stitched microtubule. For this, repeat step E3 for the generated stitched files.
Data analysis
Analyses
In this step, we present an Automatic Spatial Graph Analysis tool (ASGA) that was developed for rapid local and online analysis of segmented microtubule tracks (Figure 13A–13C; Video 3). ASGA is a fully automated package with an intuitive GUI. After loading the data, the users only have to select the type of analysis that should be performed on the microtubule tracks. The entire code is developed in R and hosted as an online service via R shiny package. In the following part, we present all available analyses in ASGA (links and GitHub: https://github.com/RRobert92/ASGA; Kiewisz and Müller-Reichert, 2022b).
Figure 13. Workflow for fully automatic microtubule analysis with ASGA. (A) The home screen of the ASGA software. The red box indicates the button used to start the analysis. (B) Loading of data into the ASGA software. The red box shows a window where users can select either single or multiple datasets at once. (C) After the data compatibility has been checked, the ASGA software welcomes the user with a setup window. This window allows users to customize their analyses. A selection of the options is needed at this point. The red box number 1 allows one to select the type of analysis. The user can switch on or of specific types of analysis. The red box number 2 indicates a button to start the selected type of analysis (Kiewisz et al., 2022; see also Video 3).
Video 3. Tutorial video describing the usage of the ASGA software v0.36. This video corresponds to Figure 13.
Data format: the ASGA workflow is operating on the ASCII Amira file format. This file can be exported from the AmiraZIBEdition package and contains information about each microtubule spline. In addition, for the analysis of the kinetochore microtubules (KMTs), the file should have separated microtubules into KMTs and non-KMTs. We have built in the ASGA workflow a file format error handler that checks each input data structure and gives clear information if any of the files is not compatible with ASGA and provides hints on how to fix it.
Fiber and KMT length distribution: the length of each microtubule is calculated as shown in Eq. 1. Furthermore, length distribution is calculated as a function of all lengths (Eq. 1).
Eq 1. Equation calculating the length of microtubule track (L) using 3D points data, where i indicates the point belonging to the microtubule track and di,i+1 indicates the 3D Euclidian distance between two consecutive points.
Statistical analysis of KMT number: ASGA performs a statistical analysis of the KMT number and compares it against the kinetochore position, the sister k-fiber KMT number, and the distance between the sister k-fiber microtubule plus-ends.
Outer-kinetochore distance and distribution: the distance between outer kinetochores is calculated as the distance between sister k-fiber plus-ends. A k-fiber plus-end is taken as a mean 3D position of all KMT plus-ends. K-fiber sisters can be indicated by the user in the spatial graph (see the New tools development paragraph) or automatically detected. Automat detection of k-fiber sisters is achieved by firstly estimating the kinetochore position with KMT plus-ends and then selecting a sister kinetochore that is facing the same direction.
Fiber and KMT curvature: to calculate KMT or k-fiber total curvature, ASGA measures the local and total tortuosity (Eq. 2). Tortuosity is the measurement of the line deformation and can be between 1 and infinity, where 1 indicates a straight line. For example, 1.57 indicates the curvature of a half-circle. It is calculated as a ratio of the true microtubule length as calculated in Eq. 1 and the length of a straight line between its endpoints. This measurement can be used as a quantification of microtubule curvature indicating the deviation from a straight line. To calculate the tortuosity of a k-fiber, ASGA computes the center of a k-fiber as a mean of all microtubules in the fiber, and the k-fiber tortuosity is a measure based on this center of a k-fiber.
Eq 2. Equation calculating microtubule tortuosity, where L indicates the total length of the microtubule calculated as in Eq1, and l indicates the microtubule length as a straight line between its endpoints.
KMT twist: the local KMT twist is calculated along KMT in steps of 500 nm along the microtubule lattice. For each step, the twist is calculated as a clockwise or anti-clockwise angle of rotation (Eq. 3). The total twist is calculated as a sum of all local twists along k-fiber. For this, at each step, we calculate the mean twist value from each KMT present in the k-fiber, and the total k-fiber twist is given as a sum of all local twists along the k-fiber (Eq. 4). This means that, for example, if the k-fiber is randomly twisted, the total twist will be close to 0°, and if the k-fiber is in its entire length negatively twisted, the total twist of a k-fiber will be for example -25°.
Eq 3. Equation calculating local microtubule twist between two consecutive points on microtubule tracks, where x and y indicate 2D vectors between the center of the k-fiber and KMT at a position i and i+1.
Eq 4. Equation calculating summed microtubule twist along k-fiber, where i indicate the number of steps along which the local twist is calculated.
K-fiber helicity: K-fiber helicity is calculated by firstly computing the k-fiber center and measuring the total twist of a k-fiber relative to the pole-to-pole axis. Helicity is then calculated as a k-fiber total twist over its length as shown in (Eq. 5).
Eq 5. Equation calculating k-fiber helicity, where the numerator indicates the total twist of a k-fiber based on a k-fiber center relative to the pole-to-pole axis, and i is the fiber length calculated with Eq1.
Fiber area: to calculate the fiber cross-section area, ASGA uses the alpha shape algorithm. This algorithm allows one to fit a convex hull polygon into the cross-section of the k-fiber. This approach allows for precise retrieval of the cross-sectional area of the k-fiber. The fiber area is calculated locally every 500 nm, or as the total fiber area given as a mean value of all local areas.
KMT neighborhood density: to calculate the local density of the KMTs in a k-fiber, the ASGA software computes the local fiber area at the specified position. The density is then derived as the number of KMTs divided by fiber area at a given position (Eq. 6).
Eq 6. Equation calculating k-fiber local density, where n indicates the number of KMTs at a given position m.
KMT branching: to detect branching of microtubules, ASGA computes the distance matrix between each KMT minus-end and every microtubule spline (Eq. 7). Next, for each minus-end, ASGA selects the closest microtubule spline. Finally, KMT branching from either microtubules or KMTs is detected by selecting KMT minus-ends, which are within a specified user distance (default: 25 nm) to the closest microtubule spline. ASGA gives out a list of KMT branches with KMT ID, microtubule ID from which KMTs branched, and the distance of KMT minus-end to the microtubule spline.
Eq 7. Equation calculating the distance between the KMT end and every other microtubule spline, where n is the number of microtubules in a system, i indicates KMT 3D end position, and j indicates microtubule spline.
Global microtubule–microtubule interactions: to identify every microtubule–microtubule interaction, ASGA computes a global distance matrix. In short, ASGA computes the distance of each microtubule to each other microtubule within a given threshold distance (default: 100 nm). Further, ASGA gives information about microtubule IDs for which the interaction was observed, for the position on the microtubule where the interaction occurred, and for the length of interaction.
New tools development
The ASGA raw code compiled packages for Windows, MacOS, or Linux is available on GitHub (Kiewisz and Müller-Reichert, 2022b; https://github.com/Robert92/ASGA). The online version of the platform is available on GitHub.
ASGA standardized data input. As with any other software, ASGA requires standardized data input. ASGA requires the input data to be in the form of an Amira Spatial Graph file format saved in the ASCII format. ASGA recognizes KMTs by detecting the KMT label; other microtubules are recognized as non-KMTs. Optionally, the user can specify extra labels, thus classifying which KMTs belong to which k-fiber. To ensure accurate computation of microtubule positions in relation to the spindle axis, it is necessary to re-orient the spindle such that both spindle poles marking the pole-to-pole axis have the same x-coordinate in the spatial graph. Furthermore, for increased accuracy in the analysis, the spatial graph should be re-sampled to a uniform point distance of at least 20 nm. The ASGA 3DViewer expects as input either CSV or XSLX file formats with all ASGA analyses as well as the corresponding Amira file spatial graph.
Spatial graph pre-processing and new tool development. ASGA performs data initialization during the loading of a spatial graph, which further standardizes the input data for all other tools to operate on. Each spatial graph is assigned an ID number, which the developer can refer to when computing multiple spatial graphs one by one. Additionally, each spatial graph is divided into three classes: non-KMTs, KMTs, and k-fibers. The optional k-fiber class is created if the spatial graph contains information for each KMT about its association with a particular k-fiber. To each microtubule, a unique ID is assigned. Developers can retrieve either individual microtubules or a set of microtubules belonging to a k-fiber by calling these IDs. By calling a specific microtubule ID, the developer retrieves access to a sorted indexed list of XYZ coordinates, where the first and last coordinate is associated with the ends of each microtubule. In addition, a developer can use the indexed list to mark only a specific area of a microtubule.
As for the code of conduct, developers are advised to develop a function that operates on either an individual or a subset of microtubules and then wrap this function over the desired classes. This approach may not always be optimized for speed. However, it allows new developers not familiar with the code to easily create new tools that can be rapidly implemented and deployed.
Visualization
The ASGA-3DViewer raw code is available on GitHub (https://github.com/RRobert92/ASGA_3DViewer; Kiewisz and Müller-Reichert, 2022a). The online version of this tool was also made available (http://cfci.shinyapps.io/ASGA_3DViewer/). The usage of this ASGA tool is shown in Figure 14A–14F and Video 4. In short, upon opening the online tool, the user is welcome to the simple and intuitive GUI to enter the main tool and select the data set of interest. After quick pre-loading of the selected data set, the user can interactively visualize the entire spindle data set, choose from different visualization options, and perform an overlay color-coded analysis.
Figure 14. Workflow for microtubule visualization using the ASGA-3DViewer web server. (A) Home screen view of the ASGA-3DViewer. The red box indicates the button initiating the tool. (B) Screenshot of the ASGA-3DViewer indicating the selection of the visualized data set. The red box indicates different data sets from which users can choose. (C) View of ASGA-3DViewer setting panel. The red box shows the selection of the sub-dataset available in the selected collection. The orange box indicates the selection of which type of microtubule class is currently visualized. By default, the ASGA-3DViewer shows only KMTs. However, all microtubules can also be selected. The light blue box indicates the k-fiber and color that is displayed. By default, all fibers are shown in red color. The dark blue box indicates the analyses that can be optionally overlaid on the selected microtubules. The green box indicates a button that is used to apply all selected changes. (D) Example of visualization shown in ASGA-3DViewer showing all kinetochore microtubules. (E) Example of the ASGA-3DViewer showing the settings in the left panel and spindle visualization of all KMTs with overlaid analysis in the right panel. (F) Another example from the ASGA-3DViewer showing selected k-fiber (Dataset – HeLa_#1; K-fiber ID - Pole1_14). This illustrates microtubule–microtubule interaction within a 35 nm distance (Kiewisz et al., 2022; see also Video 4).
Video 4. Tutorial video describing the usage of the ASGA-3DViewer software v1.3.1. This video corresponds to Figure 14.
Validation of protocol
Robert Kiewisz, Gunar Fabig, William Conway, Daniel Baum, Daniel Needleman and Thomas Müller-Reichert, (2022). Three-dimensional structure of kinetochore-fibers in human mitotic spindles. eLife (11): 1–37. DOI: 10.7554/eLife.75459.
William Conway, Robert Kiewisz, Gunar Fabig, Colm P Kelleher, Hai-Yin Wu, Maya Anjur-Dietrich, Thomas Müller-Reichert and Daniel J Needleman, (2022). Self-organization of kinetochore-fibers in human mitotic spindles. eLife (11): 1–34. DOI: 10.7554/eLife.75458.
Alejandra Laguillo-Diego, Robert Kiewisz, Carlos Martí-Gómez, Daniel Baum, Thomas Müller-Reichert and Isabelle Vernos, (2023). MCRS1 modulates the heterogeneity of microtubule minus-end morphologies in mitotic spindles. MBoC (34): 1–14. DOI: 1091/mbc.E22-08-0306-T.
General notes and troubleshooting
3D printing: The designed specimen chambers (Figure 1) can be printed using any commercially available 3D printer supplied with either biodegradable PLA or PLA/PHA plastic. The type of material used for 3D printing should have no effect on the cultivation of the cells. In addition, to avoid any potential risk of contamination with lead, we suggest changing the printing nozzle from the standard brass nozzle to a stainless-steel nozzle.
Mitotic shake-off: Make sure to prepare everything in advance that is necessary for the shake-off. The shake-off should be performed as fast as possible to minimize the time during which the cells are exposed to cold temperatures. T75 flasks should always be stored upward (with the lid cup facing upward) to avoid cooling down the cells by the cold bench. Use a long 25 mL pipette to remove or add fresh prewarmed medium to speed up the process. Optionally, you can store a metal plate in the incubator heated up to 37 °C, which can be used to help maintain the temperature of the cell culture.
Acknowledgments
We would like to thank the Dipl.-Ing. Silke Tulok and Dr. Anja Nobst of the Core Facility Cellular Imaging (CFCI, Faculty of Medicine Carl Gustav Carus, TU Dresden) for help with light microscopy and Dr. Tobias Fürstenhaupt of the EM facility at MPI-CBG (Dresden) for technical assistance with electron tomography. Figures 1–5 were reproduced from the article published in Methods in Cell Biology, Vol 162, Robert Kiewisz, Thomas Müller-Reichert and Gunar Fabig, Chapter 7: High-throughput screening of mitotic mammalian cells for electron microscopy using classic histological dyes, 151–170, Copyright Elsevier (2021). Research in the Müller-Reichert laboratory was supported by funds from the Deutsche Forschungsgemeinschaft (DFG grant MU 1423/8–2 and /10–1 to TMR). RK received funding from the European Union’s Horizon 2020 research and innovation program under the Marie Skłodowska-Curie grant agreement No. 675737 (grant to TMR), the Simons Foundation (grant No. SF349247 to Tristan Bepler) and NIH National Institute of General Medical Sciences (grant No. GM103310 to Tristan Bepler).
Competing interests
All authors disclose no competing interests.
References
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Kiewisz, R. and Müller-Reichert T. (2022a). RRobert92/ASGA_3DViewer: 3D Viewer for ASGA (v1.3.1). Zenodo.
Kiewisz, R. and Müller-Reichert. T (2022b). RRobert92/ASGA: Automatic Spatial-Graph Analysis (ASGA) v0.37 (v0.37). Zenodo.
Kiewisz, R., Fabig, G., Conway, W., Baum, D., Needleman, D. and Müller-Reichert, T. (2022). Three-dimensional structure of kinetochore-fibers in human mitotic spindles. eLife 11: e75459.
Kiewisz, R., Müller-Reichert, T. and Fabig, G. (2021). High-throughput screening of mitotic mammalian cells for electron microscopy using classic histological dyes. Methods Cell Biol. 162: 151–170.
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Ma, H. T. and Poon, R. Y. C. (2017). Synchronization of HeLa Cells. In: Banfalvi, G. (Ed.). Cell Cycle Synchronization (pp. 189–201). Methods in Molecular Biology. Humana Press, New York.
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Supplementary information
The following supporting information can be downloaded here:
File S1. File in.stl format as used for 3D printing of the incubation chambers for a single sapphire disc optimized for performing shake-off experiments. This supplementary information corresponds to Figure 1A.
File S2. File in.stl format as used for 3D printing of the incubation chambers for four sapphire discs optimized for performing shake-off experiments. This supplementary information corresponds to Figure 1B.
File S3. Protocol for the custom-designed fully automatic generation of tomograms using IMOD.
File S4. Detailed explanation of each parameter used for the semi-automatic segmentation of microtubules in Amira. This supplementary information corresponds to Figure 7.
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Correlative Conventional and Super-resolution Photoactivated Localization Microscopy (PALM) Imaging to Characterize Chromatin Structure and Dynamics in Live Mammalian Cells
DM Dushyant Mehra
EP Elias M. Puchner
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4850 Views: 521
Reviewed by: Alessandro DidonnaRevati Sumukh DewalAftab Nadeem
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Original Research Article:
The authors used this protocol in Nucleic Acids Research May 2022
Abstract
A fundamental understanding of gene regulation requires a quantitative characterization of the spatial organization and dynamics of chromatin. The advent of fluorescence super-resolution microscopy techniques such as photoactivated localization microscopy (PALM) presented a breakthrough to visualize structural features with a resolution of ~20 nm in fixed cells. However, until recently the long acquisition time of super-resolution images prevented high-resolution measurements in living cells due to spreading of localizations caused by chromatin motion. Here, we present a step-by step protocol for our recently developed approach for correlatively imaging telomeres with conventional fluorescence and PALM, in order to obtain time-averaged super-resolution images and dynamic parameters in living cells. First, individual single molecule localizations are assigned to a locus as it moves, allowing to discriminate between bound and unbound dCas9 molecules, whose mobilities overlap. By subtracting the telomere trajectory from the localization of bound molecules, the motion blurring is then corrected, and high-resolution structural characterizations can be made. These structural parameters can also be related to local chromatin motion or larger scale domain movement. This protocol therefore improves the ability to analyze the mobility and time-averaged nanoscopic structure of locus-specific chromatin with single-molecule sensitivity.
Keywords: Photoactivated localization microscopy PALM Single-molecule tracking Nanoscale Chromatin structure and dynamics Live-cell chromatin imaging CRISPR/dCas9 DNA-labeling MS2 gRNA Live-cell super-resolution imaging
Background
Gene expression is thought to be regulated by the spatiotemporal organization of chromatin from the smallest length scale of individual nucleosomes (~10 nm) up to ~100 nm [1,2]. Even larger tertiary structures such as enhancer promoter contacts or topologically associated domains may exist that regulate gene expression [1,3]. The correlated movement between small chromatin structures and the large chromatin domains they are part of has been suggested as an important feature of nuclear phase condensates [4–6]. To understand these effects of chromatin structure and dynamics on gene regulation, imaging techniques are required that can characterize both the nanoscopic structure and the dynamics of chromatin in the larger context of chromatin domains they reside in.
Recently, two main breakthroughs facilitated research advances in this field: CRISPR/dCas9-based fluorescence labeling methods, to image specific sequences of chromatin, and super-resolution microscopy techniques such as photoactivated localization microscopy (PALM). By using programmable guide RNAs (gRNA), fluorophores can be targeted via CRISPR/dCas9 to specific sequences in the genome. Tens of fluorescent probes are required to be bound to a locus of interest to create a signal that is distinguishable from the background fluorescence of all other freely diffusing and searching probes [7]. To amplify the fluorescence signal of bound probes, repetitive RNA aptamers such as MS2 sequences have been attached to gRNAs that facilitate binding of multiple fluorophores. These labeling strategies facilitated conventional fluorescence timelapse imaging and yielded valuable insights into the slow and long-term dynamics of entire loci [7–9]. However, due to the optical diffraction limit, the structural characterization of smaller chromatin structures below ~250 nm has not been possible with conventional fluorescence microscopy. The development of PALM [10–12] enabled the tracking of single molecules in living cells (also referred to as single-particle tracking PALM, sptPALM) [13,14] and the acquisition of images of intracellular structures in fixed cells with ~20 nm resolution [15]. In PALM imaging, the spatiotemporal overlap of individual fluorophores is avoided by sparsely activating photoactivatable or photoswitchable fluorophores. In this way, the precise locations of individual fluorophores can be determined by Gaussian fitting of their intensity profile. By photoactivating and localizing many or all fluorophores over time, enough localizations are obtained to resolve structures below the optical diffraction limit in fixed cells or to link molecular trajectories to characterize their diffusion in live cells. CRISPR/dCas9-based DNA labeling has recently been applied in PALM to obtain structural information of chromatin compaction or condensation in chemically fixed cells or to characterize the dynamics of chromatin in live eukaryotic and prokaryotic cells [16]. However, until recently, it was not possible to simultaneously obtain such structural and dynamic information in living cells due to the motion of DNA during the long PALM data acquisition time. This motion spreads out the localizations of bound fluorophores along the trajectory of a locus and thus results in motion blurring.
Here, we present a protocol for our recently developed correlative conventional fluorescence and PALM imaging approach in living cells that overcomes these challenges [17]. This approach is based on labeling an intracellular structure or locus with a conventional fluorophore to track its location and motion during the entire PALM data acquisition time. Each structure or locus is, in addition, labeled with a spectrally distinct PALM-compatible fluorophore to record the single molecule localizations. The trajectory of the locus determined from the conventional fluorescence signal is then subtracted from the coordinates of its single molecule localizations to correct for motion blurring. As a result, high-resolution structural parameters such as the time-averaged size or the density of bound probes can be quantified to obtain new insights into the compaction of chromatin. In addition, the dynamics of the relative single molecule rearrangement or the motion of the entire locus can be related to its structural parameters, which has not been possible with existing techniques.
We demonstrate correlative conventional and PALM imaging using the well-characterized telomere sequences as a model system labeled via dCas9 and the MS2 coat proteins (MCP) that bind to a modified telomere-targeting gRNA scaffold. However, this imaging approach can be extended to single loci or other intracellular structures that can be labeled with a sufficient number of conventional and PALM-compatible fluorophores to create signals above background. Importantly, our presented data acquisition and analysis pipeline is a primary step to any downstream analysis to quantify structural or dynamic parameters. For instance, we demonstrate that determining the location and mobility of a locus relative to the single dCas9/MCP fluorophores classifies them more reliably as bound to a locus. The relative motion of single molecules compared to the entire locus furthermore reveals how small-scale chromatin rearrangements occur within the larger-scale chromatin movements. We also relate the compaction of telomeres to the local and global chromatin mobility to yield new insights. This protocol demonstrates that correlative conventional fluorescence and PALM imaging accurately identifies Cas9 molecules bound to a locus and yields quantitative dynamic and time-averaged structural information about specific genomic loci at the nanoscale in living cells. The versatility of this protocol makes it applicable to other organelles and enables other existing or future downstream analysis techniques to extract and correlate high-resolution structural features with dynamic parameters.
Materials and reagents
Lab-Tek No. 1.5 8-well plates (Fisher Scientific, catalog number: 12-565-8)
Lipofectamine 3000 and p300 reagent (Invitrogen, catalog number: L3000001)
Opti-MEM media (Thermo Fisher, catalog number: 31985070)
MCP-HaloTag plasmid (Addgene, catalog number: 121937)
dCas9-GFP plasmid (Addgene, catalog number: 51023)
2xMS2 gRNA plasmid (Addgene, catalog number: 75389)
PA-JF646 (Luke Lavis Lab, HHMI Janelia)
GIST-T1 Cells (gastrointestinal stromal tumor cells) (Cosmo Bio, catalog number: PMC-GIST01C)
Note: We used this cell line as it is hypothesized that this cancer phenotype is impacted by changes in chromatin structure and dynamics [18]. Furthermore, living GIST-T1 cells can be imaged for long periods of time and exhibit minimal auto-fluorescence and cell death.
T25 tissue culture flask (Thermo Fisher, catalog number: 156340)
Phenol-red free trypsin EDTA (Gibco, catalog number: 15400054)
Fetal bovine serum (FBS) (Gibco, catalog number: 10437028)
Fluorobrite DMEM (Gibco, catalog number: A1896701)
Penicillin/Streptomycin (Gibco, catalog number: 15140122)
L-Glutamine (Gibco, catalog number: 25030-081)
TetraSpeck microspheres (Invitrogen, catalog number: T7279)
1.7 mL Eppendorf tubes (catalog number: 0030123611)
37 °C 5% CO2 incubator (Thermo Fisher, catalog number: 3110 or similar)
DI water (e.g., Thermo Fisher, catalog number: 751-610 or purified in house)
Distilled phosphate buffered saline (PBS) (Gibco, catalog number: 14040133)
Solutions
Fluorobrite media (see Recipes)
Serum-diluted Fluorobrite media (see Recipes)
DNA/lipid mixture (see Recipes)
Recipes
Fluorobrite media
10% FBS, 4 mM L-Glutamine, 1% penicillin/streptomycin, Fluorobrite DMEM. For 50 mL of media, add 5 mL of FBS, 500 μL of penicillin/streptomycin solution, 500 μL of L-Glutamine, and 44 mL of Fluorobrite DMEM.
Serum-diluted Fluorobrite media
1% FBS, 4 mM L-Glutamine, 1% penicillin/streptomycin, Fluorobrite DMEM solution. For 50 mL of media, add 500 μL of FBS, 500 μL of penicillin/streptomycin solution, 500 μL of L-Glutamine, and 48.5 mL of Fluorobrite DMEM.
DNA/lipid mixture
Mix 200 ng of telomere 2xMS2 gRNA along with 50 ng of MCP-HaloTag and 50 ng of dCas9-GFP plasmids with 10 μL of Opti-MEM, 1 μL of Lipofectamine 3000 reagent, and 0.5 μL of p300 reagent in a 1.7 mL Eppendorf tube in sterile cell culture environment.
Note: Telomere gRNA sequence was obtained from [7], and protocols from [8,19], and [9] were used to clone telomere gRNA sequence into 2xMS2 plasmid.
Equipment
Four OBIS lasers emitting 100 mW at 405 nm (Coherent, catalog number: 1178754), 50 mW at 488 nm (Coherent, catalog number: 1178764), 100 mW at 561 nm (Coherent, catalog number: 1253302) and 100 mW at 640 nm (Coherent, catalog number: 1178790)
Beam expander (Thor Labs, catalog number: GBE02-A)
Assorted lenses and mirrors (Thor Labs)
Inverted microscope (Eclipse Ti-E) equipped with a perfect focus system (Nikon, catalog number: MEA53100)
CFI 100× 1.49 NA oil immersion objective (Nikon, catalog number: MRD01991)
iXon 897 Ultra DU-897U EMCCD camera (Andor, catalog number: 77026047)
Quad band dichoric mirror (Chroma, catalog number: ZT405/488/561/640rpc)
Bandpass filters for the green (Chroma, catalog number: ET525/50), red (Chroma, catalog number: ET595/50), and far-red channel (Semrock, catalog number: FF731/137)
Dichroic long pass beam splitter for red/green channel experiments (Chroma, catalog number: T562lpxr BS)
Dichroic long pass beam splitter for far red/green channel experiments (Semrock, catalog number: FF652-Di01)
Motorized flat top stage for inverted microscope (ProScan II, catalog number: 77011328)
Heating insert P for Lab-TekTM S1 and temperature controller (Pecon, catalog number: 411860-9025-000 and 411860-9005-000)
Computer for microscope control and data acquisition (e.g., Dell, model: Optiplex 9020 Mini-Tower, Intel Core i7-4790 CPU @3.60GHz 4 cores, 16 GB RAM, 3.64 TB drive)
Computer for data analysis (e.g., Dell, model: PowerEdge T440, Intel Xeon Silver 4216 2.1G, 16C/32T, 9.6 GT/s, 22 M Cache, 16 GB RDIMM, 3,200 MT/s, Dual Rank, 8 TB 7.2K RPM SATA 6Gbps 512e 3.5 in Hot-plug Hard Drive)
Software
MATLAB 2018b
Insight3 Localization Software (Huang lab, UCSF or Zhuang lab, Harvard) or equivalent
Storm Control Software (https://github.com/ZhuangLab/storm-control) or equivalent
MATLAB-based trace linking and trace analysis (https://osf.io/6n4ej/)
MATLAB-based motion correction and trace separation code (https://osf.io/6n4ej/)
Python-based channel transformation code (https://osf.io/6n4ej/)
Procedure
GIST-T1 cell culture, seeding, and plasmid transfections
Seed GIST-T1 cells in 5 mL of Fluorobrite media at a density of 33% (~400,000–600,000 cells) in a T25 tissue culture flask.
Note: Fluorobrite media avoids fluorescence from phenol red.
Culture cells for ~1–2 days up to a density of 75%–80% in a humidified incubator at 37 °C and 5% CO2. Split cells by aspirating media from the flask in a cell culture hood and washing cells with 37 °C PBS twice.
After removing PBS, add 0.5 mL of phenol-red free trypsin EDTA to the flask and make sure the entire surface area of the flask is covered.
Then, place flask in incubator for 2–3 min to allow cells to lift from flask. Verify that cells have lifted.
Add 2 mL of Fluorobrite media to trypsinized cells to neutralize trypsin and add cells at density mentioned in step A1 with 5 mL of Fluorobrite media or proceed with step A6.
Seed 50 μL of trypsinized cells in 8-well plates at a density of 50,000 cells/mL two days prior to imaging. Add 400 μL of media to each well after plating cells, and culture as in step A1.
Note: Cell concentration was measured using a hemocytometer, and Fluorobrite media was used to dilute cells to find appropriate concentration.
Approximately 15–17 h before imaging, mix 200 ng of telomere 2xMS2 gRNA plasmid to generate gRNA along with 50 ng of MCP-HaloTag and 50 ng of dCas9-GFP plasmids with 10 μL of Opti-MEM, 1 μL of Lipofectamine 3000 reagent, and 0.5 μL of p300 reagent in a 1.7 mL Eppendorf tube and incubate the DNA/lipid mixture for 15 min at room temperature in a sterile cell culture environment.
During incubation of step A7, aspirate media from GIST-T1 cells in a well plate and wash cells twice with 300 μL of Fluorobrite media heated to 37 °C. Then, add 100 μL of Fluorobrite media supplemented with 4 mM L-Glutamine, 1% FBS, 1% penicillin/streptomycin, and 200 μL of Opti-MEM. Then, place cells in an incubator at 37 °C with 5% CO2 for 15 min.
After 15 min of incubation in step A8, remove cells from incubator, place in cell culture hood, and add all of the DNA/lipid mixture from the Eppendorf tube to cells in each well already supplemented with serum-diluted media by pipetting dropwise. If too much force is used, DNA/lipid complex may disassociate.
Place wells with transfecting cells in the incubator as in step A2 for 15–17 h.
Remove media from wells and wash cells twice with serum-diluted Fluorobrite media. Add 300 μL of serum-diluted Fluorobrite media with 100 nM of PA-JF646 dye and incubate for 15 min in a 37 °C incubator with 5% CO2.
Note: PA-JF646 is a far-red fluorescent dye that is photoactivated by 405 nm light and contains a ligand that attaches to HaloTag [20,21]. This dye has a higher photon budget and longer on-times compared to other photoswitchable proteins, which results in improved detection, localization precision, and longer single molecule trajectories. This improves trace mobility analysis and diffusion coefficient estimation. This dye isn’t fluorogenic and does fluoresce when not bound to HaloTag, which is why multiple rounds of washing are required before imaging.
After incubation, wash cells three times with Fluorobrite media and place in an incubator at 37 °C with 5% CO2 for an additional 30 min.
Repeat step A12 three additional times prior to imaging to remove unbound PA-JF646 dye that is still able to fluoresce.
Keep samples in an incubator at 37 °C with 5% CO2 until imaging.
Calibration and imaging experiments
Note: All experiments presented here were performed with a custom-built microscope, as recently described [22]; however, our protocol is applicable to data recorded with any microscope capable of simultaneous PALM and conventional fluorescence imaging in the respective channels. In short, a Nikon inverted microscope (Eclipse Ti-E) is equipped with a perfect focus system and an Andor iXon 897 Ultra DU-897U electron multiplying charge coupled detector (EMCCD), cooled to -70 °C and set to an amplifying gain of 30. The four 100 mW excitation lasers (405, 488, 561, and 640 nm OBIS-CW, Coherent Optics) are aligned, expanded, and focused into the back focal plane of the objective (Nikon CFI 100× 1.49 NA oil immersion) using various dichroic mirrors, beam expanders, and lenses. A quad band dichoric mirror (ZT405/488/561/640rpc; Chroma) separates fluorescence emission from excitation light. Fluorescence emission is further split into the far red and green signal using a dichroic long pass beam splitter (FF652-Di01; Semrock) and band pass filters FF731/137 (Semrock) for the far-red channel and ET525/50 (Chroma) for the green channel. Laser intensity modulation and shutter sequences are synchronized with the camera and controlled digitally with a NI-DAQ board.
Mount microscope No. 1 cover glass or 8-well plates with 10 μL of TetraSpeck microspheres diluted 1:100 in DI water on microscope stage.
Excite microspheres separately using 640 and 488 nm excitation with approximately 1.75 mW power (power density ~100 W/cm2 in sample plane). Record a 100-frame movie with approximately 10 sparsely distributed microspheres over the entire field of view without moving the sample and use the first 50 frames to record 640 nm excited microspheres and the last 50 frames to record 488 nm excited microspheres.
Move the same microspheres to different positions in the camera to sample the entire field of view and repeat steps B2 and B3 at least five times. You can also image different microspheres placed in different regions across the field of view.
Note: The number of microspheres used depends on the field of view of the camera. Our field of view was 256 × 256 pixels with a pixel size of 160 nm. You need at least 50 microspheres to create an accurate transformation with sub 20 nm registration error (see Figure 1). More microspheres or more images with microspheres shifted to different locations are necessary if your field of view is larger. Microsphere imaging can also be done after cellular imaging but should be done either before or after every imaging session, since registration parameters can change from session to session. This data will be used later to transform localizations from 640 nm channel to the 488 nm channel.
Figure 1. Example microsphere localization rendering. Example rendering of microsphere localizations across seven movies depicting the number and density of microspheres required to obtain an accurate transformation across the field of view. (A) Microspheres imaged in each channel to calculate the transformation matrix. (B) The transformation matrix is applied to transform bead localizations from the Jf646 channel to the GFP channel. This transformation matrix is then applied to single molecule localizations. Scale bars: 5 µm.
Remove microsphere sample and mount 8-well plate with transfected cells on microscope stage. Make sure the stage is heated to 37 °C and the CO2 incubator on microscope stage reads 5%.
Using the Storm Control Software (or equivalent for a different microscope system), set up a 10-frame laser shutter sequence at 20 Hz with the first frame having 488 nm excitation for GFP imaging, the second having brightfield LED and 405 nm photoactivation, and frames 3–10 having 640 nm excitation for PALM imaging.
Note: The diffusion coefficient distributions of MCP-HaloTag with and without telomere gRNA taken at 20 Hz are statistically similar to the ones obtained at 60 Hz. While faster PALM imaging speeds in principle reduce motion blurring, the required higher excitation power reduces the length of single molecule traces, and thus lowers the precision for characterizing their motion. In our experience, a 20 Hz frame rate has been a good compromise between imaging speed, localization precision in each frame, and single molecule trace lengths.
Set the 488 nm laser power to 1.75 mW (power density ~100 W/cm2 at sample plane) and the 640 nm to 17.5 mW (~1 kW/cm2). The 405 nm intensity will be adjusted to 1–251 μW (power density of ~0.06–15 W/cm2) during the experiment for constant photoactivation rates.
Turn on the 488 nm laser to visually identify cells with telomere puncta in cell nucleus prior to single molecule imaging. Keep 488 nm exposure to a minimum to reduce bleaching. Image telomere puncta conventionally using the 488 nm laser for at least 200 frames at 20 Hz for interpolation error analysis.
Note: Cells transfected with all three plasmids should show clear telomere puncta. Since cells are transiently transfected, some cells express all transfected plasmids while others do not. Look at cells that do not have gRNA but have MCP-HaloTag+PA-JF646 dye and dCas9-GFP expressed, and cells stained with only PA-JF646 dye and no transfected plasmids to characterize single molecule and conventional fluorescence background (see Figure 2).
Figure 2. Examples of fluorescence images from transfected cells suitable and unsuitable for experiments. (A) Example of cellular autofluorescence. (B) and (C) Examples of cells expressing dCas9-GFP at various levels but without distinct telomere puncta due to the lack of gRNA. This can also occur with gRNA added. (D) and (E) Examples of telomere puncta at various expression levels suitable for imaging and analysis. Nuclei are marked with red line. Scale bars: 5 μm.
Once cells with telomere puncta are identified, begin 10-frame shutter and acquisition sequence. Start with 405 nm laser power of 1 μW (power density of ~0.06 W/cm2) and slowly increase at a rate of 5–10 μW every 1,000 frames until approximately 250 μW (power density of ~0.015 W/cm2) to ensure a sparse and consistent photoactivation rate (see Figure 3).
Figure 3. Optimal photoactivation density. (A) Single frame of an optimal photoactivation density for single molecule tracking with low false linking rate. Yellow boxes are identified localizations from single-molecule emission events. (B) Example images of the maximal allowable localization density for single molecule tracking with acceptable false linking. (C) Examples of images with high-localization density that would result in a high false linking rate. False linking rates for example images are quantified in Figure 5. Scale bar: 5 μm.
Record data for 15,000–30,000 frames and stop sequence when cells start to change morphology indicating a decline of cell health (see Figure 4).
Note: Healthy GIST-T1 cells never change their morphology up to 15,000 frames. Only analyze and use frames up to 5,000 frames before morphology changes are detected, to exclude potential phototoxic effects prior to morphological changes. If cells start to change morphology sooner, cells are not healthy enough to begin with for imaging experiments (see Figure 4).
Figure 4. Cell morphology and health. LED + 405 nm photoactivation frames of healthy GIST-T1 cells (top) typically show retained morphology and size for at least 15,000 frames and show minimal autofluorescence upon 405 nm activation. Unhealthy cells (bottom) change morphology and become round within several thousand frames and show significant autofluorescence in response to 405 nm activation. Healthy cells also show much less contrast in the LED frame due to a more homogenous refractive index. Scale bars: 5 μm.
Repeat steps B5–B9 to obtain more statistics from different cells. Typically, a number on the order of 10 cells is considered satisfactory.
Data analysis
Single molecule localization
Use Insight3 localization software or a similar single molecule localization microscopy software to identify single molecule localizations and to fit them with 2D elliptical Gaussians with the following parameters: 7 × 7 pixel ROI, widths between 250 and 700 nm, and minimum intensity of 100 photons. The x and y coordinate of each localization, along with the intensity, width, background, frame number, and other parameters are also stored in the single molecule list and outputted as a .txt file for further analysis.
Note: Other localization detection software, such as Thunderstorm or SMAP, can also be used with similar parameters [23,24]. Make sure to export localization list as .txt file and to include PSF widths.
Import molecule list into provided MATLAB analysis package or equivalent single particle tracking software. If using Insight3 for localization detection, insert file path into LoadMoleculeLIst.m and execute function to generate a molecule list structure that will be used for subsequent analysis steps.
Use Thompson resolution formula or more accurate and updated version [25,26] to calculate localization precision of single molecule localizations. If using Insight3 to localize images, input molecule list into function ThompsonResolution.m to calculate localization resolution for each localization.
Note: The localization precision is provided in most localization software packages, such as Thunderstorm or SMAP along with localizations.
Single molecule trace linking and trace analysis
To perform trace-linking error analysis, execute the MATLAB function spatiotemporal_cc.m and provide the molecule list structure and desired frame range for the analysis. Use this function to measure pairwise distance of localizations in same frame (see Figure 5).
Note: This function normalizes the number of pairwise distances in each bin by the area and number of frame pairs. This modified pair correlation function quantifies the number of molecules found around each localization in the same frame. The number of localizations at your desired linking distance in the same frame will give you the estimate for the false linking rate.
Figure 5. Localization density and false linking rate. False linking rate calculations for examples in Figure 3. These plots depict the average number of localizations as a function of distance away from a localization in the same frame. The false linking rate can be estimated by obtaining the number of localizations at a specific trace linking distance. (A) Optimal false linking rates for 0.48 μm linking distance is between 0.01 and 0.05. (B) A linking error at 0.48 μm of approximately 0.1 is considered a high false linking rate and will lead to unreliable mobility measurements.
Exclude data from further analysis if localization density is too high and if the false linking rate is above 0.1. If false linking rate is too high throughout the duration of the experiment, record data with lower 405 nm photoactivation power in step B8.
Note: If you want to allow dark frames for linking a single molecule trace, incorporate the number of dark frames in the pair correlation metrics. For example, if you use one dark frame, then you need to calculate pairwise distances between every two frames to calculate false linking error.
Based on results from step 1, link localizations that are within the determined distance threshold. In the datasets presented here, a 0.48 μm linking distance was used. Only traces with a minimum of four localizations in consecutive frames are used for cross correlation and single trace fitting analysis. Input the data structure from LoadMoleculeList.m, linking distance (in μm), and number of dark frames into the MakeTraces.m function to link molecules across successive frames into traces. Any equivalent trace linking analysis package can also be used.
Calculate mean squared displacements (MSD) for each time step in all single molecule traces and average MSDs for each timestep of a single trace to obtain a time-averaged mean squared displacement (TAMSD) vs. time plot. If using the provided MATLAB code, input the output data structure from MakeTraces.m into function DiffusionDisplacment.m and execute DiffusionDisplacement.m.
Fit each TAMSD vs. time to the 2D diffusion equation <r2> = 4DΔt + 2σ2, where D is the diffusion coefficient, Δt is the time step, r2 is the TAMSD, and σ is the localization precision (see Figure 6). Only use trajectories and diffusion coefficients from fits with a coefficient of determination value (R2) of 0.7 or above and a diffusion coefficient above zero. If using the provided MATLAB code, execute DiffusionCoefficientAnalysis.m function, which calculates the diffusion coefficient and R2 value for each input TAMSD calculated from DiffusionDisplacement.m.
Figure 6. Time-averaged mean squared displacement (TAMSD) fitting. (A) Examples of acceptable TAMSD fits. These fits have R2 values of above 0.7, positive slopes, and y intercepts above 0. The y intercept can be used to calculate localization precision. (B) Examples of excluded TAMSD fits. These fits have either R2 values of below 0.7, negative slopes, or y intercepts at or below 0.
GFP cluster identification and trace linking analysis
Obtain GFP localizations by fitting GFP clusters with 2D Gaussians as in step A1 from Data analysis with the following parameters: 11 × 11 pixel ROI, widths between 250 and 7,000 nm, and a minimum of 200 photons. X and y coordinates of the GFP localization, along with the intensity, width, background, frame number, and other parameters will again be stored in the molecule list.
Note: Even though the width range is large, make sure widths from the same telomere do not deviate significantly. If the GFP localization widths from the same telomere deviate by more than ~200 nm, it indicates that the telomere is moving along z too much and cannot be used for further analysis. If using other localization software such as SMAP or Thunderstorm, make sure PSF widths are included in the output molecule list.
To link telomere GFP localizations that are within 0.48 μm of each other in consecutive conventional imaging frames (every 10 frames), repeat steps B3–B5 with the GFP localization list.
Only use traces with a minimum of five localizations for downstream analysis. The widths of consecutive localizations in a trace should be within 200 nm to be included in downstream analysis. If using steps B3–B5, this will automatically be done in the motioncorrection.m function.
Note: An axial microsphere calibration [22] showed that PSFs width deviations of 200 nm corresponded to axial deviations of 450 nm, which is similar to the lateral trace linking threshold. In our analysis, we found that traces with below five localizations had insufficient single molecule localizations for downstream motion correction analysis.
Perform a linear interpolation between the x and y coordinates of GFP localizations in consecutive conventional image frames (frame 1 and frame 10) to obtain interpolated GFP coordinates during the frames that contained single molecule localizations. If using steps B3–B5, this will automatically be done in the motioncorrection.m function.
To estimate the upper limit of the interpolation error, analyze the data of step B7 of the calibration and imaging experiments in the same way as steps B1–B4 of Data analysis. Compare interpolated positions between frames nine frames apart to the actual position of the telomere.
Note: The median interpolation error of the presented data is 45 ± 10 nm and the mean interpolation error is constant up to 20 frames (Supplementary Figure 4 in reference [17]).
Calculate TAMSD and diffusion coefficients using the same procedure described in step B5 of Data analysis. Only use the first four time steps for fitting analysis to exclude nonlinear portions of the TAMSD that occur at later time steps. In this way, non-Brownian diffusion is approximated and sub-diffusive behavior at long times is not given too much weight in the fit.
Motion correction of single molecule localizations
Localize microspheres from calibration images recorded in steps B1–B3 in both channels using Insight3 localization software or equivalent with parameters used to localize single molecules in step A1 of Data analysis.
Fit the positions of the microspheres in both channels to a third order polynomial function to extract the coordinate transformation matrix between the two channels. If using the provided procedure, input the two molecule lists and execute the python polynomial transformation code bead_calibration.py [27,28].
Apply transformation to top channel to superimpose localizations from 640 nm channel to bottom channel. If using Insight3 localization software, go to STORM math in the STORM panel, click the custom math function, and copy the output transformation equations from the python transformation code into the storm math text box. Then, in display layer options under the view tab, click the drift correction box to view the transformed localizations. Transformed localizations will show up as XC and YC in the exported Insight3 localization software molecule list text file (see Figure 7).
Figure 7. Single molecule localization microscopy (SMLM) localizations (red) before transformation (A) and after transformation (B) superimposed on GFP telomere cluster. The transformation is also used to correct for chromatic aberrations in the optical 4F emission path. Localizations that are not properly transformed cannot be accurately registered with GFP localizations and motion corrected. Apparent incomplete colocalization is due to telomere motion and is corrected during the motion correction analysis. Scale bars: 5 µm.
Calculate a distance matrix between the interpolated GFP localizations for a specific telomere and all single molecule localizations in that frame. If using the provided MATLAB code (motioncorrection.m) this will automatically be done in the cross-correlation section of the motion correction code. The code will loop through all accepted GFP trajectories.
The cross-correlation section of the provided motion correction MATLAB code (motioncorrection.m) identifies single molecule localizations whose distance to a GFP cluster is smaller than the radius of the cluster plus the localization precision of the cluster and the single molecule localizations. The radius of the last GFP localization prior to interpolation should be used as the radius of the interpolated GFP cluster coordinates. The motion correction code (motioncorrection.m) will classify single molecule traces with a minimum of four localizations and with all localizations residing within a GFP cluster as bound and will exclude single molecules traces where some but not all localizations or no localizations reside within a cluster.
To correct for motion of telomeres in PALM images, the motion correction section of provided MATLAB motion correction code (motioncorrection.m) will subtract the coordinates of GFP localizations at a specific frame from the initial GFP localization in the trajectory and apply that subtraction to the single molecule localizations belonging to each GFP cluster and the same frame (see Figure 8).
Note: A minimum of four bound trajectories with a minimum of four localizations each should be used to obtain enough localizations to be able to motion correct localizations within a telomere trajectory and to calculate downstream metrics such as area and density.
Figure 8. Motion correction example. (A) Superposition of a GFP image and conventional PALM image that includes a majority of freely diffusing and searching fluorescent probes. (B) The correlative conventional and PALM image only depicts PALM localizations that appear in proximity to a GFP cluster at any instance in time and suppresses background from freely diffusing and searching probes. (C) The motion-corrected PALM image super-resolves each moving telomere, which colocalizes with its GFP signal. Scale bar: 5 µm.
Apply a convex hull to the motion-corrected localizations in a cluster using the boundary function in MATLAB (convhull.m) to find the cluster boundary and use this boundary to calculate cluster area.
If calculating localization density, normalize the number of localizations by the cluster duration in the field of view to account for constant photoactivation rate.
Perform desired downstream analysis of structural and dynamic parameters. For instance, correlate cluster density or area with cluster mobility to correlate chromatin structural information with dynamic information. Since single molecule trajectories are assigned to specific GFP telomere trajectories, the diffusion coefficient of the single molecule trajectories can be compared to the mobility of GFP trajectories which provides meaningful information on chromatin rearrangements (see examples in Figure 9). Bulk trace analysis methods such as Gaussian mixture models, Bayesian inference techniques, or displacement analysis techniques can also be applied to trajectories.
Figure 9. Secondary downstream analysis examples of motion-corrected data. (A) The localization density of individual telomeres has a slight correlation (correlation coefficient = 0.4) with their diffusion coefficient determined with the conventional GFP signal. This result is plausible, since denser, more compacted telomeres can diffuse more freely than less dense and more extended telomeres, whose motion is slowed down. (B) The ratio of average dCas9-MCP single molecule and the telomere diffusion coefficients they reside in presents a metric of relative single molecule re-arrangement and shows a negative correlation (correlation coefficient = -0.52) with the normalized localization density of the telomeres. (C) Likewise, the ratio of average dCas9-MCP diffusion coefficients show a positive correlation (correlation coefficient = 0.48) with the area of the telomeres they reside in. These results are plausible since denser, more compacted telomeres have less ability for relative motion or re-arrangement due to tighter packing compared to less dense, more extended telomeres, which exhibit more relative mobility. Data adapted from Mehra et al. (2022) [17].
Acknowledgments
The authors thank Angel Mancebo Jr for writing the two-channel microsphere calibration code and helpful discussions. The authors acknowledge Bo Huang for providing some of the PA-JF646 dye and helpful discussions. The authors acknowledge Tamas Ordog and Yujiro Hayashi for providing the GIST-T1 cells. The authors thank Stephen C. Ekker and Karl J. Clark for providing lab space, reagents, and guidance on molecular cloning. The authors thank Jacob Ritz for the helpful discussions. This work was supported by funding from the National Institutes of Health under award number R21GM127965 and from the Mayo Graduate School of Biomedical Sciences and Mayo Foundation. This protocol is used in Mehra et al. (2022) [17].
Competing interests
The authors declare that no competing interests exist.
Ethical considerations
GIST-T1 Cells were obtained from Dr. Tamas Ordog.
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4,851 | https://bio-protocol.org/en/bpdetail?id=4851&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Human Dendritic Cell Subset Isolation by Magnetic Bead Sorting: A Protocol to Efficiently Obtain Pure Populations
GF Georgina Flórez-Grau
JE Jorge Cuenca Escalona
HL Helena Lacasta-Mambo
DR Daphne Roelofs
JB Johanna Bödder
RB Ruben Beuk
GS Gerty Schreibelt
IV I. Jolanda M. de Vries
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4851 Views: 831
Reviewed by: Chiara AmbrogioWendy Leanne Hempstock Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal for ImmunoTherapy of Cancer Apr 2022
Abstract
Dendritic cells have been investigated for cell-based immunotherapy for various applications. The low abundance of dendritic cells in blood hampers their clinical application, resulting in the use of monocyte-derived dendritic cells as an alternative cell type. Limited knowledge is available regarding blood-circulating human dendritic cells, which can be divided into three subsets: type 2 conventional dendritic cells, type 1 conventional dendritic cells, and plasmacytoid dendritic cells. These subsets exhibit unique and desirable features for dendritic cell-based therapies. To enable efficient and reliable human research on dendritic cell subsets, we developed an efficient isolation protocol for the three human dendritic cell subsets, resulting in pure populations. The sequential steps include peripheral blood mononuclear cell isolation, magnetic-microbead lineage depletion (CD14, CD56, CD3, and CD19), and individual magnetic-microbead isolation of the three human dendritic cell subsets.
Graphical overview
Scheme of the dendritic cell (DC) isolation protocol. Starting material for this process is human blood (buffy coat or aphaeresis). From that, peripheral blood mononuclear cells (PBMCs) are isolated by using ficoll gradient centrifugation. Then, an enrichment for DCs is performed using semi-automated equipment. From the enriched fraction, DC subsets are obtained by magnetic cell sorting.
Keywords: Dendritic cell subsets Human blood Sequential isolation Immunotherapy Magnetic microbeads
Background
During the past decades, dendritic cells (DCs) have been investigated for their ability to initiate antigen-specific immune responses in vivo. DC-based immunotherapies have been developed or are under investigation for the treatment of various malignancies such as cancer (Gonzalez et al., 2018), autoimmune disorders such as rheumatoid arthritis, or multiple sclerosis (Collin and Bigley, 2018). Most of the current DC knowledge is from the human DC model, monocyte-derived DCs (moDCs), which have a very high purity and can be generated in significant numbers from peripheral blood mononuclear cells (PBMCs).
Recently, attention shifted towards the utilization of the three human DC subsets circulating in blood, due to their unique and distinct functions compared with moDCs (Wang et al., 2020). Blood-circulating human DCs can be broadly divided into plasmacytoid DCs (pDCs) and conventional DCs (cDCs). cDCs are highly specialized in antigen uptake, processing, and (cross-)presenting of antigens to naïve T cells, which is a crucial step for initiating immune responses. cDCs can be subdivided into type 1 (cDC1s) and type 2 (cDC2s). In humans, cDC1s express CD141 (BDCA-3), cDC2s express CD1c (BDCA-1), and pDCs express CD123 (BDCA-2). Each DC subset presents a unique function; for example, cDC1s can potently take up apoptotic cells, cross-present externally derived antigens, and activate cytotoxic lymphocytes (Schreibelt et al., 2012). These features place cDC1s in the center of interest for mounting anti-tumor immune responses. Thus, it is necessary to investigate the phenotypical and functional differences between human DC subsets in depth, with the goal of improving DC-based therapies.
Unfortunately, investigating the human DC subsets is a rather tedious task, mainly due to their scarcity in PBMCs, ranging from < 0.2% of PBMCs in the case of cDC2s and pDCs to < 0.08% of PBMCs in the case of cDC1s (van Beek et al., 2020). In addition, currently available human DC isolation protocols result in varying levels of cell impurity or cross-contamination with other human DC subsets. Contaminating cells present in DC cultures can greatly influence the phenotypical and functional outcome of DC research (van Beek et al., 2020), hampering the elucidation of the role of individual human DC subsets.
We have developed a protocol to isolate highly pure populations of human DC subsets starting from PBMCs. Using the protocol described here, DC subsets can be isolated with higher purity and yield compared with currently available options such as cell sorting or magnetic-microbead kits without the lineage depletion step. The protocol consists of multiple steps including depletion of monocytes (CD14+), B cells (CD19+), T cells (CD3+), and NK cells (CD56+) from PBMCs with magnetic microbeads, followed by DC isolation using magnetic microbeads specific for each DC subset: cDC2s (CD1c+), cDC1s (CD141+), and pDCs (CD304+). We present the protocol with the use of semi-automated equipment (MultiMACS Cell24 Separator Plus, Miltenyi Biotec) that significantly reduces the time needed for the DC isolation process.
Materials and reagents
Beads and items for cell separation
Anti-CD3 microbeads (Miltenyi Biotec, catalog number: 130-050-101)
Anti-CD14 microbeads (Miltenyi Biotec, catalog number: 130-050-201)
Anti-CD19 microbeads (Miltenyi Biotec, catalog number: 130-097-055)
FcR blocking reagent (Miltenyi Biotec, catalog number: 130-059-901)
Anti-CD56 microbeads (Miltenyi Biotec, catalog number: 130-050-401)
Anti-CD1c biotin (Miltenyi Biotec, catalog number: 130-119-475)
Anti-biotin microbeads (Miltenyi Biotec, catalog number: 130-090-485)
Anti-CD141 microbeads (Miltenyi Biotec, catalog number: 130-090-532)
LD columns (Miltenyi Biotec, catalog number: 130-042-901)
MS columns (Miltenyi Biotec, catalog number: 130-042-201)
LS columns (Miltenyi Biotec, catalog number: 130-042-401)
Multi-24 column blocks (8×) (Miltenyi Biotec, catalog number: 130-095-691)
Single-well deep-well plates (Miltenyi Biotec, catalog number: 130-114-966)
Buffers and media
Human albumin (Sigma-Aldrich, catalog number: H0900000)
PBS (Life Technologies, catalog number: 14190-094)
Human serum (Sigma-Aldrich, catalog number: H4522-100ML)
X-VIVO15 hematopoietic cell culture medium (Life Technologies, catalog number: BE02-060Q)
UltraPureTM 0.5 M EDTA, pH 8.0 (Thermo Fisher, catalog number: 15575020)
Ammonium chloride (Sigma-Aldrich, catalog number: 213330)
Potassium bicarbonate (Sigma-Aldrich, catalog number: 237205)
Disodium EDTA (Sigma-Aldrich, catalog number: E4884)
Bovine serum albumin (Thermo Fisher, catalog number: 23209)
Sodium azide (Merck, catalog number: RTC0000068- 1L)
Trypan Blue (Sigma-Aldrich, catalog number: T6146-25G)
Diluting buffer (see Recipes)
ACK lysis buffer (see Recipes)
Washing buffer (see Recipes)
PBA (see Recipes)
Antibodies
Anti-CD20 FITC (BD Biosciences, catalog number: 345792)
Anti-CD14 PerCP (BioLegend, catalog number: 325632)
Anti-CD141 APC (Miltenyi Biotec, catalog number: 130-090-907)
Anti-CD1c PE (Miltenyi Biotec, catalog number: 130-113-864)
Anti-CD123 APC (BD Biosciences, catalog number: 560087)
Anti-BDCA2 PE (Miltenyi Biotec, catalog number: 130-097-929)
Materials
Ficoll lymphoprep (VWR, catalog number: CLUT1114547)
Tuerk’s solution for leukocyte counting (Merck, catalog number: 1092770100)
Tubes (Stem Cell Technologies, catalog number: 100-0088)
50 mL tubes (Corning, catalog number: 430828)
96-well V-bottomed plate (Thermo Fisher, catalog number: 277143)
Microbeads, antibodies, X-VIVO15 hematopoietic cell culture medium, washing buffer, and PBA were stored at 4 °C. Diluting buffer, PBS, and ACK lysis buffer were stored at RT.
Note: Products and equipment from other vendors (when available) can also be used.
Equipment
MultiMACS Cell24 Separator Plus (Miltenyi Biotec, catalog number: 130-098-637)
QuadroMACS Separator (Miltenyi Biotec, catalog number: 130-090-976)
OctoMACS Separator (Miltenyi Biotec, catalog number: 130-042-109)
FACSVerse (Becton Dickinson)
Note: This FACS has three lasers and eight colors (four in the blue laser, two in the red laser, and two in the violet laser).
Centrifuge (Hettich centrifuge, model: rotanta 460R)
Cell counting chamber (Thermo Fisher, catalog number: C10228)
Shaker (Thermo Fisher, SHKE4000-7: MaxQ)
Software
FACS verse software BD (BD FACSVerseTM Systems)
FlowJo (FlowJoTM v10.8)
Procedure
From a buffy coat/aphaeresis:
Peripheral blood mononuclear cell (PBMC) isolation
Isolate PBMCs following a ficoll-based protocol (Corkum et al., 2015).
Note: From the PBMCs isolation, a cell suspension (with a mixture of different immune cells) is obtained.
After washing the cells three times with 1 mL of washing buffer (PBS, 0.1% BSA, EDTA), count the total amount of isolated PBMCs. For accurate counting, use Tuerk’s dye.
Notes:
In this protocol, washing the cells refers to centrifuging at 252× g for 5 min at 4 °C.
Counting is performed by using a cell counting chamber and a microscope, as shown in Figure 1.
Figure 1. Schematic protocol for how to count cells
Resuspend PBMCs in 1 mL of 1× ACK lysis buffer per 100 × 106 cells.
Note: Resuspend by pipetting up and down.
Incubate for 5 min at room temperature (RT).
Add washing buffer to the ACK-cell suspension, filling up to 50 mL.
Note: Resuspend by pipetting up and down.
Centrifuge PBMCs at 252× g for 5 min at 4 °C. After centrifugation, the cells are in a pellet at the bottom.
Discard supernatant (by pouring it).
Proceed with the cell pellet to Section B.
Note: The cell pellet is resuspended by adding the lineage depletion mixture (Section B), followed by up-and-down pipetting.
Lineage depletion
Prepare the lineage depletion master mixture (Caution: This mixture is prepared using the PBMC count numbers from step A2):
Anti-CD3 microbeads at 1 μL per 1 × 106 PBMCs.
Anti-CD14 microbeads at 1 μL per 1 × 106 PBMCs.
Anti-CD19 microbeads at 1 μL per 1 × 106 PBMCs.
Anti-CD56 microbeads at 0.5 μL per 1 × 106 PBMCs.
Anti-FCR blocking at 1 μL per 1 × 106 PBMCs.
Add the depletion master mixture to the PBMC pellet and resuspend by pipetting up and down.
Incubate for 30 min at 4 °C while shaking on a shaker or manually every 5 min.
Note: Select the slowest speed of the shaker.
Add 5 mL of washing buffer to depletion mixture PBMC suspension (step B2).
Centrifuge PBMC suspension at 252× g for 5 min at 4 °C.
Discard the supernatant (by pouring it).
Resuspend the PBMCs at a concentration of 100 × 106 cells per milliliter in washing buffer.
Note: Resuspend by pipetting up and down.
Prepare the MultiMACS Cell24 Separator Plus without the elution station according to the manufacturer instructions. Initiate the preprogrammed depletion protocol.
Prewet the Multi-24 column block with 1 mL of washing buffer. Only prewet the number of columns required. Use one column per 100 × 106 PBMCs.
Place a single-well deep-well plate to recover the flowthrough containing the unlabeled cells (see Figure 2, which illustrates how the cells are collected in the system).
Figure 2. MultiMACS with the column block and the collector
Follow the indicated steps from the depletion protocol from the MultiMACS. In short, apply the PBMC suspension as 1 mL per column (Caution: The number of cells per column should be 100 × 106 PBMCs maximum). After the sample has run through the column, wash three times with 1 mL of washing buffer to ensure complete elution of target cell population.
Note: Washing the columns means adding the indicated buffer on top of the column and waiting until the fluid flows completely through.
Transfer the negative fraction (flowthrough) to a clean 50 mL tube. Centrifuge at 252× g for 5 min at 4 °C.
Note: The enriched DC fraction is the negative fraction.
Count the number of cells present in the negative fraction (with Tuerk’s dye and counting chamber).
Centrifuge at 252× g for 5 min at 4 °C.
Discard the supernatant (by pouring it).
Resuspend the pelleted cells at a concentration of 100 × 106 cells per milliliter with washing buffer.
Proceed to the final depletion step using LD columns.
Note: This step will ensure the highest purity of the negative fraction containing all DC subsets and a minute amount of other immune cells.
Prewet LD columns with 1 mL of washing buffer. Use 1 LD column per 100 × 106 cell suspension (step B13).
Run 1 mL of cell suspension per LD column and wash three times with 1 mL of washing buffer.
Collect negative fraction containing the target cell population (in a 15 mL tube, as shown in Figure 2).
Count the cells (using the counting chamber and Tuerk’s solution).
Centrifuge at 252× g for 5 min at 4 °C.
Discard the supernatant (by pouring it) and proceed with the cell pellet. The resuspension of the cell pellet will be done with the antibody mixture in Section C.
Continue with the isolation of the DC subsets.
cDC2 isolation
Add 1 μL of anti-CD1c biotin per 1 × 106 cells.
Note: Scale according to the starting cell number (step B19).
Incubate for 10 min at 4 °C while shaking.
Note: Select the lowest speed of the shaker.
Fill the 50 mL tube with washing buffer up to 50 mL.
Centrifuge at 252× g for 5 min at 4 °C.
Discard the supernatant.
Add 2 μL of anti-biotin microbeads per 1 × 106 cells.
Incubate for 15 min at 4 °C while shaking.
Note: Select the lowest speed of the shaker.
Add washing buffer up to 50 mL.
Centrifuge at 252× g for 5 min at 4 °C.
During centrifugation, prewet LS column with 1 mL of washing buffer.
Resuspend the cells in 1 mL of washing buffer.
Run the cells through the column that is placed in the magnet and wash three times with 1 mL of washing buffer.
Collect the flowthrough in a 50 mL tube (as shown in Figure 3).
Note: For the flushing step, the column should be removed from the magnet and placed in a 15 mL tube.
Figure 3. Setup of the LS column within the magnet; unlabeled cells go through, and labeled cells are retained by the magnetic field. Flush the LS column with 2 mL of washing buffer. Caution: This fraction contains the cDC2 population.
To further increase the purity of the cDC2 population, prewet one MS column with 1 mL of washing buffer.
Run the 2 mL of cDC2 cell suspension through the MS column. Wash three times with 0.5 mL of washing buffer.
Flush cDC2s out of the MS column with 2 mL of washing buffer.
Count the cDC2s (using trypan blue solution and a counting chamber, as illustrated in Figure 1).
Resuspend the cells (cDC2s) at 1 × 106 cells per milliliter with X-VIVO15 + 2% human serum and keep them at 4 °C.
cDC1 isolation
Centrifuge the flowthrough (unlabeled negative fraction) of the cDC2 isolation step (step C12) at 252× g for 5 min at 4 °C.
Discard the supernatant (by pouring it).
Count the cells (using the counting chamber and Tuerk’s solution).
Add 2 μL of anti-CD141 microbeads per 1 × 106 cells.
Incubate for 15 min at 4 °C while shaking.
Add washing buffer up to 50 mL.
Centrifuge at 252× g for 5 min at 4 °C.
During centrifugation, prewet one LS column with 1 mL of washing buffer.
Resuspend cell fraction in 1 mL.
Load the cells onto the LS column and wash the column three times with 1 mL of washing buffer.
Collect the flowthrough in a Falcon tube (50 or 15 mL).
Flush the LS column with 2 mL of washing buffer. Caution: This fraction contains the cDC1 population.
Note: For the flushing step, the column should be removed from the magnet and placed in a 15 mL tube.
To further increase the purity of the cDC1 population, prewet one MS column with 1 mL of washing buffer.
Run the 2 mL of cDC1s through the MS column that is placed in the magnet. Wash three times with 0.5 mL of washing buffer.
Flush cDC1s out of the MS column with 2 mL of washing buffer.
Count the cDC1 fraction (using trypan blue and the counting chamber).
Resuspend the cells at 1 × 106 cells per milliliter in X-VIVO15 + 2% human serum and keep them at 4 °C.
pDC isolation
Centrifuge the flowthrough (unlabeled negative fraction) from the cDC1 isolation (step D9) at 252× g for 5 min at 4 °C.
Discard the supernatant (by pouring it).
Count the cells (using the counting chamber and Tuerk’s solution).
Add 1 μL of anti-CD304 microbeads per 1 × 106 cells.
Incubate for 15 min at 4 °C while shaking.
Add washing buffer up to 50 mL.
Centrifuge at 252× g for 5 min at 4 °C.
During centrifugation, prewet one LS column with 1 mL of washing buffer.
Resuspend the cells in 1 mL of washing buffer (PBS, 0.1% BSA, EDTA).
Pass the cells through the column (add the cell suspension on top of the column that is placed in the magnet).
Discard the flowthrough (as of now, it will no longer be needed).
Run the cells through the column that is placed in the magnet and add 1 mL of washing buffer. Flush the pDC out of the LS column by washing three times with 1 mL of washing buffer and count the cells. Caution: This fraction contains the pDC population.
Note: For the flushing step, the column should be removed from the magnet and placed in a 15 mL tube.
Resuspend the cells at 1 × 106 cells per milliliter in X-VIVO15 + 2% human serum and keep them at 4 °C.
Note: Cells can be cultured for up to 48 h with high percentage of viable cells.
Purity check
Note: Flow cytometry analysis can be performed the day after the isolation of the three DC subsets. Until the analysis is performed, cells should be stored at 4 °C.
Fill the wells of a 96-well V-bottomed plate with at least 10,000 cells/well (also considering the unstained control and isotype).
Note: Three wells need to be filled per subset (unstained, isotype mixture, and the antibody mixture).
Wash 1× with 100 μL of PBA.
Prepare three different antibody (Ab) combinations (every Ab mixture identifies one of the DC subsets) with the antibodies indicated in Table 1. The dilutions of the antibodies are made with PBA.
Table 1. Antibody mixtures to measure purity of cDC2, pDC, and cDC1 populations
Population Target Label Dilution
cDC2 purity CD20 FITC 1/25
CD1c PE 1/50
CD14 PercP 1/25
CD11c APC 1/50
pDC purity BDCA2 PE 1/25
CD14 PercP 1/25
CD123 APC 1/25
CD20 FITC 1/25
cDC1 purity CD1c PE 1/50
CD14 PercP 1/25
CD141 APC 1/100
Add 25 μL of antibody mixture to the appropriate wells.
Incubate for 20 min at 4 °C in the dark.
Wash twice with 100 μL of PBA.
Resuspend in 100 μL of PBA.
Measure using a flow cytometer.
Data analysis
In the following section, examples of gating strategies for the purity assessment of the isolated DC subsets are shown after the samples were analyzed by flow cytometry.
Note: Flow cytometry data were obtained in .fcs files and uploaded to FlowJo.
For the three DC subsets, identify viable cells on FSC vs. SSC gating (Figure 4A). To evaluate the purity of the viable cDC1s, gate the cells that are negative for CD14 and CD1c (Figure 4B) and gate on CD141, the marker for cDC1s (Figure 4C).
Figure 4. Purity analysis of the cDC1 subset
To evaluate the purity of the viable cDC2s (Figure 5A), gate the cells that are negative for CD14 and CD20 (Figure 5B), and gate for CD11c and CD1c double-positive cells, markers for cDC2s (Figure 5C).
Figure 5. Purity analysis of the cDC2 subset
To evaluate the purity of the viable pDCs (Figure 6A), gate the cells that are negative for CD14 (SSC vs. CD14 plot, Figure 6B), and gate for CD123 and BDCA2 double-positive cells, markers for pDCs (Figure 6C).
Figure 6. Purity analysis of the pDC subset
Notes
In Table 2 we provide data regarding the reproducibility and variability of the DC isolation protocol. Data come from the percentage of pure cells obtained after analysis by flow cytometry. This data corresponds to three independently performed experiments (three different donors).
Table 2. Purity of DC subsets (n = 3)
Donor/independent experiment Purity of cDC1s (%)
Purity of cDC2s (%)
Purity of pDCs (%)
Donor A 87.5 94.5 98.3
Donor B 91.3 99.1 98.5
Donor C 99.2 93.2 99
Recipes
Washing buffer
2 mM EDTA and 1% bovine serum albumin (BSA) in PBS.
ACK lysis buffer
Use Milli-Q water and dissolve the following salts:
0.15 M ammonium chloride (mw: 53.49 g/mol)
0.01 M potassium bicarbonate (mw: 100.12 g/mol)
0.0001 M disodium EDTA (mw: 372.24 g/mol)
PBA buffer
Prepare by adding the reagents below to PBS. The storage of PBA is at 4 °C.
1% BSA
0.05% sodium azide (needs to be dissolved in Milli-Q water)
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Le Gall et al. (2022). Efficient targeting of NY-ESO-1 tumor antigen to human cDC1s by lymphotactin results in cross-presentation and antigen-specific T cell expansion. J. Immunother Cancer (Figure 3, panel A, B, C).
General notes and troubleshooting
Troubleshooting
This protocol has limitations. To isolate human DC subsets following this protocol, there is a need for high cell numbers (PBMCs) in the starting material. Aphaeresis material instead of a buffy coat is therefore recommended. Another limitation are the high costs for the MultiMACS separator. This could be replaced by the use of single columns and magnets. However, the isolation time will increase considerably.
Acknowledgments
This work was supported by Health Holland grant DC4Balance (LSHM18056-SGF).
Competing interests
Miltenyi Biotec GmbH was as part of the public-private DC4Balance consortium. All reagents purchased to perform these experiments and the MultiMACS were bought by Radboud University Medical Center.
Ethical considerations
Peripheral blood mononuclear cells (PBMCs) were isolated from healthy donor blood (Sanquin, the Netherlands) through density centrifugation using Lymphoprep (Axis-Shield).
References
van Beek, J. J., Flórez-Grau, G., Gorris, M. A., Mathan, T. S., Schreibelt, G., Bol, K. F., Textor, J. and de Vries, I. J. M. (2020). Human pDCs Are Superior to cDC2s in Attracting Cytolytic Lymphocytes in Melanoma Patients Receiving DC Vaccination. Cell Rep. 30(4): 1027–1038.e4.
Collin, M. and Bigley, V. (2018). Human dendritic cell subsets: an update. Immunology 154(1): 3–20.
Corkum, C. P., Ings, D. P., Burgess, C., Karwowska, S., Kroll, W. and Michalak, T. I. (2015). Immune cell subsets and their gene expression profiles from human PBMC isolated by Vacutainer Cell Preparation Tube (CPT™) and standard density gradient. BMC Immunology 16(1): e1186/s12865-015-0113-0.
Gonzalez, H., Hagerling, C. and Werb, Z. (2018). Roles of the immune system in cancer: from tumor initiation to metastatic progression. Genes Dev. 32: 1267–1284.
Schreibelt, G., Klinkenberg, L. J. J., Cruz, L. J., Tacken, P. J., Tel, J., Kreutz, M., Adema, G. J., Brown, G. D., Figdor, C. G., de Vries, I. J. M., et al. (2012). The C-type lectin receptor CLEC9A mediates antigen uptake and (cross-)presentation by human blood BDCA3+ myeloid dendritic cells. Blood 119(10): 2284–2292.
Wang, Y., Xiang, Y., Xin, V. W., Wang, X. W., Peng, X. C., Liu, X. Q., Wang, D., Li, N., Cheng, J. T., Lyv, Y. N., et al. (2020). Dendritic cell biology and its role in tumor immunotherapy. J. Hematol. Oncol. 13(1): e1186/s13045-020-00939-6.
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Immunology > Immune cell isolation > Antigen-presenting cell
Cell Biology > Cell isolation and culture > Cell isolation
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It would be nice to also mention at least roughly how many cells can be optained for example per 10^6 PBMCs. Thank you
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Oct 25, 2023
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Three-dimensional Co-culture Model for Live Imaging of Pancreatic Islets, Immune Cells, and Neurons in Agarose Gel
EM Elke M. Muntjewerff §
VJ Vijay S. Josyula
GC Gustaf Christoffersson
(§ Technical contact)
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4852 Views: 1114
Reviewed by: Mohan BabuAndrea GramaticaShun Yu Jasemine Yang
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Abstract
During the onset of autoimmune diabetes, nerve–immune cell interactions seem to play an important role; however, there are currently no models to follow and interfere with these interactions over time in vivo or in vitro. Two-dimensional in vitro models provide insufficient information and microfluidics or organs on a chip are usually challenging to work with. We present here what we believe to be the first simple model that provides the opportunity to co-culture pancreatic islets with sympathetic nerves and immune cells. This model is based on our stamping device that can be 3D printed (STL file provided). Due to the imprint in the agarose gel, sympathetic neurons, pancreatic islets, and macrophages can be seeded in specific locations at a level that allows for confocal live-cell imaging. In this protocol, we provide the instructions to construct and perform live cell imaging experiments in our co-culture model, including: 1) design for the stamping device to make the imprint in the gel, 2) isolation of sympathetic neurons, pancreatic islets, and macrophages, 3) co-culture conditions, 4) how this can be used for live cell imaging, and 5) possibilities for wider use of the model. In summary, we developed an easy-to-use co-culture model that allows manipulation and imaging of interactions between sympathetic nerves, pancreatic islets, and macrophages. This new co-culture model is useful to study nerve– immune cell– islet interactions and will help to identify the functional relevance of neuro-immune interactions in the pancreas.
Key features
• A novel device that allows for 3D co-culture of sympathetic neurons, pancreatic islets, and immune cells
• The device allows the capture of live interactions between mouse sympatheticneurons, pancreatic islets, and immune cells in a controlled environment after six days of co-culturing.
• This protocol uses cultured sympathetic neurons isolated from the superior cervical ganglia using a previously established method (Jackson and Tourtellotte, 2014) in a 3D co-culture.
• This method requires 3D printing of our own designed gel-stamping device (STL print file provided on SciLifeLab FigShare DOI: 10.17044/scilifelab.24073062).
Graphical overview
Graphical overview of co-culture model. 1) Print the stamp with a 3D printer. 2) Isolate neurons, islets, and macrophages. 3) Use the stamp to make the imprint in the agarose gel. 4) Seed the macrophages and islets in the agarose gel on their seeding points. 5) Place the coverslip with neurons on top. 6) Incubate the culture for six days. 7) Image the co-culture. Images adapted from BioRender.
Keywords: 3D co-culture Nerve-immune interactions Sympathetic nerves Macrophages Pancreatic islets Pancreatic innervation Immune cell migration Diabetes
Background
It is becoming increasingly clear that the sympathetic nervous system and its interactions with immune cells play an important role in maintaining homeostasis(Martinez-Sanchez et al., 2022). In line with this, type 1 diabetes (T1D) onset in mice halts after depletion of macrophages or sympathetic nerves, via surgical denervation or chemical ablation of nerves (Christoffersson et al., 2020). This shows that sympathetic nerves and macrophages play important roles in T1D onset. Moreover, less severe autoimmune diabetes onset was observed in non-obese diabetic mice (NOD) when the pancreatic nerve was stimulated (Guyot et al., 2019). This suggests that nerve signal modulation might be beneficial for T1D patients in the future and needs to be investigated further. However, currently there are no models to follow and interfere in pancreatic nerve–immune cell interactions over time in vivo.
Injection of neuromodulating agents such as neuropeptides, neurotransmitters, and receptor antagonists in mice has systemic effects, limiting isolation of microenvironmental changes. Separate culture and stimulation of cells in 2D cultures will, on the other hand, provide insufficient information. Microfluidics or organs on a chip are solutions used to measure hormone secretion and more complex cellular behavior in co-culture conditions, but the setup for these microfluidic models is usually challenging. Therefore, we developed a simple 3D co-culture model that allows imaging of interactions between sympathetic nerves, pancreatic islets, and macrophages.
In this protocol, we provide the instructions to construct and perform live cell imaging experiments in our co-culture model. We describe: 1) the design for the stamping device to make the imprint in the gel, 2) the isolation of sympathetic neurons, pancreatic islets, and macrophages, 3) co-culture conditions, and 4) how this can be used for live cell imaging. Finally, we discuss possibilities for a wider use of the model and suggestions for processing of the gel containing islets and cells after imaging for further data collection.
Using this new model, it is possible to study and interfere in neuro-immune interactions in the context of the pancreatic islet in a controlled environment. This environment can be manipulated by addition of neurotransmitters, peptides, or drugs. Instead of macrophages, it would be possible to seed other immune cells, such as T cells, dendritic cells, or monocytes. Due to the three seeding points for cells, it is also possible to seed different immune cells at the same time for example inflammatory and anti-inflammatory macrophages or differently activated Tcells. The versatility of this model makes it useful for studying pancreatic islet physiology and immune attack. In the future, it could also be adapted for the use of pancreatic islet organoids in combination with human immune cells. Altogether, this new co-culture model is useful to study nerve–immune cell–isletinteractions and will help to identify the functional relevance of neuro-immune interactions in the pancreas.
Materials and reagents
Biological materials
Two-day-old pups from a DsRed mouse litter [JAX strain 005441, Tg(CAG-DsRed*MST)1Nagy/J]
Mice expressing GFP [JAX strain 011106, Tg(CAG-GFP*)1Hadj/J]
Wildtype mice (Jackson Laboratory, C57BL/6 mice)
Reagents
Collagenase A, Type 4 (Roche, catalog number: 11088793001), use 10 mg/mL in media
12 mm poly-D-lysine(PLL)/laminin coated German Glass coverslip (Corning BioCoat, catalog number: 354087)
Nerve growth factor (NGF) (Peprotech, catalog number: 450-01)
Trypsin 0.25% EDTA, Mg2+ Ca2+-free (Corning Cellgro®, catalog number: 25-053-Cl)
Fetal bovine serum (FBS) (Sigma, catalog number: F7524-500ML)
Neurobasal media (Gibco, catalog number: 21103049)
B27 supplement (Gibco, catalog number: 17504044)
Penicillin-streptomycin (Pen/strep) 10,000 U/mL (Gibco, catalog number: 15140122)
UltraPure agarose (Invitrogen, catalog number: 1650010)
Low-gelling temperature agarose (Sigma, catalog number: A9045-5G)
RPMI 1640 medium (Thermo Fisher, catalog number: 21875-091)
D-(+)-Glucose (VWR, AnalaR NORMAPUR, catalog number: 101174Y)
Celltracker violet (Invitrogen, catalog number: C10094)
Macrophage colony stimulating factor (M-CSF) (Peprotech, catalog number: 3512-02)
L-glutamine (Life Technologies, catalog number: 25030-024)
Milli-Q (tapped from Purelab Ultra, model: USC214628)
Solutions
Neuron medium (see Recipes)
Islet medium (see Recipes)
Co-culture medium (see Recipes)
Agarose gel (see Recipes)
Low-gelling agarose gel (see Recipes)
Recipes
Neuron medium
Reagent Final concentration Quantity
Neurobasal media - 492.8 mL
B27 supplement 2% 200 μL
Pen/strep 0.5% 2.5 mL
L-glutamine 0.5 mM 5 mL
Islet medium
Reagent Final concentration Quantity
RPMI1640 435 mL
FBS 10% 50 mL
Pen/Strep 5% 5 mL
L-Glutamine 0.5 mM 5 mL
D-(+)-Glucose 11.1 mM [1 g in 5 mL] 5 mL
Co-culture medium
Reagent Final concentration Quantity
Islet medium (from recipe 2) - 50 mL
M-CSF 50 ng/mL 50 μL
NGF 10 ng/mL 50 μL
*M-CSF and NGF are not stable; keep frozen as aliquots (add shortly before use)
Agarose gel
Reagent Final concentration Quantity
UltraPure agarose 1.2% 0.48 g
Autoclaved Milli-Q
RPMI1640 medium
-
-
10 mL
30 mL
Low-gelling agarose gel
Reagent Final concentration Quantity
Low-gelling temperature agarose 1.2% 0.48 g
Autoclaved Milli-Q
RPMI 1640 medium
-
-
10 mL
30 mL
Laboratory supplies
6 cm tissue culture dishes (VWR, Avantor, catalog number: 10861-588)
10 cm tissue culture dishes (Thermo Scientific, catalog number: 150464)
96-well U-bottom tissue culture plate (VWR, catalog number: 734-2328)
25 cm cell scraper (VWR, catalog number: 734-2602)
50 mL tubes (Falcon, catalog number: 352070)
50 mL Erlenmeyer (VWR, catalog number: 214-1130)
Parafilm M (VWR, catalog number: 291-1214)
Water bath (Grant, catalog number: TC120)
Equipment
Surgical stereo microscope (Leica, model: MZ10F)
Brightfield microscope (VWR, model: 020025)
Fluorescent microscope (Bio-Rad, ZOE Fluorescent Imager, model: 1450031)
Confocal microscope (Leica, model: TCS SP8, objective model: HC Fluotar L 25×/0.95 W)
FormTM Steri-CylceTM CO2 incubator (Thermo Fisher, model: 371)
Laminar flow hood (Kojair, model: KBS-125)
Microwave (Whirlpool, model VIP 20)
Heating device for imaging (LCI custom heating plate for 6 cm dishes)
3D printer (Asiga, model: MAX X 27)
Cell counting chamber (Neubauer, model: HIRS8100204)
Benchtop centrifuge (Beckman Coulter, model: Allegra X30R)
Software and datasets
ImageJ/Fiji for analyzing obtained imaging data, version number: 1.8.0 (free software)
Microscopy image analysis software, IMARIS, version number: 9.9 (requires a license)
Procedure
This protocol consists of three main parts: 1) the design of the stamp to create the imprint in the gel, 2) the protocol for culture in the device including cell isolation, gel preparation, building the agarose co-culture, and imaging, and 3) suggestions for interaction studies after imaging.
The design of the stamp to create the imprint in the agarose gel
Before the start of this protocol, make sure to 3D print the stamp needed to make the imprint in the agarose gel (Figure 1). The stamping device was 3D-printed in the material PlasGray (Asiga) with an Asiga Max X 27 3D printer (Asiga) with a layer height of 50 μM. The stamp includes holes to prevent air bubble formation during stamping in the agarose gel. The STL file is provided in the supplementary materials and on the SciLifeLab FigShare website DOI: 10.17044/scilifelab.24073062.
Figure 1. Stamp design. (A) Front view of stamp and measurements: radius (26.5 mm) and width (50 mm). (B) Part of the stamp that creates the imprint in the gel. Measurements for these parts: seeding point width (0.5 mm), capillary width (0.3 mm), islet seeding point width (2 mm), distance from islet seeding point until capillary circle (10 mm). (C) Side view from the device including measurements: height of the part that stays on the dish during stamping (3 mm), height of the stamp that fits in the dish (10 mm), and the depth of the stamp (0.5 mm). (D) Tilted view of the stamp.
Protocol for the co-culture including cell isolation, gel preparation, building the co-culture, and imaging
Isolation and culturing of sympathetic neurons
Isolate superior cervical ganglia from DsRed pups (maximum three days old) as described in Jackson’s protocol “Neuron Culture from Mouse Superior Cervical Ganglion” (Jackson and Tourtellotte, 2014), followed by neuron isolation from the ganglia as described below.
Note: Steps A2–A6 are adapted from Jackson and Tourtellotte (2014) .
Place superior cervical ganglia into a 50 mL tube with ~200 μL collagenase A (10 mg/mL) for a single ganglia (use 400 μL for four ganglia) for 30 min in a 37 °C water bath and agitate every 10 min.
Remove the collagenase solution by carefully pipetting around the ganglia. Replace with approximately 200 μL of 0.25% trypsin for 1–2 ganglia (use 500 μL for four ganglia). Place in 37 °C water bath for 15 min and agitate approximately every 5 min.
Note: The 50 mL tube can be swirled to bring ganglia together in the middle of the tube.
Add 1 mL of neurobasal media to inactivate trypsin and remove both media and trypsin by aspirating media around the ganglia.
Rinse ganglia three times in 1 mL of neuron medium (see Recipes).
Re-suspend ganglia in 200 μL of neuron medium and pipette up and down with barrier tips of 200 μL until the ganglia are no longer visible (~60–70 times).
Use a cell counter (counting chamber) to determine cell number. One ganglion yields approximately 35–40,000 cells.
Dilute neurons to 6,000 neurons/100 μL in neuron medium. Seed 3,000 neurons in droplets of 50 μL per 12 mm PLL/laminin-coated glass coverslip and place in incubator (37 °C, 5% CO2).
After one hour, check with a brightfield microscope if the cells are attached to the coverslip and carefully add an additional 450 μL of pre-warmed neuron medium supplemented with NGF (10 ng/mL) using the side wall of the plate (to not disturb the cells).
Note: In case the neurons are not attached after 2 h, the cells are probably not viable and should not be used.
Culture neurons in incubator (37 °C, 5% CO2) with medium changes every two days. To not disturb the cells, half of the neuron medium can be changed each time.
Note: Since only half of the medium is changed, the added NGF should be doubled (add 250 μL with 20 ng/mL NGF per well to reach a concentration of 10 ng/mL in the full 500 μL. For medium change, tilt the plate at an angle towards you to collect the medium in the bottom and add new medium along the wall of the plate to limit disturbance of the cells.
Neurons are ready to be used for co-culture after four days and can be kept in culture for 28 days (Figure 2A).
Make and stamp agarose gel
Warm up 40 mL of islet medium to 37 °C (see Recipes).
Weigh 0.48 g of agarose in a small Erlenmeyer flask (see also agarose gel recipe).
Note: We recommend using normal agarose to prevent the gel from melting during imaging.
Add 10 mL of autoclaved Milli-Q water to the agarose in a laminar flow hood.
Cover the Erlenmeyer flask with parafilm and heat it up in the microwave until agarose is completely dissolved.
Back in the hood, add 30 mL of pre-warmed 37 °C medium. Mix medium and dissolved agarose together by swirling and pipetting.
Pipet 4 mL of agarose-medium mix into a 6 cm dish, resulting in an approximately 5 mm thick layer.
Note: The pipette tip can be placed against the wall to prevent air bubble formation.
Wait until the gel starts to set (approximately 5–10 min).
Place the stamp on the set gel in the dish and apply light pressure for 10 s to make an imprint in the gel. Remove any remaining gel from the stamp with an ethanol-drenched tissue and quickly continue to the next one (Figure 2B).
Figure 2. Co-culture setup. (A) Ds-Red neurons growing on coverslip. (B) Picture of the printed stamp. (C) Agarose gel with imprint providing seeding points for the pancreatic islets (white arrow) and the immune cells (indicated with three black arrows). Circled area is enlarged in Figure 2D. (D) Macrophages spread out from the seeding point into the half circled imprinted area. (E) Picture of coverslip and gel droplet placing to secure the location of the coverslip. (F) Picture of an islet in the co-culture model including brightfield to visualize all cells, GFP to visualize the islet, and Ds-Red to visualize the neurons. (G) Imaging setup of the co-culture. (H) Picture of the live imaging using the 488 laser to visualize the islet (indicated with white dotted line) and 512 to visualize the Ds-Red neurons.
Let the gel set for an additional 10 min at room temperature without the lid to fully solidify (Figure 2C).
Continue to use the gel for the co-culture or put the lid back on and wrap the plate with parafilm before storage at 4 °C.
Collect pancreatic islets from a GFP mouse
Islets were isolated from a global GFP-expressing mouse as described elsewhere (Bohman et al., 2006).
After isolating using the gradient, hand pick the islets using a stereomicroscope.
Collect eight islets per well in a 96-well U-bottom plate in islet medium and keep in the incubator until ready for seeding (section E).
Isolate macrophages
For macrophages in the co-culture: isolate and grow bone marrow–derived macrophages as described previously in Haag and Murthy (2021).
Collect macrophages from a 10 cm dish using a cell scraper.
If not using macrophages expressing fluorescent protein: stain the macrophages with violet or far red dye cell tracer according to manufacturer’s instructions.
Count macrophages and dilute to 100,000 cells/μL in co-culture medium.
Seed GFP-islets and macrophages in the gel
Place the 6 cm dish containing the imprinted agarose gel (from section A) in a laminar flow hood.
Spin down the plate with islets (200 g, 2 min) and remove the medium in the hood. Resuspend islets in 10 μL of islet medium per well.
Let the islets sink to the lower part of the pipette tip and inject 5 μL of medium with islets into the middle circle of the gel.
Check if the placement was successful using a brightfield microscope.
Proceed with seeding 1 μL of macrophages per seeding point (three in total).
Note: Turn off lights in the hood and use a small light from the side for better visibility of the seeding points.
Check if macrophages are in the seeding points and have spread into the crescents by the capillary effect (Figure 2D).
Note: In case the macrophages do not spread into the crescents, it will not be possible to identify the exact starting location. However, it will still be possible to image interactions between neurons, islets, and macrophages.
Add 200 μL of liquid low melting 1.2% agarose gel on top to seal islets and macrophages in (see Recipes or section B).
Quickly add coverslip containing neurons on top of the new gel, so that it is located above the islets.
Put droplets of low-melting agarose gel on the sides of the glass slide to make sure the coverslip will stay in place (Figure 2E).
Check location of cells and islets in the gel using a brightfield and/or fluorescence microscope.
Add 4 mL of co-culture medium (see Recipes) and put the dish in the incubator.
Change co-culture medium every two days. For medium changes, the complete medium can be pipetted off and replaced with new co-culture medium.
At this stage, stimulants/inhibitors can be added to the medium at any time.
Cells in co-culture can be checked using brightfield or fluorescence microscopes (Figure 2F).
Start imaging after day six to provide the neurons with enough time to reach the islet (Figure 2F).
Imaging of co-culture with neurons, pancreatic islets, and macrophages
Check and mark location of islets and interesting spots, such as macrophages or islets.
Start up the microscope and heating device.
Place the co-culture in the heating device under the microscope (Figure 2G).
Use brightfield or GFP signal to find the islets.
Set the microscope to capture immune cells (violet), pancreatic islets (GFP), and neurons (DsRed).
Set Z-stack capturing the full islet. Immune cells and neurons might be slightly below or above the islet, so check their location and include them in the Z stack if wanted.
Start imaging. For our experiments, we imaged every 2 min during 30 min per islet with a Z slice size of 1–2 μm (depending on the size of the islet) (Figure 2H).
Note: Keep an eye on the fluid levels in the dish during imaging and add more medium if necessary to keep the connection between the objective and the fluid in the dish.
If using sterile imaging in a temperature-controlled chamber, the co-culture could be followed over time by keeping it in a tissue culture incubator and re-image on the required timepoints.
Suggestions for interaction studies after imaging
To study the interactions in the islet area further after imaging, the islets could be cut out of the gel using a stereomicroscope and scalpel. The gel containing islets can be fixed in 4% PFA overnight at 4 °C and used for additional fluorescent staining. Alternatively, it would even be possible to remove the agarose and use the isolated cells for flow cytometry, qPCR, or single-cell RNA sequencing.
Data analysis
For migration analysis, we would recommend making an overview (e.g., tile-scan) image to count all the macrophages that reach the middle. Additionally, the distance from the seeding point to the middle circle is known, making it possible to calculate the migrated distance of immune cells to the islets. If a picture is acquired every day from the start of the co-culture using a fluorescence microscope, the macrophage speed under various conditions can be followed. We would recommend tracking at least 30 macrophages per gel, but this number should be optimized by the end user depending on the goal of the experiment and variation in immune cell behavior.
Images and movies made with the microscope can be further analyzed using IMARIS or ImageJ. Using this software, it is possible to determine cell numbers and cell–cell interactions and follow cell tracks of macrophages, GFP+ cells that migrate out of the pancreatic islet, and DsRed nerves.
Acknowledgments
3D printing was performed at U-PRINT: Uppsala University’s 3D-printing facility at the Disciplinary Domain of Medicine and Pharmacy. We thank Adam Engberg for his advice for the design and printing of the stamping device. Funding: E.M. is funded by a Rubicon grant from the Netherlands Organization for Scientific Research (NWO). G.C. is supported by grants from the Swedish Research Council, the Swedish Society for Medical Research and the Göran Gustafsson foundation, and the Science for Life Laboratory (SciLifeLab). Kind support for the development of this model was also provided by the Family Ernfors Fund.
Competing interests
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Ethical considerations
All mouse procedures were approved by the local animal ethics committee for the Uppsala region.
References
Bohman, S., Andersson, A. and King, A. (2006). No Differences in Efficacy Between Noncultured and Cultured Islets in Reducing Hyperglycemia in a Nonvascularized Islet Graft Model. Diabetes Technol. Ther. 8(5): 536–545.
Christoffersson, G., Ratliff, S. S. and von Herrath, M. G. (2020). Interference with pancreatic sympathetic signaling halts the onset of diabetes in mice. Sci. Adv. 6(35): eabb2878.
Guyot, M., Simon, T., Ceppo, F., Panzolini, C., Guyon, A., Lavergne, J., Murris, E., Daoudlarian, D., Brusini, R., Zarif, H., et al. (2019). Pancreatic nerve electrostimulation inhibits recent-onset autoimmune diabetes. Nat. Biotechnol. 37(12): 1446–1451.
Haag, S. and Murthy, A. (2021). Murine Monocyte and Macrophage Culture. Bio Protoc 11(6): e3928.
Jackson, M. and Tourtellotte, W. (2014). Neuron Culture from Mouse Superior Cervical Ganglion. Bio Protoc 4(2): e1035.
Martinez-Sanchez, N., Sweeney, O., Sidarta-Oliveira, D., Caron, A., Stanley, S. A. and Domingos, A. I. (2022). The sympathetic nervous system in the 21st century: Neuroimmune interactions in metabolic homeostasis and obesity. Neuron 110(21): 3597–3626.
Supplementary information
The following supporting information can be downloaded here:
STL file
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Cell Biology > Cell isolation and culture > 3D cell culture
Neuroscience > Cellular mechanisms > Cell isolation and culture
Immunology > Immune cell imaging > Confocal microscopy
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Improve Research Reproducibility
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Peer-reviewed
Efficient Large DNA Fragment Knock-in by Long dsDNA with 3′-Overhangs Mediated CRISPR Knock-in (LOCK) in Mammalian Cells
WH Wenjie Han
HL Haojun Liang
JB Jianqiang Bao
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4853 Views: 1162
Reviewed by: Xin Xu 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 May 2023
Abstract
An efficient and precise genome-editing approach is in high demand in any molecular biology or cell biology laboratory worldwide. However, despite a recent rapid progress in the toolbox tailored for precise genome-editing, including the base editors and prime editors, there is still a need for a cost-effective knock-in (KI) approach amenable for long donor DNA cargos with high efficiency. By harnessing the high-efficient double-strand break (DSB) repair pathway of microhomology-mediated end joining, we previously showed that a specially designed 3′-overhang double-strand DNA (odsDNA) donor harboring 50-nt homology arm (HA) allows high-efficient exogenous DNA KI when combined with CRISPR-Cas9 technology. The lengths of the 3′-overhangs of odsDNA donors could be manipulated by the five consecutive phosphorothioate (PT) modifications. In this protocol, we detail the stepwise procedures to conduct the LOCK (Long dsDNA with 3′-Overhangs mediated CRISPR Knock-in) method for gene-sized (~1–3 kb) KI in mammalian cells.
Graphical overview
Improvement of large DNA fragment knock-in rates by attaching odsDNA donors to Cas9-PCV2 fusion protein
Keywords: Genome editing CRISPR/Cas9 Gene knock-in (KI) Homologous recombination ssDNA donor dsDNA donor odsDNA donor Cas9-PCV2 fusion
Background
The advent of CRISPR and its derived technologies have tremendously expanded the toolbox for genome manipulation for scientists in the fields of biomedical research and innovative biotechnological development. Compared with the ZFN or TALEN approaches, which are highly reliant on the specific recognition of the target genomic DNA (gDNA) sequence by specially engineered protein readers, the CRISPR-Cas9 method only necessitates a single crRNA (~20 nt) that uniquely recognizes and pairs with the specific gDNA locus (Hsu et al., 2014). This improvement makes genome-editing technique adoptable in any individual lab because of the low cost and simplified procedures. However, it is known that the precise integration of large donor DNA sequences into the host genome is dependent on homology-directed repair (HDR), which occurs at low frequencies, in mammalian cells. In addition, the double-strand break (DSB) sites induced by Cas9 cutting could happen at undesired genomic loci (off-target), and this might result in unexpected DSB repair outcomes, e.g., insertions/deletions (indels) and translocations, which are fundamentally detrimental to cellular health. It is, therefore, critical to increase the precise targeting efficiency while minimizing the off-target effects (Doudna, 2020).
In the past decade, the scientists have made great achievements in the improvement of the precise CRISPR-Cas9 knock-in (KI) efficiency. The phosphoric backbone of the donor DNA sequences is highly negatively charged, which naturally impedes their transportation across the cellular membrane, leading to the inaccessibility of exogenous DNA donors in the local DSB sites. This has been attributed as one of the reasons causing the low efficiencies of HDR-mediated KI. As such, versatile approaches have by far been devised to promote the nuclear delivery or the structural stability of the exogenous repair DNA donors (Yu et al., 2020). For instance, the nuclear entry of DNA donors could be promoted by adeno-associated viral (AAV) vectors, by Strep-biotin labeled tethering (Gu et al., 2018), or by chromatinized packaging (Cruz-Becerra and Kadonaga, 2020).
On the other hand, it is known that while the canonical nonhomologous end joining (c-NHEJ) repair is much more frequent at the CRISPR-Cas9-induced DSB sites, the repair pathway could be biased to the precise HDR repair outcome through artificially manipulating the cellular milieu (Maruyama et al., 2015). Indeed, accumulating examples have shown that the long ssDNA (lssDNA), as exogenous HDR donors, display superior advantages over the conventional double-strand DNA (dsDNA) donors, in terms of the HDR efficiency and off-target effect (Quadros et al., 2017; Shy et al., 2023). By leveraging the distinct advantages from both dsDNA and ssDNA donors, our previous study showed that a single hybrid “3′-overhang dsDNA” donor, termed odsDNA, has proved to significantly improve the HDR efficiencies by up to 5-fold increase with low off-target effect (Han et al., 2023). We named this Long dsDNA with 3′-Overhangs mediated CRISPR Knock-in, “LOCK” for short. In this protocol, we describe this easy technique with the stepwise procedures needed to be operational in any individual lab.
Materials and reagents
Materials
Pipette tips: 10, 200, and 1,000 μL (Yeasen, catalog numbers: 83010ES20, 83040ES20, and 83070ES08)
200 μL extended pipette tips, sterilized (Yeasen, catalog number: 83061ES50)
Pipettes: 2.5, 10, 20, 100, 200, and 1,000 μL (Eppendorf, catalog numbers: 3120000216, 3120000224, 3120000232, 3120000240, 3120000259, and 3120000267)
0.2 mL PCR strips of eight tubes (Yeasen, catalog number: 83602ES10)
1.5 mL microcentrifuge tube (Axygen, catalog number: MCT-150-C)
50 mL conical tube (Thermo Fisher, catalog number: 339652)
DNA oligonucleotide primers and PCV2 linker (Sangon Biotech)
6-well cell culture plate (BIOFIL, catalog number: TCP011006)
60 mm cell culture dish (BIOFIL, catalog number: TCD010060)
10 cm cell culture dish (BIOFIL, catalog number: TCD010100)
HEK293T cells (American Type Culture Collection, catalog number: CRL-3216)
DH5α competent cells (Sangon Biotech, catalog number: B528413)
Escherichia coli Rosetta 2 (DE3) strain (Millipore EMD, catalog number: 71397)
Isopropyl β-D-1-thiogalactopyranoside (IPTG) (Invitrogen, catalog number: 15529019)
Tryptone (Gibco, catalog number: 211705)
Yeast extract (Gibco, catalog number: 211931)
Kanamycin monosulfate (Sangon Biotech, catalog number: A600286)
Agar (Thermo Fisher Scientific, catalog number: 22700025)
Terrific broth (Thermo Fisher Scientific, catalog number: 22711022)
Glycine (Invitrogen, catalog number: 15527013)
SDS (Sigma-Aldrich, catalog number: 436143)
NaCl (Sigma-Aldrich, catalog number: S9888)
KCl (Sigma-Aldrich, catalog number: P3911)
MgCl2 hexahydrate (Sigma-Aldrich, catalog number: M2670)
Tris (Invitrogen, catalog number: A32355)
HCl (Sigma-Aldrich, catalog number: 30721)
NaOH (Sigma-Aldrich, catalog number: 221465)
Imidazole (Sigma-Aldrich, catalog number: I2399)
UltraPure glycine (Invitrogen, catalog number: 15527013)
β-mercaptoethanol (Gibco, catalog number: 21985023)
EDTA (Thermo Fisher Scientific, catalog number: 17892)
HEPES (Sigma-Aldrich, catalog number: 54457)
TCEP (Thermo Fisher Scientific, catalog number: T2556)
Glycerol (Thermo Fisher Scientific, catalog number: 17904)
HisTrap fast flow, 5 mL (GE Healthcare, catalog number: GE17-5255-01)
Amicon Ultracel-100 regenerated cellulose membrane, 50 mL (Millipore, catalog number: UFC910008)
Amicon Ultracel-30 regenerated cellulose membrane, 4 mL (Millipore, catalog number: UFC803008)
0.22 μm syringe filter, PES membrane, 33 mm diameter (Millipore, catalog number: SLGPR33RS)
0.45 μm syringe filter, PES membrane, 33 mm diameter (Millipore, catalog number: SLHPR33RS)
Nalgene Rapid-Flow with PES 500 mL (Thermo Fisher Scientific, catalog number: 566-0020)
PD-10 desalting columns (GE Healthcare, catalog number: GE17-0851-01)
BeyoGel SDS-PAGE Precast Gel, Tris-Gly, 4%–20%, 12 wells (Beyotime, catalog number: P0057A)
Agarose (Thermo Fisher Scientific, catalog number: R0491)
GeneArt Precision gRNA Synthesis kit (Invitrogen, catalog number: A29377)
Invitrogen TrueCut Cas9 Protein v2 (Thermo Fisher Scientific, catalog number: A36497)
PARAFILM sealing film (Sigma-Aldrich, catalog number: HS234526B-1EA)
Kimwipes disposable wipers (Sigma-Aldrich, catalog number: Z188956)
Reagents
ddH2O, DEPC-treated water (Sangon Biotech, catalog number: B300592)
Lambda exonuclease (New England Biolabs, catalog number: M0262L)
GeneJET PCR Purification kit (Thermo Fisher Scientific, catalog number: K0702)
SpCas9 expression plasmid (Addgene, catalog number: 47327)
Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, catalog number: C11885500BT)
TrypLE express (Gibco, catalog number: 12604039)
DPBS (without Ca2+ or Mg2+) (Gibco, catalog number: 14190-144)
Fetal bovine serum (FBS) (Gibco, catalog number: 10091148)
Cell Line Nucleofector kit V (Lonza, catalog number: VCA-1003)
Q5 High-Fidelity 2× Master Mix (New England Biolabs, catalog number: M0492L)
50× TAE buffer (Sangon Biotech, catalog number: B548101)
6× Gel Loading Dye Purple (New England Biolabs, catalog number: B7024S)
SDS-PAGE Protein Loading Buffer, 5× (Beyotime, catalog number: P0286)
BeyoColor Prestained Color Protein Marker, 10–170 kDa (Beyotime, catalog number: P0077)
Imperial protein stain solution (Thermo Fisher Scientific, catalog number: 24615)
Annealing buffer for DNA Oligos, 5× (Beyotime, catalog number: D0251)
LB medium (see Recipes)
50 mg/mL kanamycin (see Recipes)
LB agar plate with kanamycin (see Recipes)
Terrific broth (TB) medium (see Recipes)
1 M IPTG (see Recipes)
Buffer A: start buffer (see Recipes)
Buffer B: elution buffer (see Recipes)
Buffer C: storage buffer (see Recipes)
Buffer D: cleavage buffer (see Recipes)
0.1 M NiSO4 (see Recipes)
0.1 M EDTA, 1 M NaCl (see Recipes)
1 M NaOH (see Recipes)
1 M HCl (see Recipes)
SDS-PAGE running buffer (see Recipes)
Low-EDTA TE buffer (see Recipes)
Recipes
LB medium (1,000 mL)
Reagent Final concentration Quantity
Tryptone
Yeast extract
NaCl
10 g/L
5 g/L
10 g/L
10 g
5 g
10 g
ddH2O n/a ~980 mL
Total n/a 1,000 mL
Adjust the pH to 7.4 with NaOH. Autoclave at 121 °C for 15 min. Store at RT.
50 mg/mL kanamycin (10 mL)
Reagent Final concentration Quantity
Kanamycin 50 mg/mL 0.5 g
ddH2O n/a 10 mL
Total n/a 10 mL
Sterilize the solution with a 0.22 μm syringe filter. Store at 4 °C.
LB agar plate with kanamycin (1,000 mL)
Reagent Final concentration Quantity
Tryptone
Yeast extract
NaCl
10 g/L
5 g/L
10 g/L
10 g
5 g
10 g
Agar 15 g/L 15 g
50 mg/mL Kanamycin 50 μg/mL 1 mL
ddH2O n/a ~960 mL
Total n/a 1,000 mL
Adjust the pH to 7.4 with NaOH.
Autoclave at 121 °C for 15 min. Agar is added after sterilization.
Stir the medium on a stir plate and allow to cool to 30–40 °C.
Add 1 mL of 50 mg/mL kanamycin solution. Stir well.
Transfer 25 mL of the solution to 10 cm Petri dishes with a sterile pipette.
Allow the agar plates to cool and dry in a laminar flow hood for 30 min.
Store the plates at 4 °C in a plastic bag.
Terrific broth medium (1,000 mL)
Reagent Final concentration Quantity
Terrific broth
Glycerol
47.6 g/L
0.4% (v/v)
47.6 g
4 mL
ddH2O n/a ~960 mL
Total n/a 1,000 mL
Adjust the pH to approximately 7.4 with NaOH. Autoclave at 121 °C for 15 min. Store at RT.
1 M IPTG (10 mL)
Reagent Final concentration Quantity
IPTG 1 M 2.38 g
ddH2O n/a ~9.8 mL
Total n/a 10 mL
Sterilize the solution with a 0.22 μm syringe filter. Store at -20 °C.
Buffer A: start buffer (1,000 mL)
Reagent Final concentration Quantity
Tris
NaCl
20 mM
500 mM
2.4228 g
29.22 g
ddH2O n/a ~960 mL
Total n/a 1,000 mL
Adjust the pH to approximately 8.0 with HCl. Sterilize the solution with Nalgene Rapid-Flow. Store at 4 °C.
Buffer B: elution buffer (1,000 mL)
Reagent Final concentration Quantity
Tris
NaCl
20 mM
500 mM
2.4228 g
29.22 g
imidazole 500 mM 34.04 g
ddH2O n/a ~960 mL
Total n/a 1,000 mL
Adjust the pH to approximately 8.0 with HCl. Sterilize the solution with Nalgene Rapid-Flow. Store at 4 °C.
Buffer C: storage buffer (1,000 mL)
Reagent Final concentration Quantity
Tris
KCl
20 mM
200 mM
2.4228 g
14.91 g
MgCl2·6H2O 10 mM 2.03 g
Glycerol 10% (v/v) 100 mL
ddH2O n/a ~860 mL
Total n/a 1,000 mL
Adjust the pH to approximately 8.0 with HCl. Sterilize the solution with Nalgene Rapid-Flow. Store at 4 °C.
Buffer D: cleavage buffer (1,000 mL)
Reagent Final concentration Quantity
HEPES
KCl
20 mM
150 mM
4.77 g
11.18 g
MgCl2·6H2O 1 mM 0.2 g
TCEP 1 mM 0.25 g
Glycerol 10% (v/v) 100 mL
ddH2O n/a ~860 mL
Total n/a 1,000 mL
Adjust the pH to approximately 8.0 with HCl. Sterilize the solution with Nalgene Rapid-Flow. Store at -20 °C.
0.1 M NiSO4 (100 mL)
Reagent Final concentration Quantity
NiSO4·6H2O 0.1 M 2.63 g
ddH2O n/a ~96 mL
Total n/a 100 mL
Sterilize the solution with a 0.22 μm syringe filter. Store at RT.
0.1 M EDTA, 1M NaCl (100 mL)
Reagent Final concentration Quantity
EDTA 0.5 M 2.92 g
NaCl 1 M 5.84 g
ddH2O n/a ~96 mL
Total n/a 100 mL
Adjust the pH to 8.0 with NaOH. Sterilize the solution with a 0.22 μm syringe filter. Store at RT.
1 M NaOH (100 mL)
Reagent Final concentration Quantity
NaOH 1 M 4 g
ddH2O n/a ~96 mL
Total n/a 100 mL
Store at RT.
1 M HCl (120 mL)
Reagent Final concentration Quantity
12 M HCl 1 M 10 mL
ddH2O n/a 110 mL
Total n/a 120 mL
Store at RT.
SDS-PAGE running buffer (1,000 mL)
Reagent Final concentration Quantity
Tris
Glycine
125 mM
1.25 M
15.1 g
94 g
SDS 0.5% (w/v) 5 g
ddH2O n/a ~800 mL
Total n/a 1,000 mL
Store at RT.
Low-EDTA TE buffer, 1× (for DNA) (100 mL)
Reagent Final concentration Quantity
Tris
EDTA
10 mM
0.2 mM
0.1211 g
0.0012 g
ddH2O n/a ~100 mL
Total n/a 100 mL
Adjust the pH to 8.0 with HCl. Sterilize the solution with a 0.22 μm syringe filter. Store at RT.
Equipment
Amaxa Nucleofector IIb device (Lonza, model: AAB-1001)
BD Accuri C6 flow cytometer (BD Biosciences, catalog number: 23432)
Heracell VIOS 160i CO2 copper chamber incubator (Thermo Fisher Scientific, catalog number: 51030476)
Biological safety cabinet (Esco Airstream, model: AB2-4S8-CN)
Sonicator Q125 (Qsonica, model: Q125-220)
1/4 diameter sonica probe (Qsonica, catalog number: 4435)
Protein purification system (AKTA pure, catalog number: 29046665)
Fluorescence spectrophotometer F-7000 (HITACHI, model: F-7000)
IX71 inverted microscope (Olympus, model: IX71)
Snowflake ice machine (XUEKE, model: IMS-20)
Water bath (EYELA, model: NTT-2200)
Mini gel DNA electrophoresis system (Thermo Fisher, model: B1)
Gel imaging system (Azure Biosystems, model: C300)
Veriti 96- Well PCR thermal cycler (Applied Biosystems, catalog number: 4375786)
TAdvanced Twin PCR thermocycler (Biometra, catalog number: 2070212)
NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, model: ND-2000)
pH meter (Mettle Toledo, catalog number: 51302807)
Milli-Q water purification system (Millipore, model: MP0024)
Eppendorf centrifuge 5424 R (Eppendorf, catalog number: 5404000090)
Eppendorf high-speed refrigerated centrifuge 5910 R (Eppendorf, catalog number: 5942000598)
Thermo shaker (Yeasen, catalog number: 80440ES03)
Software
Prism v8, for statistics and data visualization (GraphPad, version 8.0.1.244)
Snapgene, view DNA sequence and primer design (version 4.2.4)
NUPACK: Nucleic Acid Package (https://nupack.org/)
Tm Calculator (version 1.16.5, https://tmcalculator.neb.com/#!/main)
CHOPCHOP, a web-based tool for sgRNA design. The website is for non-profit and academic use only (https://chopchop.cbu.uib.no/)
Expasy, conversion of DNA sequence to amino acid sequence (https://web.expasy.org/translate/)
AAT Bioquest: Protein Concentration Calculator (https://www.aatbio.com/tools/calculate-protein-concentration)
NEBioCalculator (https://nebiocalculator.neb.com/#!/dsdnaamt)
Procedure
Preparation of dsDNA and odsDNA as DNA donors to achieve green fluorescent protein KI at varying genomic loci in HEK293T cells
Design target sites for Lamin A/C, GAPDH, and AAVS1 loci, as examples. The donor DNA sequences contain the to-be-inserted sequence flanked by 50 nt homology arm.
The target sequences were selected by the online pipeline CHOPCHOP; an example is shown in Figure 1.
Note: Target sequences with fewer mismatches and higher scores.
Figure 1. Screenshot example of the CHOPCHOP website when selecting the target sequence
All the 1,010 and 2,500 bp gene fragments as donors were cloned into plasmids following general cloning procedures, including promoter region and EGFP sequence. HA sequences are underlined. The template sequences are shown in Table 1.
Table 1. Sequences of the target insert in donor plasmids used as PCR templates
Lamin A/C locus donor sequence (EF-1α core promoter-EGFP 1110bp)
ctttggtttttttcttctgtatttgtttttctaagagaagttattttctataggtcttgaaaggagtgggtcaattggctccggtgcccgtcagtgggcagagcgcacatcgcccacagtccccgagaagttggggggaggggtcggcaattgatccggtgcctagagaaggtggcgcggggtaaactgggaaagtgatgtcgtgtactggctccgcctttttcccgagggtgggggagaaccgtatataagtgcagtagtcgccgtgaacgttctttttcgcaacgggtttgccgccagaacacaggaagcttgccaccatggtgagcaagggcgaggagctgttcaccggggtggtgcccatcctggtcgagctggacggcgacgtaaacggccacaagttcagcgtgtccggcgagggcgagggcgatgccacctacggcaagctgaccctgaagttcatctgcaccaccggcaagctgcccgtgccctggcccaccctcgtgaccaccctgacctacggcgtgcagtgcttcagccgctaccccgaccacatgaagcagcacgacttcttcaagtccgccatgcccgaaggctacgtccaggagcgcaccatcttcttcaaggacgacggcaactacaagacccgcgccgaggtgaagttcgagggcgacaccctggtgaaccgcatcgagctgaagggcatcgacttcaaggaggacggcaacatcctggggcacaagctggagtacaactacaacagccacaacgtctatatcatggccgacaagcagaagaacggcatcaaggtgaacttcaagatccgccacaacatcgaggacggcagcgtgcagctcgccgaccactaccagcagaacacccccatcggcgacggccccgtgctgctgcccgacaaccactacctgagcacccagtccgccctgagcaaagaccccaacgagaagcgcgatcacatggtcctgctggagttcgtgaccgccgccgggatcactctcggcatggacgagctgtacaagtaagactctggtcagagatacctcagtggttttatactgaaggaaaaacacaagcaaaaaaaaaaaaaaagca
GAPDH locus donor sequence (EF-1α core promoter-EGFP 1110bp)
atggcctccaaggagtaagacccctggaccaccagccccagcaagagcactaggtcttgaaaggagtgggtcaattggctccggtgcccgtcagtgggcagagcgcacatcgcccacagtccccgagaagttggggggaggggtcggcaattgatccggtgcctagagaaggtggcgcggggtaaactgggaaagtgatgtcgtgtactggctccgcctttttcccgagggtgggggagaaccgtatataagtgcagtagtcgccgtgaacgttctttttcgcaacgggtttgccgccagaacacaggaagcttgccaccatggtgagcaagggcgaggagctgttcaccggggtggtgcccatcctggtcgagctggacggcgacgtaaacggccacaagttcagcgtgtccggcgagggcgagggcgatgccacctacggcaagctgaccctgaagttcatctgcaccaccggcaagctgcccgtgccctggcccaccctcgtgaccaccctgacctacggcgtgcagtgcttcagccgctaccccgaccacatgaagcagcacgacttcttcaagtccgccatgcccgaaggctacgtccaggagcgcaccatcttcttcaaggacgacggcaactacaagacccgcgccgaggtgaagttcgagggcgacaccctggtgaaccgcatcgagctgaagggcatcgacttcaaggaggacggcaacatcctggggcacaagctggagtacaactacaacagccacaacgtctatatcatggccgacaagcagaagaacggcatcaaggtgaacttcaagatccgccacaacatcgaggacggcagcgtgcagctcgccgaccactaccagcagaacacccccatcggcgacggccccgtgctgctgcccgacaaccactacctgagcacccagtccgccctgagcaaagaccccaacgagaagcgcgatcacatggtcctgctggagttcgtgaccgccgccgggatcactctcggcatggacgagctgtacaagtaagactctggtcagagatacctaagaggaagagagagaccctcactgctggggagtccctgccacactcagt
AAVS1 locus donor sequence (EF-1α core promoter-EGFP 1110bp)
gttctgggtacttttatctgtcccctccaccccacagtggggccactaggtaggtcttgaaaggagtgggtcaattggctccggtgcccgtcagtgggcagagcgcacatcgcccacagtccccgagaagttggggggaggggtcggcaattgatccggtgcctagagaaggtggcgcggggtaaactgggaaagtgatgtcgtgtactggctccgcctttttcccgagggtgggggagaaccgtatataagtgcagtagtcgccgtgaacgttctttttcgcaacgggtttgccgccagaacacaggaagcttgccaccatggtgagcaagggcgaggagctgttcaccggggtggtgcccatcctggtcgagctggacggcgacgtaaacggccacaagttcagcgtgtccggcgagggcgagggcgatgccacctacggcaagctgaccctgaagttcatctgcaccaccggcaagctgcccgtgccctggcccaccctcgtgaccaccctgacctacggcgtgcagtgcttcagccgctaccccgaccacatgaagcagcacgacttcttcaagtccgccatgcccgaaggctacgtccaggagcgcaccatcttcttcaaggacgacggcaactacaagacccgcgccgaggtgaagttcgagggcgacaccctggtgaaccgcatcgagctgaagggcatcgacttcaaggaggacggcaacatcctggggcacaagctggagtacaactacaacagccacaacgtctatatcatggccgacaagcagaagaacggcatcaaggtgaacttcaagatccgccacaacatcgaggacggcagcgtgcagctcgccgaccactaccagcagaacacccccatcggcgacggccccgtgctgctgcccgacaaccactacctgagcacccagtccgccctgagcaaagaccccaacgagaagcgcgatcacatggtcctgctggagttcgtgaccgccgccgggatcactctcggcatggacgagctgtacaagtaagactctggtcagagatacctgacaggattggtgacagaaaagcccccatccttaggcctcctccttccta
Lamin A/C locus donor sequence (EF-1α promoter-EGFP 2600bp)
ctttggtttttttcttctgtatttgtttttctaagagaagttattttctatgcccggcgagagatcacgtggggcgcggaggcggtgctgctggggcacggccgtccagcctcggcggccatatttttgaggggctgttcatctcgttcacacgctctgtccgccatgtttgtgagtggaagcgccattaccccttcaagcgactgaaggctgcagggcctctggtggcccgcatggggagaccagacccgccaggcccgcctttccgcactcagtccgggcttactttattttgtgagacagggtctcgcctagaggctccggtgcccgtcagtgggcagagcgcacatcgcccacagtccccgagaagttggggggaggggtcggcaattgaaccggtgcctagagaaggtggcgcggggtaaactgggaaagtgatgtcgtgtactggctccgcctttttcccgagggtgggggagaaccgtatataagtgcagtagtcgccgtgaacgttctttttcgcaacgggtttgccgccagaacacaggtaagtgccgtgtgtggttcccgcgggcctggcctctttacgggttatggcccttgcgtgccttgaattacttccacgcccctggctgcagtacgtgattcttgatcccgagcttcgggttggaagtgggtgggagagttcgaggccttgcgcttaaggagccccttcgcctcgtgcttgagttgaggcttggcctgggcgctggggccgccgcgtgcgaatctggtggcaccttcgcgcctgtctcgctgctttcgataagtctctagccatttaaaatttttgatgacctgctgcgacgctttttttctggcaagatagtcttgtaaatgcgggccaagatctgcacactggtatttcggtttttggggccgcgggcggcgacggggcccgtgcgtcccagcgcacatgttcggcgaggcggggcctgcgagcgcggccaccgagaatcggacgggggtagtctcaagctggccggcctgctctggtgcctggcctcgcgccgccgtgtatcgccccgccctgggcggcaaggctggcccggtcggcaccagttgcgtgagcggaaagatggccgcttcccggccctgctgcagggagctcaaaatggaggacgcggcgctcgggagagcgggcgggtgagtcacccacacaaaggaaaagggcctttccgtcctcagccgtcgcttcatgtgactccacggagtaccgggcgccgtccaggcacctcgattagttctcgagcttttggagtacgtcgtctttaggttggggggaggggttttatgcgatggagtttccccatactgagtgggtggagactgaagttaggccagcttggcacttgatgtaattctccttggaatttgccctttttgagtttggatcttggttcattctcaagcctcagacagtggttcaaagtttttttcttccatttaaggtgtcgtgaaaactaccccaagctggcctctgaggccaccatggctgtgagcaagggcgaggagctgttcaccggggtggtgcccatcctggtcgagctggacggcgacgtaaacggccacaagttcagcgtgtccggcgagggcgagggcgatgccacctacggcaagctgaccctgaagttcatctgcaccaccggcaagctgcccgtgccctggcccaccctcgtgaccaccctgacctacggcgtgcagtgcttcagccgctaccccgaccacatgaagcagcacgacttcttcaagtccgccatgcccgaaggctacgtccaggagcgcaccatcttcttcaaggacgacggcaactacaagacccgcgccgaggtgaagttcgagggcgacaccctggtgaaccgcatcgagctgaagggcatcgacttcaaggaggacggcaacatcctggggcacaagctggagtacaactacaacagccacaacgtctatatcatggccgacaagcagaagaacggcatcaaggtgaacttcaagatccgccacaacatcgaggacggcagcgtgcagctcgccgaccactaccagcagaacacccccatcggcgacggccccgtgctgctgcccgacaaccactacctgagcacccagtccgccctgagcaaagaccccaacgagaagcgcgatcacatggtcctgctggagttcgtgaccgccgccgggatcactctcggcatggacgagctgtacaagtaaaagcttggggatcaattctctagagctcgctgatcagcctcgactgtgccttctagttgccagccatctgttgtttgcccctcccccgtgccttccttgaccctggaaggtgccactcccactgtcctttcctaataaaatgaggaaattgcatcgcattgtctgagtaggtgtcattctattctggggggtggggtggggcaggacagcaagggggaggattgggaagacaatagcaggcatgctggggatgcggtgggctctatggcttctgaggcggaaagaaccagctgggcccagtggttttatactgaaggaaaaacacaagcaaaaaaaaaaaaaaagca
DNA donor PCR primer design and synthesis
Principle for design of dsDNA donor primers:
Only two homology arms (HAs) with 50 nt sequence are designed at the 5′ end of the synthesized forward and reverse primers.
Note: Longer HA sequences generally lead to higher HDR efficiency, but also likely increase the PCR difficulties.
Principle of design for odsDNA donor primers:
i. Likewise, two 50 nt HA sequences are designed at the 5′ ends of the synthesized forward and reverse primers, respectively.
ii. Specify the five consecutive phosphorothioate modifications in the synthesized 50 nt primers at designated positions, as needed. “*” represents phosphorothioate (PT) modification. Lower-case letters are HA sequences and upper-case letters depict exogenous KI sequences. We used the first PCR product as a template in order to save the cost of primer synthesis; long primers are not needed. The primer sequences are shown in Table 2.
Table 2. Synthesis of PCR primer sequences for dsDNA and odsDNA donors
Lamin A/C locus
EF-1α core promoter-EGFP (1,110 bp) primers or
EF-1α promoter-EGFP-ploy A signal (2,600 bp) primers (5′→3′)
L-50-F ctttggtttttttcttctgtatttgtttttctaagagaagttattttctaTAGGTCTTGAAAGGAGTGGG
L-50-R tgctttttttttttttttgcttgtgtttttccttcagtataaaaccactgAGGTATCTCTGACCAGAGTC
L-30S-F ctttggtttttttcttctgtatttgtttttc*t*a*a*g*agaagttattttcta
L-30S-R tgctttttttttttttttgcttgtgtttttc*c*t*t*c*agtataaaaccactg
L-20S-F ctttggtttttttcttctgta*t*t*t*g*tttttctaagagaag
L-20S-R tgctttttttttttttttgct*t*g*t*g*tttttccttcagtat
L-15S-F ctttggtttttttctt*c*t*g*t*atttgtttttctaag
L-15S-R tgcttttttttttttt*t*t*g*c*ttgtgtttttccttc
L-10S-F ctttggttttt*t*t*c*t*tctgtatttgttttt
L-10S-R tgctttttttt*t*t*t*t*tttgcttgtgttttt
L-5S-F ctttgg*t*t*t*t*tttcttctgtatttg
L-5S-R tgcttt*t*t*t*t*ttttttttgcttgtg
GAPDH locus EF-1α core promoter-EGFP primers
G-50-F atggcctccaaggagtaagacccctggaccaccagccccagcaagagcacTAGGTCTTGAAAGGAGTGGG
G-50-R actgagtgtggcagggactccccagcagtgagggtctctctcttcctcttAGGTATCTCTGACCAGAGTC
G-30S-F atggcctccaaggagtaagacccctggacca*c*c*a*g*ccccagcaagagcac
G-30S-R actgagtgtggcagggactccccagcagtga*g*g*g*t*ctctctcttcctctt
G-20S-F atggcctccaaggagtaagac*c*c*c*t*ggaccaccagcccca
G-20S-R actgagtgtggcagggactcc*c*c*a*g*cagtgagggtctctc
G-15S-F atggcctccaaggagt*a*a*g*a*cccctggaccaccag
G-15S-R actgagtgtggcaggg*a*c*t*c*cccagcagtgagggt
G-10S-F atggcctccaa*g*g*a*g*taagacccctggacc
G-10S-R actgagtgtgg*c*a*g*g*gactccccagcagtg
G-5S-F atggcc*t*c*c*a*aggagtaagacccct
G-5S-R actgag*t*g*t*g*gcagggactccccag
AAVS1 locus EF-1α core promoter-EGFP primers
A-50-F gttctgggtacttttatctgtcccctccaccccacagtggggccactaggTAGGTCTTGAAAGGAGTGGG
A-50-R taggaaggaggaggcctaaggatgggggcttttctgtcaccaatcctgtcAGGTATCTCTGACCAGAGTC
A-30S-F gttctgggtacttttatctgtcccctccacc*c*c*a*c*a
A-30S-R taggaaggaggaggcctaaggatgggggctt*t*t*c*t*g
A-20S-F gttctgggtacttttatctgt*c*c*c*c*t
A-20S-R taggaaggaggaggcctaagg*a*t*g*g*g
A-15S-F gttctgggtactttta*t*c*t*g*t
A-15S-R taggaaggaggaggcc*t*a*a*g*g
A-10S-F gttctgggtac*t*t*t*t*atctg
A-10S-R taggaaggagg*a*g*g*c*ctaag
A-5S-F gttctg*g*g*t*a*cttttatctg
A-5S-R taggaa*g*g*a*g*gaggcctaag
Lamin A/C loucs EF-1α promoter-EGFP-ploy A signal (2,600 bp) primers for esgRNA
L-12S-F ctttggttttttt*c*t*t*c*tgtatttgtttttctaagag
L-12S-R attgagatagatgagatagatgctttttttttt*t*t*t*t*t
Primers’ synthesis by Sangon Biotech. The dry powder for each primer was first diluted to 100 μM (stock solution) using low-EDTA TE buffer (10 mM Tris and 0.2 mM EDTA, pH 8.0), and then to 10 μM (working solution) using nuclease-free ddH2O.
PCR amplification of the donor DNA:
i. Set up the following PCR assembly reaction in a 400 μL mastermix volume. Reagents and volumes are shown in Table 3. Vortex for 5 s, spin down quickly, and make 50 μL aliquots using 0.2 mL strips of eight tubes.
Table 3. Components of PCR reaction for DNA donor
Reagent Volume
Q5 High-Fidelity 2× Master Mix 200 μL
10 μM forward primer 20 μL
10 μM reverse primer 20 μL
Template DNA (plasmids or PCR product) 80 ng
Nuclease-free water to 400 μL
ii. Add the reaction components in the order given. Aliquot 400 μL of the reaction solution in 50 μL portions into 0.2 mL strips of eight tubes.
iii. Perform assembly PCR using the cycling parameters below, as shown in Table 4.
Table 4. PCR reaction for DNA donor preparation
Cycle step Temperature Time Cycles
Initial denaturation 98 °C 30 s 1×
Denaturation 98 °C 5 s
35×
Annealing 65 °C 15 s
Final extension 72 °C 1 min/2 min 20 s 1×
Hold 4 °C Hold 1×
Note: Set up the extension time for 1 min for 1,110 bp or 2 min 20 s for 2,600 bp.
Purification of PCR DNA donors using GeneJET PCR Purification kit.
Add a 1:1 volume of binding buffer to completed PCR mixture (for every 400 μL of reaction mixture, add 400 μL of binding buffer). Mix thoroughly by pipetting up and down gently.
Transfer up to 800 μL of the solution from step A3a to the GeneJET purification column. Centrifuge at 12,000× g for 60 s. Discard the flowthrough.
Add 700 μL of wash buffer to the GeneJET purification column. Centrifuge at 12,000× g for 60 s. Discard the flowthrough and place the purification column back into the collection tube.
Centrifuge the empty GeneJET purification column for an additional 1 min to completely remove any residual wash buffer.
Transfer the GeneJET purification column to a clean 1.5 mL microcentrifuge tube. Add 25 μL of nuclease-free water to the center of the GeneJET purification column membrane and centrifuge at 12,000× g for 1 min. To increase the yield, the effluent can be added to the GeneJET purification column membrane and the operation repeated.
Measure the DNA concentration using NanoDrop 2000. Wipe NanoDrop 2000 with Kimwipes disposable wipers before and after use. Store the purified dsDNA donor in -20 ℃ refrigerator.
Note: The PT-modified DNA product (odsDNA precursor) requires a subsequent Lambda exonuclease digestion treatment to prepare odsDNA donor template.
Preparation of odsDNA donor:
Set up the following digest reaction in a 300 μL volume a clean 1.5 mL microcentrifuge tube. Reagents and volumes are shown in Table 5.
Table 5. Components to odsDNA donor digestion reaction
Reagent Volume
Lambda exonuclease reaction buffer (10×) 30 μL
Lambda exonuclease 6 μL
Purified odsDNA precursor (3.f) 18 μg
Nuclease-free H2O to 300 μL
Digest at 37 °C for 60 min in a dry-bath incubator.
Purification of odsDNA donors using GeneJET PCR Purification kit:
Add a 1:1 volume of binding buffer to the PCR mixture (for every 300 μL of reaction mixture, add 300 μL of binding buffer). Mix thoroughly by pipetting up and down gently.
Transfer up to 600 μL of solution from step A5a to the GeneJET purification column. Centrifuge for 60 s. Discard the flowthrough.
Add 700 μL of wash buffer to the GeneJET purification column. Centrifuge for 60 s. Discard the flowthrough and place the purification column back into the collection tube.
Centrifuge the empty GeneJET purification column for an additional 1 min to completely remove any residual wash buffer.
Transfer the GeneJET purification column to a clean 1.5 mL microcentrifuge tube. Add 25 μL of nuclease-free water to the center of the GeneJET purification column membrane and centrifuge for 1 min. To increase the yield, the eluent can be added back to the GeneJET purification column membrane and centrifuge again.
Measure the DNA concentration using NanoDrop 2000. Store the odsDNA donor in -20 ℃ refrigerator.
Expression and purification of Cas9 and Cas9-PCV2 fusion proteins in E. coli cells
Transform the competent E. coli (Rosetta) cells with the expression plasmid: pET28b-3NLS-Cas9-3NLS-His (pET 28b-3NLS-Cas9-PCV2-3NLS-His) and grow on LB plates (kanamycin+) overnight.
The next day, inoculate one colony into 25 mL of fresh LB medium (kanamycin+) from the LB plates (kanamycin+) and incubate in a shaker (200 rpm) at 37 ℃ overnight.
Inoculate 10 mL of overnight culture to a flask containing 1L of fresh Terrific Broth (TB) medium (kanamycin+) and shake at 200 rpm at 37 ℃. Grow the bacteria until the OD600 reaches ~0.6–0.8.
Note: The optimal growth time for TB is different, which requires OD600 to be more than 1.0–1.5 prior to IPTG induction.
Cool on ice or in a cold room for 20 min, then add IPTG to a final 0.2 mM concentration. Continue the shaking at 18 ℃ overnight (18–20 h).
The next day, pellet the culture medium and weigh the pellet. Resuspend the cells with buffer A at ~6 mL/g.
Add 50 μL of β-mercaptoethanol to per 50 mL of bacterial suspension. Sonication: set up at 35% amplitude, 1.5 s ON, 8.5 s OFF, to get the whole ON time for ~10 min, for a total time of ~66.7 min.
Note: The parameters vary depending on the sonicator; the goal is to attain a homogeneous lysate (more whitish in appearance).
Spin the lysate at 16,000× g for 30 min at 4 ℃ and transfer supernatant to a new 50 mL tube.
Filter the supernatant through a 0.45 μm filter.
Equilibrate a 5 mL His-trap column (GE) with buffer A (at least 25 mL).
Note: If using a previously used column, it can be recovered through the washing cycle by NaOH, ddH2O, and HCl, and then keep the column submerged in 5 mL of ddH2O.
Cleaning of His-trap column:
Note: This step is only required for recovery of old columns.
Before use, first wash with 10 mL of 0.1 M NiSO4, 10 mL of ddH2O, and finally with 20 mL of buffer A.
In use: Slowly compress the pipette to ensure that the filter filters out drop by drop (not forming a liquid flow).
After use: ddH2O wash 10 mL
0.1 M EDTA, 1 M NaCl wash 10 mL
ddH2O wash 10 mL
1 M NaOH wash 10 mL
ddH2O wash 10 mL
1 M HCl wash 10 mL
ddH2O wash 10 mL
Note: The column must be kept in water.
Filter to ensure that there are no air bubbles in the column. Store at 4 ℃ .
Next purification step using FPLC protocol.
Note: The buffer needs to be prepared in advance! The filtered column should not be sucked by water and cannot contact other solutions (so ensure that all connections are correct). Instrument settings: place the start buffer in Pump A1, place the elution buffer in Pump B1, select the column connection line of position 2, place the connection point of A1 upstream, and place the connection point of B1 downstream.
Wash step:
i. Open the AKTA Explorer UNICORN software, select Execute... in Manual, select Pumps B in wash, and then click Inlet.
ii. Select position 2 for the middle circle and waste for the filtered interface.
iii. Select Pumps A wash after the automatic stop. The steps are as above.
iv. Finally, clean the whole system flow, select the flow rate of 5 mL/min, pressure OFF, wash for 10 min.
Note: Use all buffer A and gradient options as 0% buffer B.
All tubes after elution need to be replaced with new ones. Starting from No. 1, approximately forty 1.5 mL tubes are placed in order. Lift the outflow tip up, move directly onto the first tube, and position the column in place.
Set up the flow rate and channel in the Manual option.
i. Flow rate: 1.5 mL/min
ii. Pressure: System pressure
iii. Gradient: Target 4% buffer B
iv. Fraction collection-fraction volume: 5 mL
v. Alarms: Alarm system pressure: High clarm-1.00 Mpa
Note: After completing the setting, double-check whether the elution system functions automatically in order.
Once the mAU value stabilizes, change the target of gradient to 10% buffer B and set the time to 15 min, in order to wash possible non-specifically bound protein. This gradient takes ~15 min to go from 4% buffer B to 10% buffer B.
Wait again until it stabilizes and the peaks disappear; then, change the gradient to 20% buffer B and set the time to 15 min.
Note: The safety imidazole concentration for Cas9 protein ranges between 50 and 250 mM. After reaching 250 mM, it indicates substantial absence of impurities if no peak present.
Collect the eluent at peaks approximately within the range of 50–250 mM imidazole.
Run SDS-PAGE for gel verification of the protein molecular weight.
In the peak effluent, take 8 μL of crude protein product from each tube, add 2 μL of SDS-PAGE protein loading buffer (5×), and denature at 95 ℃ for 10 min.
Note: The first flowthrough serves as control. Marker: add 5 μL BeyoColor Prestained Color Protein marker 10–170 kDa. Running conditions: 150 V, 30 min.
Stain in protein staining solution after completion.
Note: Microwave for 15 s or shake for 5 min.
Then, shake for 30 min.
Check the protein sizes: Cas9 is ~163 kDa and Cas9-PCV2 is ~179 kDa. Gels are shown in Figure 2.
Figure 2. Purification and activity detection in vitro of Cas9 and Cas9-PCV2 fusion proteins. (A) Verification of purified spCas9 protein by SDS-PAGE. (B) Verification of purified Cas9-PCV2 fusion protein by SDS-PAGE.
Compare the results to find 5–6 brightest fractions and conduct the protein concentration subsequently as below.
Note: During the running process, the desktop centrifuge is precooled at 4 ℃.
Protein concentration:
Centrifuge filters (pre-cool the 10 K 50 mL and 3 K 15 mL filters on ice).
Add the selected 5–6 fractions of the sample to the pre-cooled 10 K 50 mL filter.
Spin down in the pre-cooled centrifuge for 30 min, observe the upper liquid, and stop the centrifuge any time when the leftover is ~2.5 mL volume.
Using PD-10 column to desalt the concentrated samples.
Approximately 10 tubes of 100 μL of Coomassie Brilliant Blue are required to detect in real time when elute has protein presence and to compare protein concentrations.
Wash the PD-10 column with Cas9 storage buffer three times for 5 min every time.
Then, add 2.5 mL of concentrated solution to PD10, followed by centrifugation. Add 500 μL of storage buffer, collect the elute with a 1.5 mL centrifuge tube, add 500 μL each time for a total of seven times, and collect seven tubes for each 500 μL product.
Take out 3 μL of the product and mix with Coomassie Brilliant Blue, distinguish the blue brightness, select the brightest five tubes of product, transfer to the second PD-10, which has been exchanged with Cas9 buffer, and perform the second desalting.
Measure the brightness, pick up four tubes with the highest blue staining, and proceed to the next protein concentration.
Note: Perform these steps in a cold room; load PD-10 column with less than or equal to 2.5 mL volume.
Concentrate the desalted protein.
Add 2 mL of the desalted product to a 3 K 15 mL pre-cooled centrifuge filter and then centrifuge in a pre-cooled centrifuge at 4,000× g for approximately 30 min. Pay attention to the remaining volume and stop centrifugation when the leftover volume reaches 500 μL.
Prepare 20 μL aliquots in the 1.5 mL tubes in a cold room.
Store at -80 ℃ freezer.
Keep one tube of product for concentration determination.
The absorption peak of protein at 280 nm was measured by NanoDrop 2000.
Calculate final concentration in software Expasy and AAT Bioquest: Protein Concentration Calculator.
Testing enzymatic activity in vitro.
Examine the activity of Cas9 protein, with an in vitro cleavage assay as below.
i. Set up the following cleavage reaction in a 20 μL volume. Reagents and volumes are shown in Table 6; add the reaction components in the order given.
Table 6. Reaction mixture for in vitro cleavage assay
Reagent Volume
Cleavage buffer (10×) 2 μL
Substrate DNA 100 ng
Cas9 (or Cas9-PCV2) 0.3 μg
sgRNA 130 ng
Nuclease-free water up to 20 μL
ii. Incubate at 37 °C for 30 min.
iii. Visualize the cleaved bands by 3% agarose gel electrophoresis with 1× TAE buffer. Gel image is shown in Figure 3.
Figure 3. In vitro cleavage assay for examination of the Cas9 and Cas9-PCV2 activity
Measurement of the enzymatic activity for the Cas9-PCV2 fusion protein.
i. The ssDNA probe was synthesized by Sangon Biotech:
PCV2-linker Sequence (5′→3′): BHQ1-TAAGTATTACCAGAAA/i6FAMdT/cctcttgtcccacagat atccagaaccctgaccctgccgtgtaccagct
ii. Dilute the PCV2 linker with ddH2O to a final 0.02, 0.06, and 0.10 μM, respectively.
iii. The ssDNA probe was co-incubated with Cas9-PCV2 at 37 °C, and the fluorophore intensity was detected using 0.02 μM Cas9-PCV2 protein along with varying concentrations of ssDNA probes by fluorescence spectrophotometer F-7000.
Note: When the ssDNA probe is cleaved by the Cas9-PCV2 fusion protein, the FAM fluorophore is released from the quenching group BHQ1 and begins to emit fluorescence, which is monitored in real time by a fluorescence spectrometer.
Synthesis and purification of sgRNA to target gDNA in HEK293T cells (GeneArt Precision gRNA Synthesis kit)
Design sequences as below for the Target F1 forward and Target R1 reverse primers required for synthetic sgRNA template assembly. Sequences are shown in Table 7:
Target F1: TAATACGACTCACTATAG + target sequence (20 nt)
Target R1: TTCTAGCTCTAAAAC + target sequence reverse complement (20 nt)
Note: If the target sequence already contains a 5′ G, you can keep it, which will result in an extra G being added from the T7 promoter primer. Alternatively, you can remove the first G from the target sequence, which will be added back by the T7 promoter primer.
Table 7. Primers for in vitro translation of sgRNA
Locus Primer sequence (5′→3′)
Lamin A/C TAATACGACTCACTATAGAGAGAAGTTATTTTCTACAG
TTCTAGCTCTAAAACCTGTAGAAAATAACTTCTCT
GAPDH TAATACGACTCACTATAGAGCCCCAGCAAGAGCACAAG
TTCTAGCTCTAAAACCTTGTGCTCTTGCTGGGGCT
AAVS1 TAATACGACTCACTATAGACAGTGGGGCCACTAGGGAC
TTCTAGCTCTAAAACGTCCCTAGTGGCCCCACTGT
Primers were synthesized by Sangon Biotech and were diluted using nuclease-free water to final 0.3 μM concentration (working solution).
sgRNA transcription in vitro using GeneArt Precision gRNA Synthesis kit:
Set up the following PCR assembly reaction in a 25 μL volume and prepare the reaction components as below; reagents and volumes are shown in Table 8.
Table 8. Components to make gRNA DNA template
Reagent Volume
Phusion High-Fidelity PCR Master Mix (2×) 12.5 μL
Tracr Fragment + T7 Primer Mix 1 μL
0.3 μM Target F1/R1 primer mix 1 μL
Nuclease-free water 10.5 μL
Perform assembly PCR using the cycling parameters below, as shown in Table 9.
Table 9. PCR parameters to generate gRNA DNA template
Cycle step Temperature Time Cycles
Initial denaturation 98 °C 10 s 1×
Denaturation 98 °C 5 s
32×
Annealing 55 °C 15 s
Final extension 72 °C 1 min 1×
Hold 4 °C Hold 1×
In vitro transcription (IVT) of gRNAs; reagents and volumes are shown in Table 10.
Table 10. Components of IVT reaction
Reagent Volume
NTP mix (100 mM each of ATP, GTP, CTP, UTP) 8 μL
gRNA DNA template (from PCR assembly) 6 μL
5× TranscriptAidTM Reaction Buffer 4 μL
TranscriptAidTM Enzyme Mix 2 μL
Mix the tubes thoroughly and spin down quickly. Set up the IVT reaction for 3 h at 37 °C.
Note: All operations are preferably conducted in RNase-free workstation.
Removal of the DNA template by DNase I digestion:
Incubate the IVT reaction mix with 1 μL of DNase I (1 U/μL) immediately following the IVT reaction and incubate at 37 °C for 15 min.
Purification of sgRNA using GeneArt gRNA Clean-up kit.
Dilute the IVT reaction to 200 μL with nuclease-free water and add 100 μL of binding buffer. Mix by pipetting.
Add 300 μL of ethanol (> 96%) and mix by pipetting.
Transfer the mixture to the GeneJET RNA Purification Micro Column (preassembled with a collection tube) and centrifuge for 30–60 s at 14,000× g. Discard the flowthrough.
Wash the bound RNA with 700 μL of wash buffer 1 and wash buffer 2.
Centrifuge the empty purification column for an additional 60 s at 14,000× g to completely remove any residual wash buffer.
Transfer the purification column to a new 1.5 mL RNase-free EP tube.
Add 10 μL of nuclease-free water to the center of the column filter and centrifuge at 14,000× g for 60 s to elute the sgRNA.
Measure the concentration of sgRNA using NanoDrop 2000.
Store the sgRNA at -80 °C.
Note: All operations are preferably conducted in RNase-free workstation.
Preparation of the RNP complex for odsDNA attaching to Cas9-PCV2 fusion protein
Design and synthesis of the PCV2 linker.
Because PCV2 protein recognizes DNA sequence (AAGTATT^ACCAGAAA) in 5′ to 3′ direction, odsDNA has 3′ overhang, and a PCV2 linker is needed as a bridge to attach odsDNA to Cas9-PCV2 fusion protein.
Note: “^” represents the cleavage position of the PCV2 protein.
The PCV2 linker is divided into two parts: the 5′ end is a fixed sequence recognized by the PCV2 protein for cleavage, followed by subsequent covalent linkage to PCV2 protein, and the 3′ end is a sequence that anneals to the 3′ overhang of odsDNA.
The PCV2 linker sequence for the Lamin A/C locus is shown in Table 11.
Table 11. PCV2 linker sequence for the Lamin A/C locus
Lamin A/C locus Cas9-PCV2 linker sequence（5′→3′）
PCV2 linker AAGTATTACCAGAAAtgcttttttt
Note: The upper-case letters are the set sequences recognized by the PCV2 protein and the lower-case letters are the sequences that anneal to the odsDNA of the Lamin A/C locus.
Synthesis of PCV2 linker was by Sangon Biotech and diluted to 10 μM with nuclease-free water.
PCV2 linker annealing to odsDNA.
We use odsDNA with 10 bases-overhang to anneal to PCV2 linker with 10 nt paired bases.
Set up the following annealing reaction in a 10 μL volume; reagents and volumes are shown in Table 12.
Table 12. Components of annealing reaction
Reagent Molar amount Volume
PCV2 linker 7.5 pmol 0.75 μL
odsDNA (1 μg/μL) 7.5 pmol 5 μL
5× annealing buffer - 2 μL
Nuclease-free water - to 10 μL
Perform annealing reaction using the cycling parameters as shown in Table 13.
Table 13. Annealing reaction to generate odsDNA/PCV2 linker complex
Cycle step Temperature Time Cycles
Denaturation 65 °C 5 min 1×
Annealing 65 °C (-0.1 ℃/8 s) to 25 °C ~60 min 1×
Hold 4 °C Hold 1×
Prepare RNP complex composed of odsDNA/PCV2 linker/Cas9-PCV2 fusion protein.
In order to verify that odsDNA/PCV2 linker can be effectively attached to the Cas9-PCV2 protein, different molar ratios of 0:1, 1:1, 2:1, 3:1, and 4:1 were incubated with Cas9-PCV2 protein and then verified by 2% agarose gel. As shown in Figure 4, when the molar ratio of Cas9-PCV2 fusion protein to odsDNA/PCV2 linker was 3:1, the free odsDNA/PCV2 linker below was almost completely reacted, and all odsDNA/PCV2 linker was attached to the Cas9-PCV2 fusion protein.
Figure 4. Verification for the attachment of odsDNA to Cas9-PCV2 protein.
Efficient tethering of odsDNA donors to Cas9-PCV2 when the ratio of Cas9-PCV2 to pre-annealed odsDNA/PCV2 linker reaches > 3:1.
Add 30 pmol Cas9-PCV2 fusion protein to the odsDNA/PCV2 linker complex from step D2.
Because Cas9-PCV2 fusion protein is much more abundant than odsDNA, almost all odsDNA is reacted completely attached to Cas9-PCV2 fusion protein.
The working condition of PCV2 protein requires the presence of at least 1 mM Mg2+. Since the buffer C: storage buffer of Cas9-PCV2 protein has 10 mM MgCl2, no additional Mg2+ is needed here.
Nucleofection of the odsDNA/Cas9-PCV2/sgRNA RNP complex into HEK293T cells
Before nucleofection, sub-culture the HEK293T cells three days prior to nucleofection experiments using Dulbecco’s modified Eagle’s medium and 10% fetal bovine serum.
Nucleofection of HEK293T cells with odsDNA/Cas9-PCV2/sgRNA:
Cell preparation
i. Optimal cell confluency for nucleofection: 80%–90%. Higher cell densities may cause lower Nucleofection® efficiencies.
ii. Remove the medium from the cultured cells and wash cells once with 1× PBS. Wash with at least the same volume of 1×PBS as culture media.
iii. For harvesting, incubate the cells for ~5 min at 37 °C with TrypLE Express reagent.
iv. Neutralize the trypsinization reaction with culture medium containing 10% FBS. The majority of the cells (>90%) will be detached. The culture medium, PBS, and TrypLE are opened and sealed using PARAFILM sealing film.
Preparation for odsDNA/Cas9-PCV2/sgRNA RNP complex:
i. To prepare the Cas9-PCV2 RNPs complex, mix the odsDNA/PCV2 linker/Cas9-PCV2 fusion protein complex with 90 pmol sgRNA and incubate for 10 min at 37 °C.
ii. Perform nucleofection immediately as below.
Nucleofection
i. Ensure that the entire supplement is added to the Nucleofector® solution. The ratio of Nucleofector® solution to supplement is 4.5:1. For a single reaction, use 82 μL of Nucleofector® solution plus 18 μL of supplement to make 100 μL of total reaction volume.
ii. Prepare 6-well plates by filling the appropriate number of wells with 1 mL of supplemented culture media and pre-incubate/equilibrate plates in a humidified 5% CO2 incubator at 37 °C.
iii. Harvest the cells by trypsinization.
iv. Count the cell numbers with a hemacytometer or an automated cell counter as preferred.
v. Centrifuge the required number of cells (1 × 106 cells per sample) at 200× g for 10 min at room temperature. Remove supernatant completely.
vi. Resuspend the cell pellet carefully in 100 μL of room-temperature Nucleofector® solution per sample.
Note: Do not leave the cells in Nucleofector® solution for an extended time (i.e., longer than 15 min), as this may reduce both the cell viability and the gene transfer efficiency.
i. Combine 100 μL of cell suspension with odsDNA/Cas9-PCV2/sgRNA complex. Table 14 shows the different samples.
Table 14. One Nucleofection® sample contains
odsDNA sample odsDNA sample dsDNA sample
1 × 106 HEK293T cells 1 × 106 HEK293T cells 1 × 106 HEK293T cells
odsDNA/Cas9-PCV2/sgRNA complex odsDNA, Cas9/sgRNA complex dsDNA, Cas9/sgRNA complex
100 μL Cell Line Nucleofector® Solution V 100 μL Cell Line Nucleofector® Solution V 100 μL Cell Line Nucleofector® Solution V
ii. Transfer cell/DNA suspension into a certified cuvette (sample must cover the bottom of the cuvette without air bubbles). Close the cuvette with the cap.
iii. Select the appropriate Nucleofector® Program D-032 for Nucleofector® 2b Device.
iv. Insert the cuvette with cell/DNA suspension into the Nucleofector® Cuvette Holder and apply the selected program by pressing the X-button.
v. Take the cuvette out of the holder once the program is finished.
vi. Immediately replenish with ~500 μL of the pre-equilibrated culture medium to the cuvette and gently transfer the mixture into the prepared 6-well plate (final volume 1.5 mL media per well). Use the supplied pipettes and avoid repeated aspiration of the sample.
Post nucleofection:
Incubate the cells in a humidified 37 °C/5% CO2 incubator until further analysis at defined time points. Nucleofection of odsDNA with Cas9/sgRNA into HEK293T cells showed that universal odsDNA donors had higher KI rates than dsDNA donors in Figure 5. When Cas9-PCV2 was used to attach to odsDNA, the KI rate was 5-fold higher than dsDNA in Figure 6.
Figure 5. Knock-in (KI) rates in HEK293T cells with different donors. A panel of three different genomic loci were selected for the 1,010 bp odsDNA donor KI in HEK293T cells. The dsDNA donors were used as controls. Data were collected 15 days after nucleofection. The KI rates were calculated as the averaged percentages for the total number of EGFP-positive cells (donor KI) divided by the total number of RNP-electroporated cells.
Figure 6. Different lengths of DNA fragment knock-in (KI) using the odsDNA attach to Cas9-PCV2 protein strategy. (A) The 1,010 bp donor templates, including Cas9-bound dsDNA and 10 nt odsDNA as well as Cas9-PCV2-bound 10 nt odsDNA, were electroporated to target the Lamin A/C locus in HEK293T cells. The KI rates were plotted for side-by-side comparison. Data were collected 15 days after nucleofection. (B) The 2,500 bp donor templates, including Cas9-bound dsDNA and 10-nt odsDNA as well as Cas9-PCV2-bound 10-nt odsDNA, were electroporated to target the Lamin A/C locus in HEK293T cells. The KI rates were plotted for side-by-side comparison. Data were collected 21 days after nucleofection.
Data analysis
The KI rates of different donors across varying loci in HEK293T cells were measured by BD Accuri C6 flow cytometer and plotted in Prism 8 software.
The KI rates of odsDNA attached to Cas9-PCV2 fusion protein at Lamin A/C locus in HEK293T cells was measured by BD Accuri C6 flow cytometer and plotted in Prism 8 software.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Han et al. (2023). Efficient precise integration of large DNA sequences with 3′-overhang dsDNA donors using CRISPR/Cas9. Proc. Natl. Acad. Sci. U.S.A. (Figure 4, panel B, C and D).
Tei et al. (2023). Comparable analysis of multiple DNA double-strand break repair pathways in CRISPR-mediated endogenous tagging. bioRxiv. (Figure 3, panel C and D).
Acknowledgments
This work was supported by the Ministry of Science and Technology of China (2019YFA0802600 and 2020YFA0710700), the National Natural Science Foundation of China (Nos. 21991132, 52033010, 52021002, 31970793, and 32170856). The current protocol is mainly derived from Han et al. (2023).
Competing interests
The authors declare no competing financial interests.
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4,854 | https://bio-protocol.org/en/bpdetail?id=4854&type=0 | # Bio-Protocol Content
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Princeton RAtlas: A Common Coordinate Framework for Fully cleared, Whole Rattus norvegicus Brains
ED Emily Jane Dennis
PB Peter Bibawi
ZD Zahra M. Dhanerawala
LL Laura A. Lynch
SW Samuel S.-H. Wang
CB Carlos D. Brody
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4854 Views: 910
Reviewed by: Oneil Girish BhalalaXiaochen SunEmma Puighermanal
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Abstract
Whole-brain clearing and imaging methods are becoming more common in mice but have yet to become standard in rats, at least partially due to inadequate clearing from most available protocols. Here, we build on recent mouse-tissue clearing and light-sheet imaging methods and develop and adapt them to rats. We first used cleared rat brains to create an open-source, 3D rat atlas at 25 μ resolution. We then registered and imported other existing labeled volumes and made all of the code and data available for the community (https://github.com/emilyjanedennis/PRA) to further enable modern, whole-brain neuroscience in the rat.
Key features
• This protocol adapts iDISCO (Renier et al., 2014) and uDISCO (Pan et al., 2016) tissue-clearing techniques to consistently clear rat brains.
• This protocol also decreases the number of working hours per day to fit in an 8 workday.
Graphical overview
Keywords: Tissue clearing Light-sheet imaging Rattus norvegicus Image registration iDISCO uDISCO Rat Neuroscience
Background
Recent developments in tissue clearing and imaging allow us to gather whole-brain connectivity in intact brains, together with classic neuronal tracing techniques.Though tissue fixation and preparation predated neuroscience, its use was refined by Golgi and further modified and popularized by Ramón y Cajal (1911). To this day, after experiments, neuroscientists often remove the brain and slice it into sections that are then mounted to microscope slides for imaging. This allows for both post-hoc verification of experimental targeting and, if paired with staining or immunohistochemistry, learning about the tissue. However, this process is hard to automate and time-consuming: each step of fixation, removal, slicing, and imaging for each slice can take hours. Additional immunohistochemistry or staining can take weeks. To decrease the amount of labor required to process each brain, researchers will frequently only keep a subset of the brain slices near the relevant regions of study. Often this is more than sufficient, but this approach can easily miss potentially relevant information in the discarded sections, and sectioning itself can introduce information and aberrations to data that are difficult to identify post-hoc. Address this issue, several groups have developed methods to clear the lipids and pigments from whole brains, allowing for imaging through the entire tissue without the need for sectioning and often dramatically decreasing imaging time.
Building on advances from the early 1900s, tissue clearing has experienced a resurgence in neuroscience over the last 20 years. Methods like CUBIC (Susaki et al., 2015), CLARITY (Chung and Deisseroth, 2013), FluoClearBABB (Schwarz et al., 2015), and the DISCO methods (Renier et al., 2014; Pan et al., 2016) have become popular with researchers studying mice (see Molbay et al., 2021 for a review) yet those of us studying rats have not yet fully embraced this new methodology. One major impediment for broad acceptance of these methods in larger-brained animals is that existing clearing techniques do not consistently produce fully cleared, intact brains, prohibiting the study of subcortical structures in whole brains and decreasing their utility over slice histology. However, several advances have been made: Woo and colleagues nearly doubled the length of the mouse-centric CUBIC method to 20 days to improve clearing rats (Woo et al., 2018); Branch and colleagues successfully cleared entire rat brains by splitting samples into two hemispheres before clearing, together with increasing the timing of stages of the mouse iDISCO protocol (Branch et al., 2019); and a third method, FluoClearBABB, cleared the majority of tissue from an intact rat brain in 8 days while controlling the pH during dehydration, which, similar to uDISCO, preserved fluorescence, though the clearing at the center of the tissue is unclear from their manuscript (Stefaniuk et al., 2016).
Similar advances in imaging techniques allow for faster image acquisition. Typically, slices are imaged on confocal or 2-photon microscopes, which use point scanning methods that scan across and through tissues to excite and observe fluorescence. In contrast, light-sheet microscopy images planes instead of points, decreasing the time needed to go through the same volume of tissue. Like other methods key to modern developments, light-sheet microscopy also was first proposed in 1902 (Siedentopf and Zsigmondy, 1902), but recent developments in CMOS sensors that can acquire large images quickly, as well as advances in light sculpting (Keller et al., 2008), moving illumination and detection (Ahrens et al., 2013), and others [see review Hillman et al. (2019)] were key in making whole-brain imaging available to neuroscientists.
Inspired by these advances, we set out to produce two reliable, worker-friendly protocols for efficient clearing of intact rat brains, based on the uDISCO and iDISCO protocols. Coupled with light-sheet imaging, we can now image intact rat brains, including subcortical structures. We use these clear brains to form a light-sheet-based rat brain template for the rat neuroscience community.
Whole-brain clearing of Rattus norvegicus brains
We first set out to improve existing clearing techniques to work consistently on whole adult Rattus norvegicus brains of any size or sex. Inspired by many, we primarily focused on the DISCO techniques and the timing of stages, as this was the most effective in our preliminary explorations. We successfully made a worker-friendly protocol (Figure1A, maximum 8 h days, compared with 10–16 h days in other protocols) that also resulted in consistently clear brains, visible in the field-standard qualitative evaluations (Figure1B and 1C) and a newer quantitative evaluation method, Fourier ringcorrelation–based quality estimation (FRC-QE) (Figure 1E) (Preusser et al., 2021). When applied appropriately, FRC-QE allows for comparable measurements across experiments and provides quantitative data in addition to the field-standard qualitative data.
Figure 1. Modifications to iDISCO and uDISCO protocols allow for fully cleared ratbrains. A. Schematics of iDISCO protocol and the modified rat iDISCO and uDISCOprotocols (abbreviations DCM, BABB, and DBE all refer to solutions inRecipes). B. Example images of brains before clearing and after clearing with each protocol from the schematic. The uncleared brain is approximately 2.5 cm long and 1.7 cm wide. All images were taken on the same piece of paper for easy comparison. C. Computational slices through each clearedbrain in B. Labels (a253, k320, z268) reflect the animal’s name. D. Drawing showing the orientation of the brain (left) and the FRC-QE scores (right) for each brain in B along the coronal axes in slice units. E. Quantification of the values in (D), with a Bonferroni-corrected t-test. Horizontal white bars are median values, and the violin limits are the 10% (bottom) and 90% (top) of the data. Asterisks indicate significance p < 0.05 compared with iDISCO a253.
Princeton RAtlas
Once we could reliably obtain fully cleared brains, we set out to make a common coordinate framework for rats using whole-brain, light-sheet imaging. We imaged 18 brains and chose the best eight for an averaged template brain. We chose these eight based on a manual inspection of quality and based on external genitalia: we chose four male and four female brains without obvious damage or imaging aberrations, like streaking. The four female brains were all of sufficient quality that we did not need to procure additional brains, and we chose four of the male brains to match our number of females. We used an iterated, paired averaging scheme to create the seed brain (Figure 2A) and then aligned each brain to this seed, averaged the result to create the new seed, and repeated this process two additional times. The final averaged result of all brains aligned to this final seed brain became the Princeton RAtlas (PRA, Figure 2B). This coordinate space is agnostic to the sex of the animal. We also provide mPRA, a common coordinate space using only animals with visible penis and testes, and fPRA, a common coordinate space using only animals without visible penis or testes.
Figure 2. Creation of the Princeton RAtlas (PRA). A. Schematics showing the paired alignments to create the seed brain (left, middle) and the multilevel refinement resulting in the PRA (right). B.Summed axial projection of the PRA. mBrains and fBrains are male and femalebrains respectively, defined by external genitalia.
Materials and reagents
Biological materials
Long-Evans rats [Hilltop Lab Animals Inc., Long Evans Hla(LE)CVF, Scottdale, PA] of any sex. Animals used in this study were 3–12 months and fed ad libitum from 290 to 597 g, though older and larger animals have since been successfully cleared but are not included in this focused manuscript.
Reagents
Ketamine (MedVet International, catalog number: RXKETAMINE, CAS: 6740-88-1)
Xylazine (MedVet International, catalog number: RXXYLAZINE, CAS: 7361-61-7)
Isoflurane (shopmedvet.com, catalog number: RXISO-100)
Phosphate buffered saline (PBS) (Thermo Fisher Scientific, catalog number: 14190136)
Heparin (Patterson Veterinary, catalog number: 07-805-8262, CAS 9005-49-6)
4% paraformaldehyde in PBS (Fisher Scientific, catalog number: AAJ19943K2)
Methanol (Carolina Biological Supply, catalog number: 874195, CAS: 67-56-1)
30% Hydrogen peroxide (Sigma-Aldrich, catalog number: H1009, CAS: 7722-84-1)
Dichloromethane (DCM) (Millipore-Sigma, catalog number: 270997-2L, CAS: 75-09-2)
Benzyl ether (DBE) (Sigma-Aldrich, catalog number: 108014-1KG, CAS: 103-50-4)
Vitamin E (A17039 DL-alpha-Tocopherol, 97%+) (Alfa Aesar, catalog number: 10191-41-0, CAS: 10191-41-0)
Benzyl alcohol (Sigma-Aldrich, catalog number: W213705, CAS: 100-51-6)
Benzyl benzoate (Sigma-Aldrich, catalog number: B9550, CAS: 120-51-4)
Diphenyl ether (DPE 99%) (A15791, 101-84-8, Alfa Aesar, Lancashire, UK, CAS: 101-84-8)
Tert-butanol (Sigma-Aldrich, catalog number: 471712, CAS: 75-65-0)
(Optional) Sodium azide (Sigma-Aldrich, catalog number: S2002, CAS: 26628-22-8)
Solutions
Methanol/diH2O (see Recipes)
70% ethanol (see Recipes)
Peroxide and methanol solution (see Recipes)
Dichloromethane (DCM) 66%/33% Methanol (see Recipes)
Tert-butanol in diH2O (see Recipes)
Benzyl alcohol:benzyl benzoate (BABB) (see Recipes)
BABB-D15 (see Recipes)
1× PBS (see Recipes)
Recipes
Recipes can be scaled up to clear multiple brains in parallel. The volumes below are for a single brain.
Methanol/diH2O series
Make these fresh the day of or the day before, and allow them to reach room temperature, which can take 1.5 h.
Reagent Final concentration Volume
Methanol 20%, 40%, 60%, 80% 4 mL, 8 mL, 24 mL, 16 mL
diH2O n/a 16 mL, 12 mL, 8 mL, 4 mL
Total n/a 20 mL
70% ethanol (higher concentrations are ok, used to wipe down and remove DBE)
This can be made ahead and stored for up to a year if in a sealed container.
Reagent Final concentration Volume
Ethanol (absolute) 70% 700 mL
H2O n/a 300 mL
Total n/a 1,000 mL
Peroxide and methanol solution
This can be made ahead and stored for up to a year if in a sealed container.
Reagent Final concentration Volume
30% Hydrogen Peroxide (H2O2) 5% 4 mL
Methanol 80% 20 mL
Total n/a 24 mL
66% Dichloromethane (DCM) and 33% methanol solution
CAUTION: Use double or chemical nitrile gloves, work under hood or snorkel, expel slowly. This should be made within 24 h and not stored long term because of its volatility.
Reagent Final concentration Volume
Dichloromethane 66% 16 mL
Methanol 33% 8 mL
Total n/a 24 mL
Tert-butanol solutions
Make within 24 h and ideally 1.5 h ahead of use. Allow to reach 37 °C, which can take 1.5 h. In some labs, room temperature is too cold for tert-butanol, and it can solidify at higher concentrations. If so, bring up to 37 °C to make dilutions and for use either in an incubator or water bath. If your lab is too cold for liquid tert-butanol, also bring the brains up to 37 °C as described in the procedure.
Reagent Final concentration Volume
Tert-butanol 30%, 50%, 70%, 80%, 90%, 96% 7.2, 12, 16.8, 19.2, 21.6, 23.04 mL
diH2O n/a 16.8, 12, 7.2, 4.8, 2.4, 0.96 mL
Total n/a 24 mL
BABB (Benzyl alcohol benzyl benzoate)
This can be made ahead and stored for up to a year if in a sealed container. Other groups have successfully replaced BABB with ethyl cinnamate (Henning, 2019), which is less toxic.
Reagent Final concentration Volume
Benzyl alcohol 33.33% 6 mL
Benzyl benzoate 66.66% 12 mL
Total 100% 18 mL
BABB-D15
This can be made ahead and stored for up to a year if in a sealed container.
Reagent Final concentration Volume
BABB (see above) 93.38% 15 mL
Diphenyl ether (DPE) 6.22% 1 mL
Vitamin E 0.4% 0.064 mL
Total n/a 16.064 mL
1× PBS
This can be made ahead and stored for up to a year if in a sealed container. Recommend adding 0.1% sodium azide and/or storing at 4 °C if possible, but not required.
Reagent Final concentration Volume
10× Phosphate Buffered Saline 10% 50 mL
diH2O 90% 450 mL
Total 500 mL
Laboratory supplies
Rongeurs (Fine Science Tools, catalog number: 16022-15)
20 mL scintillation vials (Millipore Sigma, catalog number: Z190535)
3D-printed holder (we used a Form3 SLA 3D-printer and Black Resin from Formlabs, catalog number: RS-F2-GPBK-04) https://github.com/emilyjanedennis/PRA/blob/main/rat_brainholder.STL
Objective MI Plan 1.1×, NA 0.1 (Miltenyi Biotec, catalog number: 150-000-493)
Filter (Semrock, catalog number: FF01-525/39-25)
5 mL Eppendorf tubes (Genesee Scientific, catalog number: 86-899SC)
Super glue. We used Loctite Super Glue (Loctite, catalog number: 212111) but any similar product should be acceptable.
Equipment
Rocker (Genesee Scientific, model: 27-529)
Ultramicroscope II (LaVision Biotec. Bielefeld, Germany)
LifeCanvas SmartSPIM (LifeCanvas SmartSPIM, Cambridge, MA)
Fume hood/snorkel
Perfusion equipment. This should not be sensitive to brand or type. If you see a lot of capillaries in your cleared brains, this usually means that the perfusion is incomplete. We used MasterFlex L/S (NA5183975, VWR, Radnor, PA) and tubing size 16 (3.10 mm ID). Please also see Gage et al. (2012)
Access to a workstation computer and/or a computational server or cluster. There are no specific requirements, but the more powerful the computer, the faster each step will take. We recommend at least an Intel i7 Processor and 64 GB of available RAM or equivalent. More RAM, processing cores, and faster processor speeds will produce noticeable improvements
Waxholm Space Atlas (WHS) https://www.nitrc.org/projects/whs-sd-atlas from Papp et al. (2014)
Software and datasets
ImSpector Microscope controller software Version 5.1.347 (free)
Abberior Instruments Development Team, Imspector Image Acquisition & Analysis Software v16.3, http://www.imspector.de
Elastix 4.8 or 5.0 (free)
Custom software (free) http://www.github.com/emilyjanedennis/PRA Scripts are written in Python (3.0+). The server-side implementation is also written in bash (2.0+)
Procedure
Tissue preparation
There is nothing unique about this section compared to methods common in most rodent neuroscience laboratories. Please see Gage et al. (2012) for a detailed protocol with videos. There is no particular peristaltic pump required; modify the general recommendations below based on the manufacturer’s instructions for your pump.
Prepare animal and environment
Weigh the animal.
Prepare 1× PBS for flushing the system, fixative solution, as well as an anesthetic cocktail based on your animal use protocol. We used ketamine (80 mg/kg) and xylazine (10 mg/kg).
Flush your peristaltic pump with PBS. We use PBS. CAUTION: Ensure no bubbles are visible in any of the tubing. If you see any bubbles, continue to flush until all bubbles are gone. Stop the flow of PBS and ensure that you can see a protruding meniscus is at the end of the tube before transferring the intake tube to the fixative solution to avoid a bubble.
Perfuse animal (at room temperature)
Anesthetize the animal based on your animal care requirements. We used 3% isoflurane in oxygen. Once the animal is asleep, administer the anesthetic cocktail. Wait and ensure the animal is in a deep, surgical plane of anesthesia following your animal care requirements.
Make an incision under the ribcage across the animal’s abdomen. Separate the liver and diaphragm. Cut the diaphragm. Then, cut the ribs on either side. Be careful not to harm the heart or lungs; use a hemostat to hold the ribcage and slowly pull up and away from the body towards the head to expose the heart and lungs.
Insert a 15 gauge needle into the posterior left ventricle. You may snip the area with scissors if the needle does not easily puncture the ventricle. You may clamp the needle in place, either over the aorta or at the incision site.
Use scissors to snip the animal’s right atrium only.
Begin perfusing PBS through the animal. Adjust the needle if needed [see Gage et al. (2012) and troubleshooting tip 1 for details]. Run until the liver loses its color and the fluid exiting the right ventricle is clear; this should be approximately 200 mL.
Switch the perfusion solution from PBS to fixative. The animal may twitch; this is expected. After 200 mL of fixation fluid, the animal should feel stiff all over. You can make your own fixative solution, but we purchased 4% paraformaldehyde in PBS.
Extract brain
Remove the head with scissors or a guillotine.
Use scissors to trim muscles and to expose the skull by making an incision from the neck to the face.
Carefully remove the skull with rongeurs. If the skull chips into pieces, the edges can damage your sample. Take care to avoid this.
Use a spatula, a surgical blade, or small scissors to sever the nerves around the edges of the brain until it comes free.
Place brain in fixative solution in a 15–20 mL Eppendorf or scintillation vial overnight (10–24 h). Use vials made of either glass or polypropylene.
Wash brain
Wash brain with 1× PBS three times for 30 min each, rocking or shaking at room temperature.
OPTIONAL: Samples can be stored in PBS for 3 days or less at 4 °C.
Tissue processing (uDISCO, Figure 1A)
You should follow either A (uDISCO) or B (iDISCO, below) but not both. To transfer the brains to a new solution, you may reuse the same vial throughout unless otherwise noted. To do this, we recommend decanting the current solution without decanting the brain, then filling the vial with the next solution until the brain is floating and completely covered. We used glass scintillation vials, but polypropylene is also acceptable.
Dehydration
Mix tert-butanol solutions to allow them to reach room temperature (~1.5 h).
5 min rocking at room temperature in 15–20 mL Eppendorf tubes or scintillation vials in 0.1 M PBS. Use vials made of either glass or polypropylene.
Transfer brain to 30% tert-butanol, rocking at 37 °C overnight (10–24 h).
Transfer brain to 50% tert-butanol, rocking at 37 °C overnight (10–24 h).
Transfer brain to 70% tert-butanol, rocking at 37 °C overnight (10–24 h).
Transfer brain to 80% tert-butanol, rocking at 37 °C overnight (10–24 h).
Transfer brain to 90% tert-butanol, rocking at 37 °C overnight (10–24 h).
Transfer brain to 96% tert-butanol, rocking at 37 °C overnight (10–24 h).
Transfer brain to 100% tert-butanol, rocking at 37 °C overnight (10–24 h).
Clearing
Transfer brain to 100% DCM, rocking at room temperature for 2.5–3 h. CAUTION: Wear double nitrile gloves and work under a ventilation hood or snorkel.
Carefully wipe with 70% (or higher) ethanol in water to remove any solution on the exterior of the vial or tube.
Transfer brain to a fresh final vial with BABB-D15 for three or more hours. Inverting (not rocking) several times (ideally every 30 min but can be flexible between 15–45 min).
Carefully wipe with 70% (or higher) ethanol in water to remove any solution on the exterior of the vial or tube.
Cleared brains can be stored indefinitely at room temperature or 4 °C. We recommend 4 °C for long-term storage.
Tissue processing (iDISCO, Figure 1A)
You should follow either A (uDISCO, above) or B (iDISCO) but not both. To transfer the brains to a new solution, you may reuse the same vial throughout unless otherwise noted. To do this, we recommend decanting the current solution without decanting the brain, then filling the vial with the next solution until the brain is floating and completely covered. We used glass scintillation vials, but polypropylene is also acceptable.
Dehydration
Mix fresh methanol solutions to allow them to reach room temperature (~1.5 h).
Transfer brain to 20% methanol in diH2O rocking at room temperature for 3 h.
Transfer brain to fresh 20% methanol in diH2O rocking at room temperature for 3 h.
Transfer brain to 40% methanol in diH2O rocking at room temperature for 2 h.
Transfer brain to 40% methanol in diH2O rocking at room temperature overnight.
Transfer brain to 60% methanol in diH2O rocking at room temperature for 3 h.
Transfer brain to 60% methanol in diH2O rocking at room temperature for 3 h.
Transfer brain to 80% methanol in diH2O rocking at room temperature for 2 h.
Transfer brain to 80% methanol in diH2O rocking at room temperature overnight.
Transfer brain to 100% methanol rocking at room temperature for 3 h.
Transfer brain to fresh 100% methanol rocking at room temperature for 3 h. Can be stored for months in 100% methanol at 4 °C in the dark without rocking.
Bleaching
48–72 h in peroxide and methanol solution (5% H2O2 in methanol) at room temperature with rocking.
Wash brain three times in 100% methanol rocking at room temperature for 2 h each wash. Can be stored for months in 100% methanol at 4 °C in the dark without rocking.
Clearing
Transfer brain to 66% DCM/33% methanol rocking at room temperature for 6 h (or up to 8 h). CAUTION: Wear double nitrile gloves and work under a ventilation hood or snorkel when working with DCM. Wipe outside of tube or vial with 70% or greater ethanol.
Transfer brain to 100% DCM rocking at room temperature for 1 h and wipe outside of tube or vial with 70% or greater ethanol.
Transfer brain to fresh 100% DCM for 1 h.
Transfer brain to fresh 100% DBE at room temperature without rocking for 3–24 h. When possible, invert sample several times.
Transfer brain to a fresh tube or vial of 100% DBE at room temperature and wipe outside of tube or vial with 70% or greater ethanol.
Samples can be stored indefinitely in DBE at room temperature or 4 °C without rocking. We recommend 4 °C in the dark for long-term storage and ensuring minimal air in the vial.
Imaging the sample: Imaging details will depend on your microscope and experimental goals
Examples could include using higher magnification (at least 4×) for counting labeled cell centers, or lower (1.1×) for general imaging of gross anatomy to, for example, make a common coordinate system or identify a probe’s location. You will likely need to use tiles to cover the entire brain. We recommend at least 20% overlap for all tiles to assist the fusion and alignment process later. Also see Pisano et al. (2022).
Set up the microscope with appropriate filters. For autofluorescence, shorter wavelengths provide more information. We recommend 488 nm excitation, 525 nm emission (or lower wavelengths). Ensure the light-sheets are aligned according to the manufacturer’s specifications. This process will be slightly different for every microscope, so please check your manual before proceeding.
Don gloves and ensure you are not using any plastic items unless you are sure they are compatible with the chemicals in use.
Add the refractive index matched solution (BABB-D15 for uDISCO or DBE for iDISCO) to the microscope chamber according to the manufacturer’s instructions. Take care, as some microscopes will not be compatible and either solution could melt some plastics.
Clean the brain holder (custom 3D-printed or manufacturer-provided) with ethanol and let dry. Place a cleared brain onto a paper towel, then use the smallest possible amount of super glue to affix the brain to the holder. We typically super glue either the brainstem or olfactory bulbs, depending on project needs (Figure S2).
Insert the brain into the solution in the microscope.
Image the brain: turn on lasers, camera, stage, and software. We recommend using 20% overlap or greater for tiles. We also recommend imaging with the smallest axis parallel to the detection axis (the objective) and perpendicular to the light-sheets (such that the left light-sheet enters the left side of the brain, the right light-sheet enters the right side of the brain, and the objective images the z-axis from dorsal to ventral).
After acquisition, remove the brain, and replace it back in DBE or BABB-D15. Power off all items, clean thoroughly.
Some imaging platforms will automatically blend the light-sheets or stitch the tiles to create single z planes. Often, this is sufficient, but acquiring raw data before blending and stitching is recommended in case these steps are suboptimal and need to be repeated.
Create a 3D volume: there are many tools available for these steps
We provide free code to handle images from either LaVision Ultrascope II or SmartSPIM microscopes. However, the code can be altered to allow for alternative inputs.
On a cluster, run either:
spim_downsize_n_register.sh providing the required inputs: the path to the main image folder, the registration channel subfolder name, and the cell channel subfolder name. If you only have one channel imaged, just repeat the registration channel folder name.
sub_registration.sh
Align to the Princeton RAtlas
This section assumes you have already installed all the necessary software. Detailed instructions are available in the GitHub repository README.
Data analysis
We quantified the opacity of our cleared tissues in two ways: the field-standard qualitative assessment of reading text through a cleared brain (Figure 1B), and quantitatively, using FRC-QE (Figure 1D–1E) (Preusser et al., 2021). Next, inspired by the Scalable Brain Atlas approach (Bezgin et al., 2009; Sergejeva et al., 2015), we quantified the process of importing annotations into the Princeton RAtlas (PRA) space by asking four experienced humans to annotate points in anonymized volumes (Figure 3). These volumes included a cleared brain (k320) pre-alignment, after affine alignment, after affine and 1 b-spline alignment, and after affine and 3 b-spline alignment (final alignment) to the atlas. Details on these alignment types can be found in Figure S1. We also included a volume containing the Waxholm Space Atlas (WHS) to measure the accuracy of our human annotators and, unbeknownst to them, a second copy of the fully aligned k320 brain to quantify their precisions.
Figure 3. Quantification of computational alignment efficacy compared with humans. A. Slices from the Waxholm Space Atlas of the Adult Rat Brain demonstrating the location (white arrow, +) of identifiable points of interest. Abbreviations: d/v dorsal/ventral, c/r caudal/rostral, L/R left/right. B. Drawing of the location of each slice from A shown in sagittal and axial spaces. C. Sagittal (left) and axial (right) examples of four human annotators’ points for location VM1 (ventricle middle 1) in a single brain compared with imported and aligned ground truth data from the Waxholm atlas. D. Sagittal (left) and axial (right) examples of four human annotators’ points for location VM1 made in Waxholm atlas space, compared with the ground truth data from the atlas. E. Simplified diagram demonstrating how to calculate the distance metric used in F. F. Quantification of distances between user-annotated points in distinct stages of alignment and the imported annotations (WHS). G. Quantification of distances demonstrating the accuracy (left) and precision (right) of human annotators for all four annotated points.
Validation of protocol
In addition to the eight animals used to create the PRA, we discarded two brains due to damage or streaking in the imaging, and six additional successfully cleared brains were used for supplementary figures and further internal validation on various sizes of brains.
General notes and troubleshooting
Waste disposal. Please refer to your local requirements and consult with your Environmental Health and Safety or appropriate agency. In New Jersey, where these experiments were conducted, all reagents except PBS and water are considered hazardous waste and should be handled with care. Aside from PBS and water, none of these reagents should be discarded in a standard sink.
Perfusion: The quality of the perfusion can strongly impact the clearing quality. For example, if you see lots of autofluorescence that look like capillaries in your final cleared brains when imaging, this is often a sign of a bad perfusion. See Gage et al. (2012) for details on how to properly perfuse, and also the specific notes below for common problems.
If the heart stops before needle insertion, clots will begin to form. Practice until you can quickly dissect and insert the needle while the heart is still bleeding.
During perfusion, if you see bloody discharge from the nose or mouth of the animal, you have likely inserted the needle incorrectly. Check that the needle is in the left atrium but do not insert too deep as you can puncture the internal wall between the atria.
During perfusion, if anything is punctured except the left ventricle, attempt to clamp the wound.
Installing Elastix 5.0 is often the hardest step of the analysis.
On Windows, follow the instructions on “read the docs” https://simpleelastix.readthedocs.io/GettingStarted.html#compiling-on-windows starting with step 3, the command line, and skip the IDE in steps 4 and 5. If the Python wrapping step fails, see this known issue https://github.com/SuperElastix/SimpleElastix/issues/243 .
On Linux or MacOS: follow the instructions in the Elastix 5.0 manual under “the easy way” and not the “super easy way.” Ensure you have ITK 5.0.
Acknowledgments
Yisi S. Zhang for sharing unpublished protocols. Meg Younger, Ahmed El Hady, Yisi S. Zhang for helpful discussions. Annie Chen, Sara Guariglia, and Sanjeev Janarthanan for additional imaging assistance. Austin Hoag for sharing unpublished code. Adrian Bondy, Charles Kopec, Jess Breda, Marino Pagan, Thomas Luo, and Tyler Boyd-Meredith for manual annotations. ZMD, LAL, and SS-HW were funded by U19 NS104648-01 (IBW), NIH R01 MH128775 and NIH R01 NS045193. EJD was funded by Howard Hughes Medical Institute, the Helen Hay Whitney Foundation Fellowship, and Hanna H. Gray Fellowship. CDB was funded by Howard Hughes Medical Institute and U19 NS104648-01 (IBW). The research paper in which the protocol is used is Bondy et al. (2023).
Competing interests
The authors have no competing interests to declare.
Ethical considerations
Animal use procedures were approved by the Princeton University Institutional Animal Care and Use Committee (IACUC; Protocols #1837 and #1853). Rats were housed in pairs. “Male” and “female” animals were operationally defined by external genitalia. A total of 14 male and 4 female Long-Evans rats (Hilltop Lab Animals Inc., Long Evans Hla(LE)CVF), Scottdale, PA) between the ages of 3 and 12 months, weighing between 290 and 597 g, were used for this study.
References
Ahrens, M. B., Orger, M. B., Robson, D. N., Li, J. M. and Keller, P. J. (2013). Whole-brain functional imaging at cellular resolution using light-sheet microscopy. Nat. Methods 10(5): 413–420.
Bezgin, G., Reid, A. T., Schubert, D. and Kötter, R. (2009). Matching Spatial with Ontological Brain Regions using Java Tools for Visualization, Database Access, and Integrated Data Analysis. Neuroinformatics 7(1): 7–22.
Branch, A., Tward, D., Kolstad, A. C., Pulyadi, V., Vogelstein, J. T., Wu, Z. and Gallagher, M. (2019). An optimized tissue clearing protocol for rat brain labeling, imaging, and high throughput analysis. bioRxiv: e1101/639674.
Chung, K. and Deisseroth, K. (2013). CLARITY for mapping the nervous system. Nat. Methods 10(6): 508–513.
Henning, Y., Osadnik, C., and Malkemper, E. P. (2019). EyeCi: Optical clearing and imaging of immunolabeled mouse eyes using light-sheet fluorescence microscopy. Exp. Eye Res. 180:137-145.
Hillman, E. M., Voleti, V., Li, W. and Yu, H. (2019). Light-Sheet Microscopy in Neuroscience. Annu. Rev. Neurosci. 42(1): 295–313.
Keller, P. J., Schmidt, A. D., Wittbrodt, J. and Stelzer, E. H. (2008). Reconstruction of Zebrafish Early Embryonic Development by Scanned Light Sheet Microscopy. Science 322(5904): 1065–1069.
Molbay, M., Kolabas, Z. I., Todorov, M. I., Ohn, T. and Ertürk, A. (2021). A guidebook for DISCO tissue clearing. Mol. Syst. Biol. 17(3): e20209807.
Gage, G. J., Kipke, D. R., Shain, W. (2012). Whole Animal Perfusion Fixation for Rodents. J. Vis. Exp. (65): e3564.
Pan, C., Cai, R., Quacquarelli, F. P., Ghasemigharagoz, A., Lourbopoulos, A., Matryba, P., Plesnila, N., Dichgans, M., Hellal, F., Ertürk, A., et al. (2016). Shrinkage-mediated imaging of entire organs and organisms using uDISCO. Nat. Methods 13(10): 859–867.
Papp, E. A., Leergaard, T. B., Calabrese, E., Johnson, G. A. and Bjaalie, J. G. (2014). Waxholm Space atlas of the Sprague Dawley rat brain. NeuroImage 97: 374–386.
Preusser, F., dos Santos, N., Contzen, J., Stachelscheid, H., Costa, Ã. T., Mergenthaler, P. and Preibisch, S. (2021). FRC-QE: a robust and comparable 3D microscopy image quality metric for cleared organoids. Bioinformatics 37(18): 3088–3090.
Ramón y Cajal, S. (1911). Histologie du système nerveux de l'homme & des vertébrés. Maloine: e48637.
Renier, N., Wu, Z., Simon, D. J., Yang, J., Ariel, P. and Tessier-Lavigne, M. (2014). iDISCO: A Simple, Rapid Method to Immunolabel Large Tissue Samples for Volume Imaging. Cell 159(4): 896–910.
Schwarz, M. K., Scherbarth, A., Sprengel, R., Engelhardt, J., Theer, P. and Giese, G. (2015). Fluorescent-Protein Stabilization and High-Resolution Imaging of Cleared, Intact Mouse Brains. PLoS One 10(5): e0124650.
Sergejeva, M., Papp, E. A., Bakker, R., Gaudnek, M. A., Okamura-Oho, Y., Boline, J., Bjaalie, J. G. and Hess, A. (2015). Anatomical landmarks for registration of experimental image data to volumetric rodent brain atlasing templates. J. Neurosci. Methods 240: 161–169.
Siedentopf, H. and Zsigmondy, R. (1902). Uber Sichtbarmachung und Größenbestimmung ultramikoskopischer Teilchen, mit besonderer Anwendung auf Goldrubingläser. Ann. Phys. 315(1): 1–39.
Stefaniuk, M., Gualda, E. J., Pawlowska, M., Legutko, D., Matryba, P., Koza, P., Konopka, W., Owczarek, D., Wawrzyniak, M., Loza-Alvarez, P., et al. (2016). Light-sheet microscopy imaging of a whole cleared rat brain with Thy1-GFP transgene. Sci. Rep. 6(1): e1038/srep28209.
Susaki, E. A., Tainaka, K., Perrin, D., Yukinaga, H., Kuno, A. and Ueda, H. R. (2015). Advanced CUBIC protocols for whole-brain and whole-body clearing and imaging. Nat. Protoc. 10(11): 1709–1727.
Woo, J., Lee, E. Y., Park, H. S., Park, J. Y. and Cho, Y. E. (2018). Novel Passive Clearing Methods for the Rapid Production of Optical Transparency in Whole CNS Tissue. J. Vis. Exp.: e3791/57123-v.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Demonstration of the selection process for parameter files
Figure S2. Example diagram of glued brains
Article Information
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© 2023 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Cell Biology > Tissue analysis > Tissue imaging
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4,855 | https://bio-protocol.org/en/bpdetail?id=4855&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Maize Seedlings Colonization with Serendipita indica and Its Colonization Efficiency Analysis
Om Prakash Narayan
BY Bindu Yadav
NV Nidhi Verma
MD Meenakshi Dua
AJ Atul Kumar Johri
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4855 Views: 666
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Original Research Article:
The authors used this protocol in The Plant Cell Apr 2021
Abstract
Maize is one of the most important crops in the world, and ensuring its successful growth and productivity is crucial for global food security. One way to enhance maize growth and productivity is by improving the colonization of its roots by beneficial microorganisms. In this regard, Serendipita indica, a plant growth–promoting fungus, has gained attention for its ability to enhance plant growth and productivity, especially in cereal crops and medicinal plants. Previous studies have shown that S. indica can colonize various plant species, including maize, but the efficiency of the colonization process in maize seedlings has not been extensively characterized. This protocol outlines a method for efficient colonization of maize seedlings with the beneficial fungus S. indica. The protocol includes the preparation of stock solutions, maintenance and growth of S. indica, surface sterilization and germination of seeds, preparation of S. indica chlamydospores, and colonization of maize plants with S. indica. The advantages of this protocol include the use of surface sterilization techniques that minimize contamination, the production of a large number of viable chlamydospores, and efficient colonization of maize seedlings with S. indica. This protocol may be useful for researchers studying the role of S. indica in promoting plant growth and combating biotic and abiotic stress. Additionally, this protocol may be used in the development of biofertilizers using S. indica as a means of increasing crop yields and reducing dependence on synthetic fertilizers. Overall, this protocol offers a reliable and efficient method for colonizing maize seedlings with S. indica and may have potential applications in the agricultural industry. This study also provides a valuable tool for researchers interested in studying plant–microbe interactions in maize and highlights the potential of S. indica as a biocontrol agent to enhance maize productivity under adverse conditions.
Key features
• This protocol builds upon the method developed by Narayan et al. (2022), and its application optimized for the root endophytic symbiotic fungus S. indica.
• This protocol also allows for histochemical analysis to visualize the colonized fungal spores in the root cells of host plant species.
• This protocol helps in mathematical calculation of the percent colonization or efficiency of colonization.
• This protocol utilizes readily available laboratory equipment, including a light microscope, autoclave, and laminar flow hood, ensuring ease of reproducibility in other research laboratories.
Graphical overview
Keywords: Maize Serendipita indica Colonization Plant microbe interaction Endophytic fungus Plant biotic stress Symbiosis PGPF
Background
Maize is an important cereal crop and an essential source of food, feed, and biofuel worldwide. However, maize is highly susceptible to biotic and abiotic stresses, such as pests, diseases, and adverse environmental conditions, which can significantly reduce crop yield and quality (Erenstein et al., 2022; Chávez-Arias et al., 2021). Therefore, it is becoming increasingly important to create sustainable and eco-friendly agricultural methods that can improve the productivity and resilience of maize crops. One such strategy is the use of plant growth–promoting fungi (PGPF) to improve plant growth and health (Adedayo and Babalola, 2023).
S. indica (formerly known as Piriformospora indica) is a PGPF that has been reported to enhance the growth and stress tolerance of various plant species, including maize (Singhal et al., 2017; Narayan et al., 2021). The colonization of maize seedlings with S. indica has been shown to increase root growth, nutrient uptake, photosynthetic efficiency, and resistance to various biotic and abiotic stresses (Varma et al., 1999; Narayan et al., 2017, 2021 and 2022; Prasad et al., 2019; Verma et al., 2022). Therefore, the optimization and standardization of the maize seedlings’ colonization protocol with S. indica are crucial to ensure the reproducibility and reliability of the results.
The protocol described in this paper can help explore the mechanisms underlying the beneficial effects of S. indica on maize growth and stress tolerance and identify potential molecular and biochemical markers of plant–fungi symbiosis. Moreover, this protocol can be used to evaluate the performance of different S. indica strains and inoculation methods and to assess the effects of environmental factors on colonization efficiency, and plant growth and development.
Several methodologies have been developed to study the interactions between plant growth–promoting fungi (PGPF) and plants such as wheat (Triticum aestivum), rice (Oryza sativa), barley (Hordeum vulgare), bean (Phaseolus vulgaris), soybean (Glycine max), and chickpea (Cicer arietinum) using S. indica (Rocha et al., 2019; Wahid et al., 2019; El-Maraghy et al., 2020; Mahdi et al., 2022; Verma et al., 2022). One common approach is the use of in vitro culture systems, such as agar dishes or hydroponic cultures, to assess the effects of S. indica on plant growth and stress responses under controlled conditions (Osman et al., 2020). Another approach is the use of field trials to evaluate the performance of S. indica in enhancing crop productivity and quality under natural conditions. Additionally, various molecular and biochemical techniques, such as quantitative polymerase chain reaction (qPCR), metabolomics, and proteomics, were used to analyze the gene and protein expression levels and metabolic pathways involved in the plant–fungi interaction (Gouda et al., 2018).
The protocol described in this paper has several advantages over other published methods (Kumar et al., 2009; Hosseini et al., 2018; Zhang et al., 2018). Firstly, this protocol provides a standardized and reproducible procedure for the colonization of maize seedlings with S. indica. Secondly, this protocol allows for the evaluation of the colonization efficiency and distribution of S. indica in different plant tissues, which provides insights into the spatial and temporal dynamics of the plant–fungi interaction. Thirdly, this protocol can be combined with various physiological, biochemical, and molecular analyses to study the mechanisms underlying the plant–fungi symbiosis.
In addition to the research applications, this protocol has several other possible applications. Firstly, this protocol can be used to develop inoculum production and delivery methods for S. indica, which can be used for commercial purposes, such as biofertilizer or biostimulant production. Secondly, this protocol can be used to screen and identify potential S. indica strains with improved plant growth–promoting and stress tolerance properties, which can be used to develop new and more effective PGPF-based products. Thirdly, this protocol can be used to investigate the interaction between S. indica and other beneficial microbes, such as mycorrhizal fungi or rhizobia, and their combined effects on plant growth and health. Overall, this protocol has the potential to advance research in the field of plant–microbe interactions and to contribute to the development of sustainable and environmentally friendly agricultural practices.
Materials and reagents
Biological materials
Maize seeds (Variety HQPM-5 from Indian Agriculture Research Institute, Pusa, New Delhi, India)
Serendipita indica fungus (strain DSM11827 gifted from Prof. Ajit Verma) (Varma, Kost, Rexer & Franken, 1997, European patent office, Muenchen, Germany. Patent No. 97121440.8-2105, Nov 1998)
Reagents
Potassium hydroxide (KOH) (Sigma-Aldrich, catalog number: 1310-58-3)
Hydrochloric acid (HCl) (Sigma-Aldrich, catalog number: 7647-01-0)
Phenol (Sigma-Aldrich, catalog number: 108-95-2)
Lactic acid (Sigma-Aldrich, catalog number: 50-21-5)
Glycerol (Sigma-Aldrich, catalog number: 56-81-5)
Trypan blue (Sigma-Aldrich, catalog number: 72-57-1)
Glucose (Sigma-Aldrich, catalog number: 50-99-7)
Calcium chloride (CaCl2) (Sigma-Aldrich, catalog number: 10043-52-4)
Ferrous chloride (FeCl2) (Sigma-Aldrich, catalog number: 7705-08-0)
Calcium nitrate [Ca(NO3)2] (Sigma-Aldrich, catalog number: 13477-34-4)
Magnesium sulfate (MgSO4.7H2O) (Sigma-Aldrich, catalog number: 7487-88-9)
Potassium nitrate (KNO3) (Sigma-Aldrich, catalog number: 7757-79-1)
Potassium phosphate monobasic (KH2PO4) (Sigma-Aldrich, catalog number: 7778-77-0)
Boric acid (H3BO3) (Sigma-Aldrich, catalog number: 10043-35-3)
Manganese sulfate monohydrate (MnSO4.H2O) (Sigma-Aldrich, catalog number: 10034-96-5)
Copper sulfate (CuSO4.5H2O) (Sigma-Aldrich, catalog number: 7758-98-7)
Zinc sulfate (ZnSO4.7H2O) (Sigma-Aldrich, catalog number: 7733-02-0)
Ammonium molybdate [(NH4)2MoO4] (Sigma-Aldrich, catalog number: 13106-76-8)
Sodium-hypochlorite (NaClO) (Sigma-Aldrich, catalog number: 7681-52-9)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: 7447-40-7)
Manganese(II) chloride (MnCl2) (Sigma-Aldrich, catalog number: 7773-01-5)
Iron(II) sulfate heptahydrate (FeSO4.7H2O) (Sigma-Aldrich, catalog number: 7782-63-0)
Cobalt(II) chloride (CoCl2) (Sigma-Aldrich, catalog number: 7646-79-9)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: 7647-14-5)
Sucrose (Sigma-Aldrich, catalog number: 57-50-1)
Ethylenediaminetetraacetic acid ferric sodium salt (NaFeEDTA) (Sigma-Aldrich, catalog number: 15708-41-5)
Potassium iodide (KI) (Sigma-Aldrich, catalog number: 7681-11-0)
Manganese(II) chloride (MnCl2) (Sigma-Aldrich, catalog number: 7773-01-5)
Sodium molybdate dihydrate (Na2MoO4.2H2O) (Sigma-Aldrich, catalog number:10102-40-6)
Glycine (Sigma-Aldrich, catalog number: 56-40-6)
Ammonium phosphate dibasic [(NH4)2HPO4] (Sigma-Aldrich, catalog number: 7783-28-0)
Iron(III) chloride (FeCl3) (Sigma-Aldrich, catalog number: 7705-08-0)
Ethanol (Sigma-Aldrich, catalog number: 64-17-5)
Peptone (Sigma-Aldrich, catalog number: 73049-73-7)
Yeast extract (Sigma-Aldrich, catalog number: 8013-01-2)
Casamino acid (Sigma-Aldrich, catalog number: 65072-00-6)
Agar (Sigma-Aldrich, catalog number: 9002-18-0)
BD DifcoTM BactoTM Agar (Sigma-Aldrich, catalog number: DF0140-15-4)
Biotin (Sigma-Aldrich, catalog number: 7646-79-9)
Nicotinamide (Sigma-Aldrich, catalog number: 98-92-0)
Pyridoxal phosphate (Sigma-Aldrich, catalog number: 853645-22-4)
Aminobenzoic acid (Sigma-Aldrich, catalog number: 150-13-0)
Riboflavin (Sigma-Aldrich, catalog number: 83-88-5)
Thiamine hydrochloride (Sigma-Aldrich, catalog number: 67-03-8)
Pyridoxine hydrochloride (Sigma-Aldrich, catalog number: 58-56-0)
Nicotinic acid (Sigma-Aldrich, catalog number: 59-67-6)
Trypticase peptone (Sigma-Aldrich, catalog number: 91079-40-2)
Malt extract (Sigma-Aldrich, catalog number: 8002-48-0)
Myo-Inositol (Sigma-Aldrich, catalog number: 87-89-8)
Solutions
Sterile water (1 L)
Double-distilled water (DI water) (1 L)
Half-strength modified Hoagland solution (1 L) (see Recipes)
Ethanol (1 L) 70% (w/v) (see Recipes)
Sodium hypochlorite (NaClO) solution (1 L) 0.75% (w/v) (see Recipes)
0.8% Bacto agar (500 mL) (see Recipes)
Lactophenol (50%) (see Recipes)
Trypan blue (100 mL) (see Recipes)
10% KOH (100 mL) (see Recipes)
Aspergillus modified medium (AMM) or Kaefer (KF) medium (1 L) (see Recipes)
MN-Agar medium (1L) (see Recipes)
Recipes
Half-strength modified Hoagland solution (1 L)
Reagent Final concentration Quantity
Macronutrients
Ca(NO3)2·4H2O 4 mM 944.5 mg
MgSO4·7H2O 2 mM 492.9 mg
KNO3 6 mM 606.6 mg
KH2PO4 1 mM 136.08 mg
Micronutrients
H3BO3 45 μM 2.78 mg
MnSO4·4H2O 20 μM 3.01 mg
CuSO4·5H2O 0.4 μM 99.87 mg
ZnSO4·7H2O 0.7 μM 201.32 mg
(NH4)2MoO4·2H2O 0.2 μM 41.18 mg
H2O 1,000 mL
Total 1,000 mL
70% ethanol (1 L)
Reagent Quantity
Ethanol (absolute) 700 mL
H2O 300 mL
Total 1,000 mL
0.75% sodium-hypochlorite (NaClO) (bleach) solution (100 mL)
Reagent Quantity
Sodium hypochlorite solution (6%) 12.5 mL
ddH2O 87.5 mL
Total 100 mL
0.8% Bacto agar (500 mL)
Reagent Quantity
Bacto agar 4g
ddH2O 500 mL
Total 500 mL
Lactophenol (50%)
Reagent Quantity
Phenol 150 mL
ddH2O 150 mL
Lactic acid 125 mL
Glycerol 125 mL
Total 600 mL
Trypan blue (100 mL)
Reagent Quantity
Trypan blue 0.1 g
Lactophenol 100 mL
Total 100 mL
10% KOH (100 mL)
Reagent Quantity
KOH 10 g
ddH2O 100 mL
Total 100 mL
Aspergillus modified medium/Kaefer medium (1 L)
Reagent Quantity
Glucose 20 g
Peptone 2 g
Yeast Extract 1 g
Casamino acid 1 g
Vitamin stock solution 1 mL
Macro-element stock solution 50 mL
Micro-element stock solution 2.5 mL
CaCl2 (0.1 M) 1 mL
FeCl2 (0.1 M) 1 mL
Agar 10 g
pH (Adjust with 1 N HCl) 6.5
ddH2O 944.5 mL
Total 1,000 mL
Macro-elements stock
KCl 10.4 g
MgSO4·7H2O 10.4 g
KH2PO4 30.4 g
ddH2O 1,000 mL
Total 1,000 mL
Micro-elements stock
ZnSO4·7H2O 22 g
H3BO3 11 g
MnCl2·4H2O 5 g
FeSO4·7H2O 5 g
CoCl2·6H2O 1.6 g
CuSO4·5H2O 1.6 g
ddH2O 1,000 mL
Total 1,000 mL
Vitamin stock
Biotin 5 mg
Nicotinamide 50 mg
Pyridoxal Phosphate 10 mg
Aminobenzoic Acid 10 mg
Riboflavin 25 mg
ddH2O 100 mL
Total 100 mL
0.1 M FeCl2
FeCl2 1.62 g
ddH2O 100 mL
Total 100 mL
0.1 M CaCl2
CaCl2 1.11 g
ddH2O 100 mL
Total 100 mL
MN-Agar medium (1 L)
Reagent Quantity
NaCl 23.4 mg
KNO3 80 mg
Ca(NO3)2·4H2O 288 mg
Sucrose 10 g
NaFeEDTA 8 mg
KI 0.8 mg
MnCl2·4H2O 6 mg
Na2MoO4·2H2O 0.0024 mg
Glycine 3 mg
KH2PO4 272.2 mg
(NH4)2HPO4 40.8 mg
CaCl2 81.6 mg
MgSO4·7H2O 731 mg
FeCl3 583.9 mg
Thiamine hydrochloride 67.4 mg
Pyridoxine hydrochloride 67.4 mg
Nicotinic acid 0.5 mg
Trypticase peptone 1 g
Glucose 10 g
Malt extract 50 g
Myo-Inositol 50 mg
Bacto agar 10 g
KCl 65 mg
H3BO3 1.5 mg
MnSO4·H2O 6.0 mg
ZnSO4·7H2O 2.7 mg
CuSO4·5H2O 0.2 mg
pH 5.8
Total 1,000 mL
Notes:
All the stock solutions are stored at 4 °C and the vitamin stock at -20 °C. The stock of FeSO4·7H2O is prepared separately.
Filter sterilize the vitamin stocks.
Laboratory supplies
Gloves (Genesee Scientific, catalog number: 44-100M)
Lab coat (Fisher Scientific, catalog number: 19-181-570)
Pipettes of different sizes (Fisher Scientific, catalog number: 13-690-029)
Tip boxes (Fisher Scientific, catalog number: 01-670-713)
1 L Glass bottles (Fisher Scientific, catalog number: 13951L)
100 mL Glass bottles (Fisher Scientific, catalog number: 06-414-1A)
Petri dishes 20mm (Fisher Scientific, catalog number: 08-757-099)
Petri dishes 150mm (Fisher Scientific, catalog number: 50-403-868)
Plastic trays (Fisher Scientific, catalog number: 11-394-455)
Blotting papers (Fisher Scientific, catalog number: 09-301-199)
Germination paper (Fisher Scientific, catalog number: NC1466201)
Spreader (Fisher Scientific, catalog number: 14-665-230)
Beaker (Fisher Scientific, catalog number: 07-250-056)
250 mL flask (Fisher Scientific, catalog number: 10-040F)
Surgical blade (Fisher Scientific, catalog number: 22-079-774)
Muslin cloth ( Amazon.in, item model number: HAZC017639)
Nail paint (Amazon.in, item model number: CC4407)
Equipment
Laminar flow hood (Thermo ScientificTM, catalog number: 1323TS)
Light microscope (Leica Microscope, Type 020-518.500, Germany and Nikon Eclipse Ti)
Autoclave (Thymol autoclave, India, product code: TAI-902)
Incubator shaker (Multitron Incubator Shaker, HT-Infors, Switzerland)
Glass house (School of Life Sciences, Jawaharlal Nehru University, New Delhi India)
Software and datasets
Microsoft Office Excel 10
GraphPad Prism 8
Procedure
Preparation of stock solutions
To ensure the successful execution of all procedures outlined in this protocol and the proper maintenance of the S. indica culture, it is necessary to prepare the listed stock solutions mentioned above, and then prepare working solutions and mixtures in sterile water and store in glass bottles at room temperature, as required during the different steps.
Sterilize the KF medium using the autoclave at 121.6 °C under 15 psi for 20 min before plating.
Maintenance and growth of fungal species (S. indica)
Prepare 25 mL of KF-agar medium in a 100 mL glass bottle and autoclave it.
Add the vitamin stock into this KF-agar medium at a temperature of 40 °C (kept in a water bath) and mix gently to avoid bubble formation.
Pour this KF-agar medium into a 90 mm Petri dish in a laminar flow hood.
Allow 20–30 min to complete the solidification of the KF media in the Petri dish inside the laminar flow hood. Note: To improve solidification and prevent water droplet accumulation inside the lids, it is recommended to keep the Petri dish lid open.
Inoculate a single ball/colony of S. indica from the liquid KF medium (the sample has already been incubated in a shaker incubator at 30 ± 2 °C for 7–9 days with shaking at 100 rpm) or cut a slice sized 5 × 5 mm of pregrown S. indica on a solid dish culture using a sterilized surgical blade and place this slice of KF-agar-containing S. indica culture on the center of the 90 mm fresh KF medium Petri dish.
Note: A fine pressure can be applied onto the slice to make sure the S. indica slice can make full contact with the fresh KF medium.
Incubate the dishes at 28–30 °C incubator in darkness for 5–7 days for full growth.
After incubation, observe the growth of the S. indica by visualizing the grey-colored, cotton-like growth (see the right panel of Figure 1).
Figure 1. Schematics showing how to inoculate the S. indica on a solid KF-agar 90 mm Petri dish
Now, S. indica is ready for the preparation of chlamydospores and further colonization study with maize seedlings.
Note: For liquid broth medium growth, S. indica was maintained routinely on solidified Aspergillus (Aspergillus niger) modified medium (Hill and Kafer, 2001). Growth of S. indica was studied in 250 mL culture flasks with constant shaking at 100 rpm, and at 30 ± 2 °C for 7–9 days in a metabolic shaker (Multitron Incubator Shaker, HT-Infors, Switzerland).
Maize seed surface sterilization and germination
Take the required quantity of desired functional maize seeds (we used the HQPM5 variety from IARI, New Delhi, India).
Take approximately 120 seeds (in our case) and dip them into 100 mL of autoclaved DD water for 1 h using a 250 mL beaker.
Wash the maize seeds with detergent (teepol or Triton X-100) using 3–5 drops in 50 mL of water.
Repeat washing of the maize seeds with autoclaved DD water by changing the water approximately 5 times for a total of 10 min.
Sterilize the maize seeds by soaking them in a 250 mL beaker containing 70% ethanol for 40 s, manually swirling it.
Remove the ethanol and wash five times with autoclaved DD water for a total of 10 min.
Again, sterilize the maize seeds by soaking them in a 100 mL beaker containing 50 mL of 6% NaOCl (final concentration 0.75%) for 1 min, manually swirling it.
Wash the maize seeds with autoclaved DD water six times for 10 min.
Note: In this final washing, make sure there is no remaining NaOCl.
Transfer the maize seeds into a new beaker and heat it in a water bath at 60 °C for 5 min.
Place the maize seeds in water agar dishes/germination sheets using sterile forceps, spacing them 1 inch apart for germination at 28 °C in the dark for 3 days (10–15 seeds per dish).
Note: Use bi size (150 mm) glass or plastic Petri dishes for making 0.8% water agar dishes (0.8% Bacto Agar, Difco, Detroit, MI, USA).
Preparation of chlamydospores of S. indica
Take the fully grown S. indica KF dishes (6–10 days old) to prepare the chlamydospores.
Note: Use the completely grown S. indica on a KF medium and open the dish in laminar air flow.
Take approximately 2–3 mL of 2% filter-sterilized glucose solution prepared in DD water and pour on the top surface of the fully grown S. indica KF dishes.
Spread the 2% glucose solution gently using a sterile plastic spreader and aim to just collect the chlamydospores in the solution phase.
Note: Try not to disturb the bottom part of the S. indica-containing hyphae.
Collect the solution containing chlamydospores in 2 mL Eppendorf tubes.
Filter this solution using a sterile muslin cloth to remove the fungal hyphae and any part of solid medium, transferring the chlamydospores to another tube.
Centrifuge the S. indica spores (in Eppendorf tubes) at 3× g at 30 °C for 3–4 h.
The quality of the spores can be checked by observation under a light microscope using glass slides.
The number of S. indica chlamydospores can be maintained at 5 × 105/mL using a hemocytometer for further colonization with maize seedlings.
Colonization of S. indica with maize plants
Take out the germinated maize seedlings from the water agar dishes or germination sheets (3–4 days old) and wash the roots with sterilized water, then dip the roots in S. indica chlamydospore suspensions (maintained at 5 × 105 spores per mL) to start the inoculation.
Note: Forceps can be used to dip the roots into the S. indica chlamydospore suspensions.
Transfer the inoculated maize seedlings with the chlamydospores of S. indica to the sterile MN-agar medium.
Grow the inoculated maize seedlings in the MN medium for a minimum of 7 days for colonization.
Alternatively, grow the inoculated maize seedlings in pots filled with a mixture of sterile sand and soil in the ratio of 3:1.
Grow the inoculated plants under controlled conditions in a glasshouse with an 8 h light (1,000 Lux)/16 h dark period at a temperature of 28 °C with a relative humidity of 60%–70%.
Supplement the plants weekly with half-strength modified Hoagland solution.
For control plants (without S. indica inoculated), mock inoculate with autoclaved DD water–mixed sand. Check the S. indica colonization at different time points under the light microscope. In our case, plant roots were harvested at different dpi (5, 10, 15, and 20 days) and were assessed for colonization by trypan blue staining as shown below.
Histochemical analysis
Harvest the plant roots at 5, 10, 15, and 20 days after inoculation, and choose approximately 10–20 random samples of the root system for colonization estimation.
Note: Use small surgical scissors to cut the root sample from the maize plants.
To soften the root samples, treat them with 10% KOH solution for 15 min and then acidify them with 1 N HCl for 10 min. Finally, stain them with 0.02% trypan blue overnight (Narayan et al., 2021).
Samples were destained with 50% Lactophenol for 1–2 h. Keep the samples on a rotary shaker.
Cut the root samples into 1 cm small fragments using the surgical blade and mount them on glass slides.
Note: Label the glass slides with sample number and root number (see Figure 2).
Figure 2. Schematics showing a glass slide mounted with small fragments of the S. indica-colonized maize roots after the trypan blue staining, for microscopic study
Cover the root samples with the glass coverslips, pressing the coverslips gently to crush the roots, and seal slides using nail paint (see Figure 2)
Observe these slides under a light microscope to see the blue-stained and pear-shaped chlamydospores of S. indica under the different magnifications. Please see Figure 3 to see how chlamydospores of S. indica look like in a colonized state inside the root cortical cells, under light microscope observation.
Figure 3. Blue-colored chlamydospores of S. indica inside the maize root cortical cells, viewed under a light microscope.
Count the colonized root segments for further calculation.
The distribution of chlamydospores within the root was taken as an index for studying colonization. Percent colonization was calculated for the inoculated plants according to the method described previously (Narayan et al., 2017 and 2021). Specifically, it was calculated using the following formula. The calculation involved the total number of randomly selected roots, the number of colonized root segments observed under a microscope, and the total number of root segments taken from a random sample to observe chlamydospores under a microscope. Both biological and technical repeats were combined to calculate the standard deviation and determine the significance of the data. Table 1 provides an example of percent colonization.
Table 1. Percent colonization of S. Indica with maize plant root
Time after inoculation (days) Percent colonization (%)
5 7.5 ± 5
10 27.5 ± 9.6
15 45 ± 12.9
20 70 ± 11.5
Data analysis
Data can be analyzed using Excel. In our case, we used three biological and three technical repeats. Each time we used 120 seeds of maize. Statistical tests like standard deviation and standard error analysis can be done by using sigma plot or GraphPad Prism 8.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Kundu et al. (2022). Piriformospora indica recruits host-derived putrescine for growth promotion in plants. Plant Physiology. (Figure 6, panel h).
Narayan et al. (2021). Sulfur transfer from the endophytic fungus Serendipita indica improves maize growth and requires the sulfate transporter SiSulT. Plant Cell. (Figure 1, panel c).
Verma et al. (2022). Functional characterization of a high‐affinity iron transporter ( PiFTR ) from the endophytic fungus Piriformospora indica and its role in plant growth and development. Environmental Microbiology. (Figure 6, panel c).
Narayan et al. (2017). Antioxidant enzymes in chickpea colonized by Piriformospora indica participate in defense against the pathogen Botrytis cinerea. Scientific Reports. (Figure 1).
Jogawat et al. (2013). Piriformospora indica rescues growth diminution of rice seedlings during high salt stress. Plant signaling & behavior. (Figure 1, panel b).
Kumar et al. (2009). Antioxidant enzyme activities in maize plants colonized with Piriformospora indica. Microbiology. (Figure 1).
General notes and troubleshooting
General notes
The colonization efficiency of S. indica may vary depending on the maize cultivar, growth conditions, and chlamydospore quality.
Overstaining can hide the chlamydospores with the root tissues.
To improve colonization efficiency, use a fresh culture of S. indica.
Limitations associated with this protocol:
The colonization efficiency of S. indica may vary depending on the maize cultivar, growth conditions, and inoculation methods, which may affect the reproducibility of the results.
The use of high-throughput molecular techniques, such as RNA sequencing or metabolomics, may require a large amount of plant tissue, which may limit the application of this protocol to small-scale experiments.
This protocol only focuses on the early stages of maize growth, and the long-term effects of S. indica on plant growth and stress tolerance are still unclear.
Troubleshooting (Table 2)
Table 2. Troubleshooting
Common problems Troubleshooting
Low colonization efficiency Check the viability of the chlamydospores by growing on a liquid broth or solid KF-agar dishes and compare with normal S. indica growth.
Low germination efficiency Check the quality of seeds. Avoid using very old, dormant seeds.
Chlamydospores are not visible in the microscope Stain chlamydospores with trypan blue for a longer time.
Blue-colored chlamydospores are hidden in root tissues Try to destain the roots for a longer time using Lactophenol.
Contamination of other fungal species Ensure all equipment and reagents are in sterile conditions.
Acknowledgments
Jawaharlal Nehru University, New Delhi, and University Grants Commission, New Delhi India, ICMR, New Delhi, India is highly acknowledged for the financial support. We also acknowledge the original research paper in which this protocol was described and validated was published by Narayan et al., 2021, The Plant Cell (doi: 10.1093/plcell/koab006) from School of Life Sciences, Jawaharlal Nehru University, New Delhi, India.
Competing interests
All the authors declare no conflict of interests.
References
Adedayo, A. A. and Babalola, O. O. (2023). Fungi That Promote Plant Growth in the Rhizosphere Boost Crop Growth. J. Fungi 9(2): 239.
Chávez-Arias, C. C., Ligarreto-Moreno, G. A., Ramírez-Godoy, A. and Restrepo-Díaz, H. (2021). Maize Responses Challenged by Drought, Elevated Daytime Temperature and Arthropod Herbivory Stresses: A Physiological, Biochemical and Molecular View. Front. Plant Sci. 12: e702841.
El-Maraghy, S. S., Tohamy, T. A. and Hussein, K. A. (2020). Role of plant-growth promoting fungi (PGPF) in defensive genes expression of Triticum aestivum against wilt disease. Rhizosphere 15: 100223.
Erenstein, O., Jaleta, M., Sonder, K., Mottaleb, K. and Prasanna, B. (2022). Global maize production, consumption and trade: trends and R&D implications. Food Secur. 14(5): 1295–1319.
Gouda, S., Kerry, R. G., Das, G., Paramithiotis, S., Shin, H. S. and Patra, J. K. (2018). Revitalization of plant growth promoting rhizobacteria for sustainable development in agriculture. Microbiol. Res. 206: 131–140.
Hill, T. W. and Kafer, E. (2001). Improved protocols for Aspergillus minimal medium: trace element and minimal medium salt stock solutions. Fungal Genet Rep. 48(1): 20–21.
Hosseini, F., Mosaddeghi, M. R., Dexter, A. R. and Sepehri, M. (2018). Maize water status and physiological traits as affected by root endophytic fungus Piriformospora indica under combined drought and mechanical stresses. Planta 247(5): 1229–1245.
Kumar, M., Yadav, V., Tuteja, N. and Johri, A. K. (2009). Antioxidant enzyme activities in maize plants colonized with Piriformospora indica. Microbiology 155(3): 780–790.
Mahdi, L. K., Miyauchi, S., Uhlmann, C., Garrido-Oter, R., Langen, G., Wawra, S., Niu, Y., Guan, R., Robertson-Albertyn, S., Bulgarelli, D., et al. (2022). The fungal root endophyte Serendipita vermifera displays inter-kingdom synergistic beneficial effects with the microbiota in Arabidopsis thaliana and barley. ISME J. 16(3): 876–889.
Narayan, O. P., Kumar, P., Yadav, B., Dua, M. and Johri, A. K. (2022). Sulfur nutrition and its role in plant growth and development. Plant Signal. Behav.: e2030082.
Narayan, O. P., Verma, N., Jogawat, A., Dua, M. and Johri, A. K. (2021). Sulfur transfer from the endophytic fungus Serendipita indica improves maize growth and requires the sulfate transporter SiSulT. Plant Cell 33(4): 1268–1285.
Narayan, O. P., Verma, N., Singh, A. K., Oelmüller, R., Kumar, M., Prasad, D., Kapoor, R., Dua, M. and Johri, A. K. (2017). Antioxidant enzymes in chickpea colonized by Piriformospora indica participate in defense against the pathogen Botrytis cinerea. Sci. Rep. 7(1): 13553.
Osman, M., Stigloher, C., Mueller, M. J. and Waller, F. (2020). An improved growth medium for enhanced inoculum production of the plant growth-promoting fungus Serendipita indica. Plant Methods 16(1): 1–7.
Prasad, D., Verma, N., Bakshi, M., Narayan, O. P., Singh, A. K., Dua, M. and Johri, A. K. (2019). Functional Characterization of a Magnesium Transporter of Root Endophytic Fungus Piriformospora indica. Front. Microbiol. 9: e03231.
Rocha, I., Duarte, I., Ma, Y., Souza-Alonso, P., Látr, A., Vosátka, M., Freitas, H. and Oliveira, R. S. (2019). Seed Coating with Arbuscular Mycorrhizal Fungi for Improved Field Production of Chickpea. Agronomy 9(8): 471.
Singhal, U., Prasad, R. and Varma, A. (2017). Piriformospora indica (Serendipita indica): The Novel Symbiont. Mycorrhiza - Function, Diversity, State of the Art: 349–364.
Varma, A., Verma, S., Sudha, Sahay, N., Bütehorn, B. and Franken, P. (1999). Piriformospora indica, a Cultivable Plant-Growth-Promoting Root Endophyte. Appl. Environ. Microbiol. 65(6): 2741–2744.
Verma, N., Narayan, O. P., Prasad, D., Jogawat, A., Panwar, S. L., Dua, M. and Johri, A. K. (2022). Functional characterization of a high‐affinity iron transporter ( PiFTR ) from the endophytic fungus Piriformospora indica and its role in plant growth and development. Environ. Microbiol. 24(2): 689–706.
Wahid, F., Sharif, M., Fahad, S., Adnan, M., Khan, I. A., Aksoy, E., Ali, A., Sultan, T., Alam, M., Saeed, M., et al. (2019). Arbuscular mycorrhizal fungi improve the growth and phosphorus uptake of mung bean plants fertilized with composted rock phosphate fed dung in alkaline soil environment. J. Plant Nutr. 42(15): 1760–1769.
Zhang, W., Wang, J., Xu, L., Wang, A., Huang, L., Du, H., Qiu, L. and Oelmüller, R. (2018). Drought stress responses in maize are diminished byPiriformospora indica. Plant Signal. Behav. 13(1): e1414121.
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Microscopy and Plate Reader–based Methods for Monitoring the Interaction of Platelets and Tumor Cells in vitro
VT Veeresh Toragall *
EH Elizabeth J. Hale *
KH Kenneth R. Hulugalla
TW Thomas A. Werfel
(*contributed equally to this work)
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4856 Views: 916
Reviewed by: Dipak Kumar PoriaJibin SadasivanEduardo Listik
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Abstract
Platelets and their activation status play an essential role in cancer metastasis. Therefore, the anti-metastatic potential of antiplatelet drugs has been investigated for many years. However, the initial screening of these antiplatelet drugs to determine which agents can inhibit the interactions of platelets and tumor cells is very limited due to reliance upon expensive, time-consuming, and low-throughput animal experiments for screening. In vitro models of the platelet–tumor cell interaction can be a useful tool to rapidly screen multiple antiplatelet drugs and compare their ability to disrupt platelet–tumor cell interactions, while also identifying optimal concentrations to move forward for in vivo validation. Hence, we adopted methods used in platelet activation research to isolate and label platelets before mixing them with tumor cells (MDA-MB-231-RFP cells) in vitro in a static co-culture model. Platelets were isolated from other blood components by centrifugation, followed by fluorescent labeling using the dye CMFDA (CellTrackerTM Green). Labeling platelets allows microscopic observation of the introduced platelets with tumor cells grown in cell culture dishes. These methods have facilitated the study of platelet–tumor cell interactions in tissue culture. Here, we provide details of the methods we have used for platelet isolation from humans and mice and their staining for further interaction with tumor cells by microscopy and plate reader–based quantification. Moreover, we show the utility of this assay by demonstrating decreased platelet–tumor cell interactions in the presence of the T-Prostanoid receptor (TPr) inhibitor ifetroban. The methods described here will aid in the rapid discovery of antiplatelet agents, which have potential as anti-metastatic agents as well.
Key features
• Analysis of platelet–tumor cell binding dynamics.
• In vitro methods developed for measuring platelet–tumor cell binding to enable rapid testing of antiplatelet and other compounds.
• Complementary analysis of platelet–tumor cell binding by imaging and fluorimetry-based readings.
• Representative results screening the effect of the antiplatelet drug, ifetroban, on platelet–tumor cell binding using the protocol.
• Validation results were presented with both a TPr agonist and ifetroban (antagonist).
Graphical overview
Representative overview of the process to isolate and label platelets, incubate platelets and tumor cells in the presence of antiplatelet agents, and image and/or quantify platelet–tumor cell interactions.
Keywords: Platelet Cancer metastasis Anti-platelet therapy Platelet–tumor interaction In vitro
Background
Platelets are well known to play various crucial roles in hemostasis and thrombosis. However, growing evidence suggests that platelets are significantly involved in cancer metastasis as well (Gay and Felding-Habermann, 2011). Further, preclinical and clinical evidence has shown that platelets promote tumorigenesis and metastasis by paracrine interactions with cancer cells (Labelle et al., 2011). Cancer cells change platelet behavior by directly inducing tumor–platelet aggregates and triggering platelet activation (granule and extracellular vesicle release, altered phenotype, etc.). Further, platelet activation has been directly or indirectly linked to the dissemination of cancer cells, cancer cell survival within the circulation, and the extravasation of cancer cells at distant sites of metastasis (Xu et al., 2018). Since platelet activation pathways are likely contributors to cancer growth and metastasis, antiplatelet drugs have immense potential for the treatment of cancer metastasis by inhibiting a myriad of events that drive cancer growth and metastasis (Gay and Felding-Habermann, 2011; Wojtukiewicz et al., 2017; Lucotti et al., 2019; Tao et al., 2021). For instance, we recently discovered single nucleotide polymorphisms in TBXA2R, a gene expressed on platelets, that correlate significantly with metastasis of many solid cancers, including breast, melanoma, colon, and lung (Werfel et al., 2020). Further, we found that both hyperactivated and wild-type TPr (Thomboxane A2-Prostanoid receptor)—the protein encoded by TBXA2R—contribute to cancer metastasis in animal models. The safe and selective inhibitor of TPr, ifetroban, was able to reverse this effect and significantly reduce metastasis of cancer cell lines in the same animal models. However, antiplatelet drug efficacy against platelet–tumor cell interactions is challenging to study in vitro due to difficulties in fully replicating the dynamic and intricate processes that occur in vivo and often requires screening directly in rodent models of metastasis. These rodent studies are expensive, time-consuming, and low-throughput, which is particularly burdensome when a large number of antiplatelet drugs need to be screened. Hence, in order to rapidly screen for agents with anti-metastatic potential through disrupting platelet–tumor cell interactions, we have developed methods for the fluorescent labeling of platelets and the study of platelet–tumor cell interactions in vitro using Red Fluorescent Protein (RFP)-expressing breast cancer cells (RFP-MDA-MB-231). Using these methods, we have carried out studies to quantify the interaction of platelets and tumor cells in culture, alongside other reports also showing that platelets play key roles in the survival and spreading of tumor cells early during the metastatic cascade (Ponert et al., 2018; Lucotti et al., 2019).
Materials and reagents
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A3059)
Coulter counter (Beckman; 50 μM aperture tube, 3–30 fl particles)
CellTrackerTM Green CMFDA Dye (Thermo ScientificTM, catalog number: C7025)
Dextrose (Sigma-Aldrich, catalog number: D9434-250)
DPBS (Ca2+/Mg2+ free) (Gibco, catalog number: 14190359)
Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, catalog number: 11965118)
EDTA (Sigma-Aldrich, catalog number: E5134)
Fetal bovine serum (FBS) (Gibco, catalog number: 26140079)
Calcium-free Hank’s Balanced Salt Solution (HBSS) (Sigma-Aldrich, catalog number: H9394)
HEPES (Sigma-Aldrich, catalog number: H2034)
Hydrochloric acid (Thermo Fisher ScientificTM, catalog number: A466-2)
Ifetroban (Medchem express, catalog number: HY-105218)
Mycoplasma-free MDA-MB-231-RFP cells (Gentarget Inc, catalog number: SC057-Bsd), and other tumor cells/xenografts fluorescent protein–incorporated or later staining in the confocal step models can be tested in a similar way
Paraformaldehyde 16% aqueous solution EM grade (Electron Microscopy Sciences, catalog number: 15710-S)
PBS, pH 7.4 (Thermo Fisher ScientificTM, catalog number: 10010023)
Penicillin-streptomycin (P/S; 10,000 U/mL) (Gibco, catalog number: 15140122)
Potassium chloride (KCl) (Thermo Fisher ScientificTM, catalog number: P217-500)
Prostaglandin E1 (PGE1) (Thermo Fisher ScientificTM, catalog number: P5515)
Sodium citrate (Sigma-Aldrich, catalog number: C8532)
Sodium chloride (NaCl) (MP Biomedicals, catalog number: 194848)
Sodium hydrogen carbonate (NaHCO3) (Sigma-Aldrich, catalog number: S6297-250)
Sodium phosphate monobasic (NaH2PO4) (Sigma-Aldrich, catalog number: 71505-250)
TrypLETM Express (Gibco, catalog number: 12604-021) or 0.05% Trypsin/EDTA (Gibco, catalog number: 15400-054)
U46619 (Cayman Chemicals, catalog number:16450)
Vectashield Antifade Mounting Medium with DAPI (Cole-Parmer®, catalog number: H-1200-10)
Triton-X 100 (Thermo Fisher ScientificTM, catalog number: A16046)
Solutions
Sodium citrate for anticoagulation of blood (see Recipes)
HBSS with EDTA (see Recipes)
DMEM complete media (see Recipes)
Washing buffer for platelets (see Recipes)
Resuspension buffer for platelets (see Recipes)
MTH buffer (see Recipes)
Recipes
Sodium citrate for anticoagulation of blood
Prepare a 3.8% solution of sodium citrate by dissolving 380 mg of sodium citrate in 7 mL of water (ddH2O).
Adjust pH to 7 with concentrated hydrochloric acid.
Add water to a final volume of 10 mL.
Store the solution at 2–8 °C for six months. Allow the solution to reach room temperature before use.
To anticoagulate blood with 0.38% sodium citrate final concentration, add 1 mL of 3.8% sodium citrate solution to 9 mL of whole blood. Adjust the volume of sodium citrate according to the final blood volume.
HBSS with EDTA
Add 250 μL of 1 M EDTA to 50 mL of HBSS and mix well.
Adjust pH to 6.4 with concentrated hydrochloric acid.
Store the solution at 2–8 °C for two weeks. Allow the solution to reach room temperature before use.
DMEM complete media
Remove the 50 mL of plain media from the fresh DMEM bottle in the cell culture hood.
Add 50 mL of heat-inactivated FBS to the DMEM bottle containing 450 mL of plain media.
Add 5 mL of Pen/Strep reagent (100×) to the above mixture and mix it gently.
Washing buffer for platelets
Add 0.1% wt/vol NaHCO3 + 0.1% wt/vol BSA + 1 mM EDTA (Mw = 380.35 g/mol).
Add 10% MTH (Recipe 6) vol/vol in dH2O.
Prepare all of them in ultrapure distilled water.
Sterilize each buffer after preparation using a sterile filter unit inside the cell culture hood.
Resuspension buffer for platelets
Add 0.1% wt/vol NaHCO3 + 0.1% wt/vol BSA.
Add 10% MTH vol/vol in dH2O.
Prepare all of them in ultrapure distilled water.
Sterilize each buffer after preparation using a sterile filter unit inside the cell culture hood.
MTH buffer
Add 134 mM NaCl (Mw = 58.44) + 0.3 mM NaH2PO4·2H2O (Mw = 156.01).
3 mM KCl (Mw = 74.55) + 5 mM HEPES (Mw = 238.3) + 5 mM dextrose (Mw = 180.16) + 2 mM MgCl2 (Mw = 203.3)
Prepare all of them in ultrapure distilled water.
Sterilize each buffer after preparation using a sterile filter unit inside the cell culture hood.
Laboratory supplies
Pipette tips (Gilson, catalog numbers: F167104, F167103, F167101)
Pipettes (Eppendorf, catalog numbers: 3123000012, 3123000020, 3125000109, 3123000055, 3123000063)
1 mL syringe (BD, catalog number: 309628)
10 mL syringe (BD, catalog number: 305959)
50 mL syringe (BD, catalog number: 300865)
0.22 μm syringe filter (Millex®, catalog number: SLGP033RS)
21 G syringe needle (Sarstedt, catalog number: 85.1162)
23 G syringe needle (BD, catalog number: 300800)
27 G syringe needle (BD, catalog number: 302200)
1.5 mL microcentrifuge tubes (Thermo ScientificTM, catalog number: 69715)
2 mL microcentrifuge tubes (Thermo ScientificTM, catalog number: 69720)
15 mL centrifuge tubes (Thermo ScientificTM, catalog number: 339650)
Black 96-well plate (Thermo ScientificTM, catalog number: 229411)
100 mm cell culture dishes (Thermo ScientificTM, catalog number: 150464)
48-well cell culture plate (Thermo ScientificTM, catalog number: 8715064)
Collagen-coated 35 mm dish coverslip (MatTek Corporation, catalog number: NC9125998)
Cell counting chamber slides (Thermo ScientificTM, catalog number: C10314)
Transwell® inserts (Corning, catalog number: 3401)
Round-bottom 10 mL Plastic centrifuge tube (Thermo Fisher ScientificTM, catalog number: 02-689-6, BD VacutainerTM supplier)
EDTA/heparin-coated or Microhematocrit capillary tube (Thermo Fisher ScientificTM, catalog number: 22-362566)
Equipment
-20 °C and -80 °C freezers
Centrifuge with swing-out rotor suitable for 15 mL centrifugation tubes (Eppendorf, model: 5810R)
Liquid nitrogen dewar (Thermo Fisher ScientificTM, catalog number: CY509113)
37 °C incubator (Thermo Fisher ScientificTM, catalog number: FSGPD20)
Mouse restrainer (Braintree Scientific, model: SHORTI STD)
Analytical weighing scale (Mettler Toledo, ME104E and ME204E)
Laminar flow cabinet (Thermo ScientificTM 1300 Series Class II, catalog number: 1323TS)
CO2 cell culture incubator (Thermo Fisher ScientificTM, HERAcell VIOS 160i, catalog number: 51030403)
Benchtop centrifuge (Eppendorf, model: 5810R)
Improved Neubauer cell counting chamber (EMS, catalog number: 68052-16)
LSETM low-speed orbital shaker (CorningTM, catalog number: 6780FP)
Countess II automated cell counter (Invitrogen, Thermo Fisher ScientificTM, catalog number: AMQAX1000)
Brightfield microscope (Thermo Fisher ScientificTM, catalog number: LMI6PH1)
Microhematocrit capillary tubes (Thermo Fisher ScientificTM, catalog number: 22-362574)
Microplate reader (BioTek, model: Synergy H1). Fluorescence intensity details: Top, fluorescein 2.5 pM (0.25 fmol/well 384-well plate); bottom, fluorescein 4 pM (0.4 fmol/well 384-well plate); bandpass = 18 nm (excitation and emission), fluorescein 0.25 pM (0.025 fmol/well 384-well plate); light source: Xenon flash lamp, wavelength range: 230–999 nm, 1 nm increment, bandpass: 4 nm (230–285 nm), 8 nm (> 285 nm) and wavelength range: monochromators: 250–700 nm, detection system: PMT detectors
Confocal microscope (LEICA SP8X, Wetzlar, Germany):
Objectives: HC PL APO CS2 40×/1.10 water
Lasers: 405 Diode (on), 442 diode, WLL (on, 70.00%)
Channels: Green, Gray, Blue, and Red
Laser lines (nm): 405, 442, 491, 554, 472, 491, 667, 670
Optimal settings: Scan mode (xyz), Scan speed (400 Hz), Magnification (40), Immersion (water), Pinhole (77.2 μm) and Emission wavelength (580 nm)
Software
Gen5, 3.05 (BioTek Instrument)
Prism, 9.2.0 (GraphPad)
Confocal microscopy image analysis software (Leica application suite 3.7.6)
Photoshop 21.1 (Adobe)
ImageJ 13.0.6 (imagej.nih.gov/ij/download/)
Procedure
Perform all protocol steps for live platelets at room temperature. Do not put platelets on ice. A Schematic overview of the experimental process is shown in the Graphical overview.
Collection of human blood
Draw 5 mL of venous blood into a 10 mL syringe with a 21 G syringe needle from a consenting adult donor. This should be performed by a competent phlebotomist.
Remove the needle from the syringe and immediately and gently transfer blood into a round-bottom 10 mL centrifuge tube containing 1 mL of 3.2% sodium citrate (see Recipes).
Mix blood with sodium citrate by gently inverting the tube three times.
Collection of mouse blood
Collect 75 μL of blood from the mouse dorsal pedal vein or another venous non-terminal collection site. For blood collection from the pedal vein, place the mouse into a restrainer and puncture the vein with a 23 G and 27 G syringe needle.
Collect the blood by holding an EDTA/heparin-coated capillary tube to the puncture site. Pre-warming the mice using a heated gel pack greatly aids the identification of the pedal vein.
Alternatively, collect 900 μL of blood by cardiac puncture under terminal anesthesia with a 23 G needle and 1 mL syringe. Remove the needle from the syringe and transfer blood immediately to a 2 mL microcentrifuge tube containing 100 μL of 3.2% sodium citrate.
Production of platelet-rich plasma (PRP) from whole blood
Centrifuge whole blood at 350× g for 20 min at room temperature (RT) or 1,300× g for 10 min at 22 °C.
The blood will have separated into two layers: the lower, dark layer of packed red blood cells and the upper yellow layer of PRP. The PRP volume will be approximately 50% of the whole blood volume.
Carefully remove the upper PRP layer and transfer it to a new round-bottom centrifuge tube.
PRP may be used directly in cytometry experiments. Further processing is required to generate washed platelet suspensions.
Platelet separation and washing
Wash platelets by diluting PRP (see Procedure C) with 5 volumes of the HBSS, supplemented with 4 mM EDTA, pH 6.4, or washing buffer (see Recipes).
Centrifuge at 350× g for 20 min at RT or 1,300× g for 10 min at 22 °C.
Wash the platelet pellet gently without resuspending it three times with HBSS (supplemented with 4 mM EDTA, pH 6.4) or wash buffer containing 0.25 μM PGE1.
Washed platelets can be resuspended directly into DMEM complete media (see Recipes) or resuspension buffer (see Recipes) suitable for the experiment.
Count platelets using a coulter counter.
Stain the washed platelets (8 × 105 cells/mL) with CellTrackerTM Green dye (see section E3 below) and readjust the required concentration in resuspension buffer or directly into DMEM complete media (see Recipes).
Examination of platelets and tumor cell interactions by confocal microscopy
Cell revival and maintenance
Recover MDA-MB-231-RFP or other tumor cells from liquid nitrogen and dip the lower half of the vial into the 37 °C water bath or bead bath to thaw.
Do not let the whole content of the vial completely thaw (a small piece of ice should be visible inside the vial), wipe the vial from outside with 70% ethanol, and move to a laminar flow cell culture hood.
Open the vial and add contents into a 15 mL centrifuge tube containing pre-warmed complete media.
Centrifuge the 15 mL centrifuge tube containing cell and DMEM complete media (see Recipes) at 250× g for 5 min at RT.
After centrifugation, wipe the outside of the 15 mL centrifugation tube with 70% ethanol, move to a laminar flow cell culture hood, and decant the media in the liquid discarder without disturbing the cell pellet.
Pipette out the 1 mL fresh DMEM complete media (see Recipes) in the 15 mL centrifuge tube containing the cell pellet and resuspend the cells by slowly inverting the tube. Disperse the cells in a 100 mm dish plate containing 10 mL OF DMEM complete media (see Recipes) and keep the cultured dish plate in a 37 °C, 5% CO2 humidified cell culture incubator.
Replace the media with fresh DMEM complete media every 2–3 days.
Seeding cells into collagen-coated 35 mm dish cover glass
Once the cells reach 80%–90% confluency, remove the media from a 100 mm dish by aspiration.
Wash cells with 2–4 mL of DPBS (Ca2+/Mg2+ free) and remove the solution by aspiration.
Pipette 2–4 mL of pre-warmed 0.05% Trypsin solution into the 100 mm dish and incubate the dish in a 37 °C, 5% CO2 humidified cell culture incubator for 5 min.
Swirl the dish to ensure the solution covers all cells.
Observe the trypsinization process under the microscope at RT.
Once cells are suspended, pipette 6 mL of DMEM complete media to the flask to inhibit trypsin activity.
Transfer the cell suspension into a 15 mL centrifuge tube and centrifuge at 250× g for 5 min to pellet the cells.
Aspirate the media from the tube without disturbing the cell pellet.
Resuspend the cells in 1 mL of complete medium by gently pipetting the cells up and down and count the cells using a cell counting chamber.
Seed cells (4 × 104 cells/dish) immediately into the 35 mm collagen-treated cover glass plates or 48-well glass bottom plates (clear-culture plates) at a density of 3.5 × 104 cells/well in 500 μL of DMEM complete media (see Recipes).
Following the addition of cells, swirl the media in the dish to distribute the cells and place the 35 mm collagen-treated cover glass and 48-well glass bottom plates in a 37 °C, 5% CO2 humidified cell culture incubator.
Platelet staining
Fluorescently label the isolated platelets using CellTrackerTM Green according to the manufacturer’s protocol with slight modifications. Platelets isolated at a concentration of 8 × 105 cells/mL were reconstituted in resuspension buffer.
Prepare the CellTrackerTM Green stock solution (1 mM) by adding the 20 μL of DMSO to a vial containing 50 μg of CellTrackerTM Green.
Prepare the working CellTrackerTM Green solution to 2 μM by diluting it in 1× PBS.
Add 200 μL of working CellTrackerTM Green solution (2 μM) to 1 mL of platelet suspension.
Immediately mix the sample by gentle pipetting and incubate the sample for 10 min at 37 °C.
After incubation, centrifuge the sample at 350× g for 20 min at RT.
Discard the supernatant and wash the pellet with warm PBS or washing buffer thrice by resuspending the pellet in PBS/washing buffer before centrifugation during each wash and after centrifugation. PBS should be discarded without disturbing the pellet.
Resuspend the platelets pellet (6 × 106/100 μL) in the resuspension buffer for treatment.
Treatment with agonist and antagonist
After the labeling, activate the platelets by incubation with an agonist (U46619, 0.3 μM in PBS) and/or antagonist (Ifetroban, 10 μM in PBS) (Table 1).
Table 1. Examples of platelet agonists or antagonists and concentrations for platelet activation
Platelet agonist or antagonist Concentration Incubation temperature Incubation time
U46619 0.3 μM 37 °C 15 min
ADP 10 μM 37 °C 1 min
Thrombin 1 U/mL 37 °C 1 min
Collagen 5 μg/mL 37 °C 15 min
Ifetroban 10 μM 37 °C 15 min
Incubate platelets for 15 min at 37 °C.
After incubation, centrifuge the sample at 350× g for 20 min at RT.
Discard the supernatant and wash the pellet with warm PBS or washing buffer thrice by resuspending the pellet in PBS/washing buffer before centrifugation during each wash and after centrifugation. PBS should be discarded without disturbing the pellet.
Resuspend the platelets pellet (6 × 106/100 μL) in the resuspension buffer or cell culture media (DMEM complete media, see Recipe) for treatment.
Treatment and confocal examination
Observe cell growth in 35 mm collagen-treated cover glass (see step E2k). If the cells are confluent (80%–90%), then add the agonist- or antagonist-treated CellTrackerTM Green-labeled platelets (6 × 106 in 100 μL of DMEM complete media or PBS) (see step E4e).
Incubate the cultures in a 37 °C, 5% CO2 humidified cell culture incubator for another 30 min.
Wash the cells thrice with warm PBS to remove unbound platelets.
Fix the cells with 2% paraformaldehyde (diluted from 16% paraformaldehyde) in PBS for 15 min in a dark room.
Gently wash the cells twice with warm PBS.
Add 0.5 mL of mounting media with DAPI to fix cells and either examine immediately under confocal microscopy or store the slides at 4 °C until further analysis.
Platelet quantification by microplate reader analysis
Repeat steps E5a–E5c.
Lyse platelets bound to tumor cells (see step E5c) with 200 μL of Triton X-100 (10% in PBS) agitated for 5 min.
Transfer cell lysates to black 96-well plates and measure the fluorescence at excitation and emission wavelengths of 485 and 525 nm, respectively, using a microplate reader. Use at least triplicate samples to quantify platelet levels.
Data analysis
Confocal imaging
For the confocal imaging, at least three independent experiments, each of which originated from a different treatment, should be performed, and multiple images should be captured for each experiment.
Capture the images of all corners and center of the culture dishes. Captured images are analyzed with ImageJ software to measure the average fluorescence pixel intensities of green fluorescence in the various treatments (such as in Figure 1A, 1B, 1C, and 1D) (Wu et al., 2013) and processed with the Photoshop software [these will be the analytical regions of interest (ROIs)].
Figure 1. Representative pictures of CellTrackerTM Green–labeled platelets interacting with MDA-MB-231-RFP cells stained with DAPI. CMFDA-labeled platelets (green color), MDA-MB-231 RFP cells (red color), and DAPI (blue color). Representative confocal images control (platelet + tumor cells) (A), thromboxane A receptor agonist, U46619-treated platelet + tumor cells (B), platelet antagonist, Ifetroban-treated platelet + tumor cells (C), and co-treatment of Ifetroban- and U46619-treated platelet + tumor cells (D). Objectives: HC PL APO CS2 40×/1.10 water, scale bar is 15 μm, and the zoom of ROI 5×. ImageJ software is used to measure the average fluorescence intensities per pixel in all the treatments.
Fiji (Fiji is just ImageJ) can be downloaded at https://fiji.sc/. Export the raw image data in a format compatible with ImageJ (e.g., tif) from the imaging system and import into ImageJ.
Go to Analyze > Set Measurements and check off the box next to Limit to Threshold. Then use Image > Adjust > Threshold to highlight the area you want to analyze, and then Analyze > Measure will give you intensity measurements in just your thresholded area.
Alternatively, draw a ROI in an area of the image around your object with one of the drawing tools (in the toolbar) and then Analyze > Measure will limit its measurement to that area. Do the same to another image to analyze the same size/shape area.
Export the green fluorescence pixel intensities data to Microsoft Excel. Calculate the mean and standard deviations (n = 3) of data from at least three independent experiments.
Present the data in mean ± SD (n = 3) and analyze the data by one-way ANOVA followed by Tukey’s test using Prism (GraphPad).
Changes in the treatment can be represented with respect to the percentage of control (normalization), where bars with different letters indicate significant (p < 0.05) differences between the groups (Toragall et al., 2023).
The represented confocal image results should be confirmed by examining three independent experiments.
Plate reader fluorometric assay
For the fluorometry assay, collect the green fluorescence reading data (n = 3) from at least three independent experiments, each of which originated from a different treatment from the plate reader.
Calculate the mean and standard deviations (n = 3) using Microsoft Excel from the above recorded data.
Present the data in mean ± SD (n = 3) and analyze the data by one-way ANOVA followed by Tukey’s test using Prism (GraphPad).
Changes in the treatment can be represented with respect to the percentage of control (Figure 2) and represent bars with different letters indicating significant (p < 0.05) differences between the group (Toragall et al., 2023) (Materials and reagents section).
Figure 2. Representative quantification of platelet (CellTrackerTM Green–labeled) and tumor cell (MDA-MB-231-RFP) interactions using a microplate reader. Data are expressed as mean ± SD (n = 3) and analyzed by one-way ANOVA followed by Tukey’s test; bars with different letters indicate significant (p < 0.05) differences between the groups. Changes in the treatment are represented with respect to the percentage of control.
Validation of protocol
The protocol presented here was validated using confocal microscopy to image platelet–tumor cell binding and fluorimetry on a plate reader to measure the overall fluorescence of platelets in platelet–tumor co-culture. Moreover, the utility of these assays to determine changes in platelet–tumor cell binding in the presence of platelet agonists and antagonists was assessed using the TPr agonist U46619 and TPr antagonist Ifetroban. Figure 1 shows confocal microscopy images of RFP-expressing MDA-MB-231 tumor cells in co-culture with platelets that were pre-labeled with CellTrackerTM Green. These images show that platelets have low, but observable, levels of binding to tumor cells under resting conditions (Figure 1A). As expected, platelet binding to tumor cells increases drastically when the TPr agonist U46619 is added to the media (Figure 1B). Since platelet–tumor cell binding was fairly low under basal conditions, the treatment of platelets with TPr antagonist ifetroban had little effect on platelet–tumor cell interactions (Figure 1C). However, pre-treatment of platelets with ifetroban prior to the addition of U46619 completely abrogated the effect of U46619 on platelet–tumor cell binding (Figure 1D), suggesting that ifetroban is effective at limiting TPr-mediated platelet–tumor cell interactions. Next, fluorimetry readings on a microplate reader were used to confirm the imaging results. Again, there was a significant increase in platelet–tumor cell binding in U46619-treated co-cultures (Figure 2), as indicated by increased platelet fluorescence after washing, whereas the fluorescence in co-cultures with ifetroban-pre-treated platelets returned back to control levels. Importantly, the trends observed by plate reader–based fluorimetry matched the relative trends between groups obtained when quantifying confocal images as well (Figure 3). Taken together, these results validate that isolated platelets and tumor cells can be co-cultured, imaged via high-resolution confocal microscopy, quantified for total fluorescence using a microplate reader, and used to rapidly screen the effectiveness of antiplatelet drugs such as ifetroban.
Figure 3. Representative quantification of platelet (CellTrackerTM Green–labeled) and tumor cell (MDA-MB-231-RFP) interactions using confocal images by ImageJ analysis of fluorescence intensities. Data are expressed as mean ± SD (n = 3) and analyzed by one-way ANOVA followed by Tukey’s test. Bars with different letters indicate significant (p < 0.05) differences between the groups. Changes in the treatment are represented with respect to the percentage of control.
General notes and troubleshooting
General notes
There are stopping points at this blood collection steps (sections A and B). The collected blood must be processed immediately for the PRP production. However, once the PRP is separated, it can be stored at room temperature for another 5 days for further experiments.
After the centrifugation, the platelets have pelleted down and supernatants (platelet poor plasma; PPP) are separated and must be recentrifuged to ensure the platelet yield/losses by handling errors (pipetting) (section B)
Ideally, the ratio of blood to sodium citrate (1:10) works well; this is what we have used in the present protocol. However, the ratio may vary with rodent models and geographical location of animals. Hence, we suggest optimizing different citrate ratios before proceeding with actual experiments (section B).
Human blood platelets have been used to label and treat tumor cells throughout the study. However, the steps can be used for mice blood platelets as well. For additional experiments with flow cytometry, 10 μL of PRP (step C3) can be used to verify the platelet activation markers (Burzynski et al., 2019) (section C).
A low level of leukocytes will be present in washed platelets. Repeated rounds of the washing protocol will reduce leukocyte load, but it does not significantly reduce the platelet count (section D).
We have used the resuspension buffer (500 μL, which should be no more than 25% of total complete media in the culture dish) for redissolving the platelet pellet to get a uniform distribution of platelets (homogenous mixture). However, one can use the complete media to dissolve it (there is no volume restriction) (section D).
For the confocal studies, seed the tumor cells in the center of the 35 mm collagen-treated cover glass plates for better examination of tumor cell and platelet interactions. Further, the 96-well clear-bottom plates can be used for the lysate assay with MDA-MB-231 RFP cell seeding density of 8.5 × 103 cells per well in 200 μL of complete media (section E2).
To avoid the platelet clustering on coverslips, avoid using coverslips that are less than 95% confluent with tumor cells for the confocal microscopy studies, keep an extra control of coverslip treated with only labeled platelets, and image under confocal microscopy (section E2).
While labeling the platelets with fluorescent dyes, make sure to count the platelets (coulter counter) before and after labeling and optimize the different concentration fluorescent dyes to the number of platelets using the plate reader (excitation and emission wavelengths of 485 and 525 nm). Consider the highest number of platelets with higher fluorescence reading for better platelet labeling (section E3).
Platelets are sensitive and should be handled gently to avoid activation. Platelet-rich plasma and washed platelets should be kept at room temperature.
Sodium citrate is the recommended anticoagulant for these protocols, as its chelation of divalent cations is reversible with the addition of CaCl2, allowing for the easy production of serum. These protocols can also be performed using other non-reversible anticoagulants, if necessary (Burzynski et al., 2019).
Repeated washes/centrifugation in case of HBSS (supplemented with 4 mM EDTA, pH 6.4) may lead to activation of platelets; this is reported from the previous publication by Burzynski et al. (2019). However, the use of the PGE-1 with washing buffer can be incorporated to limit activation of platelets during wash steps (Lucotti et al., 2019). Further, the PGE1-containing buffer should only be used after the first wash (section D).
The detection of small fluorescently labeled particles by confocal microscopy (platelets) depends on the respective area of focus. The dish or slide must be scanned thoroughly for many representative portions and the confocal images can be analyzed using ImageJ software for the quantification of bound platelets around the tumor cells. The software is suitable to detect the platelet population and can be used to calculate the percentage of tumor cells covered with platelets.
For additional information related to confocal image processing for fluorescent intensity using ImageJ software, please visit the website below.
https://www.unige.ch/medecine/bioimaging/files/1914/1208/6000/Quantification.pdf
Acknowledgments
This work was funded by American Cancer Society (ACS) grant number: RSG-21-114-01-MM. This protocol was modified from the previous publications Lucotti et al. (2019) and Ponert et al. (2018) and verified by current lab examinations.
Competing interests
The authors declare that they have no conflicts of interest, financial or otherwise.
Ethical considerations
The collection of blood from rodents is approved by the institutional animal care and use committee (IACUC) at the University of Mississippi under protocol #23-007. For the representative studies shown, all human blood was acquired commercially from BioIVT.
References
Burzynski, L. C., Pugh, N. and Clarke, M. C. (2019). Platelet Isolation and Activation Assays. Bio Protoc 9(20): e3405.
Gay, L. J. and Felding-Habermann, B. (2011). Contribution of platelets to tumour metastasis. Nat. Rev. Cancer 11(2): 123–134.
Labelle, M., Begum, S. and Hynes, R. O. (2011). Direct Signaling between Platelets and Cancer Cells Induces an Epithelial-Mesenchymal-Like Transition and Promotes Metastasis. Cancer Cell 20(5): 576–590.
Lucotti, S., Cerutti, C., Soyer, M., Gil-Bernabé, A. M., Gomes, A. L., Allen, P. D., Smart, S., Markelc, B., Watson, K., Armstrong, P. C., et al. (2019). Aspirin blocks formation of metastatic intravascular niches by inhibiting platelet-derived COX-1/thromboxane A2. J. Clin. Invest. 129(5): 1845–1862.
Ponert, J. M., Schwarz, S., Haschemi, R., Müller, J., Pötzsch, B., Bendas, G. and Schlesinger, M. (2018). The mechanisms how heparin affects the tumor cell induced VEGF and chemokine release from platelets to attenuate the early metastatic niche formation. PLoS One 13(1): e0191303.
Tao, D. L., Tassi Yunga, S., Williams, C. D. and McCarty, O. J. T. (2021). Aspirin and antiplatelet treatments in cancer. Blood 137(23): 3201–3211.
Toragall, V., Muzaffar, J. and Baskaran, V. (2023). Lutein loaded double-layered polymer nanocarrier modulate H2O2 and CoCl2 induced oxidative and hypoxia damage and angiogenic markers in ARPE-19 cells. Int. J. Biol. Macromol. 240: 124378.
Werfel, T. A., Hicks, D. J., Rahman, B., Bendeman, W. E., Duvernay, M. T., Maeng, J. G., Hamm, H., Lavieri, R. R., Joly, M. M., Pulley, J. M., et al. (2020). Repurposing of a Thromboxane Receptor Inhibitor Based on a Novel Role in Metastasis Identified by Phenome-Wide Association Study. Mol. Cancer Ther. 19(12): 2454–2464.
Wojtukiewicz, M. Z., Sierko, E., Hempel, D., Tucker, S. C. and Honn, K. V. (2017). Platelets and cancer angiogenesis nexus. Cancer and Metastasis Reviews 36(2): 249–262.
Wu, M. M., Ma, X. G. and He, J. M. (2013). Measurement of Endogenous H2O2 and NO and Cell Viability by Confocal Laser Scanning Microscopy. Bio Protoc 3(19): e920.
Xu, X. R., Yousef, G. M. and Ni, H. (2018). Cancer and platelet crosstalk: opportunities and challenges for aspirin and other antiplatelet agents. Blood 131(16): 1777–1789.
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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4,857 | https://bio-protocol.org/en/bpdetail?id=4857&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Preparation of Cardiac Extracts from Embryonal Hearts to Capture RNA–protein Interactions by CLIP
GB Giulia Buonaiuto *
VT Valeria Taliani *
CN Carmine Nicoletti
FD Fabio Desideri
MB Monica Ballarino
(*contributed equally to this work)
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4857 Views: 521
Reviewed by: Gal Haimovich Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Mar 2023
Abstract
The interaction of RNA with specific RNA-binding proteins (RBP) leads to the establishment of complex regulatory networks through which gene expression is controlled. Careful consideration should be given to the exact environment where a given RNA/RBP interplay occurs, as the functional responses might depend on the type of organism as well as the specific cellular or subcellular contexts. This requisite becomes particularly crucial for the study of long non-coding RNAs (lncRNA), as a consequence of their peculiar tissue-specificity and timely regulated expression. The functional characterization of lncRNAs has traditionally relied on the use of established cell lines that, although useful, are unable to fully recapitulate the complexity of a tissue or organ. Here, we detail an optimized protocol, with comments and tips, to identify the RNA interactome of given RBPs by performing cross-linking immunoprecipitation (CLIP) from mouse embryonal hearts. We tested the efficiency of this protocol on the murine pCharme, a muscle-specific lncRNA interacting with Matrin3 (MATR3) and forming RNA-enriched condensates of biological significance in the nucleus.
Key features
• The protocol refines previous methods of cardiac extracts preparation to use for CLIP assays.
• The protocol allows the quantitative RNA-seq analysis of transcripts interacting with selected proteins.
• Depending on the embryonal stage, a high number of hearts can be required as starting material.
• The steps are adaptable to other tissues and biochemical assays.
Graphical overview
Identification of RNA/protein interactions from developing hearts
Keywords: Embryonal cardiomyocytes Cardiomyocyte isolation RNA LncRNA RNA–protein interactions CLIP
Background
The dynamic interaction between RNA and proteins controls many aspects of gene expression and disease [1, 2]. RNA binding proteins (RBP) have been shown to affect every aspect of RNA metabolism by positively or negatively regulating transcription and splicing, cytoplasmic export, and stability of different classes of RNAs [3]. RBP can also regulate mRNA subcellular localization and promote localized translation [4]. On the other hand, non-coding regulatory sequences in the mRNA, such as 5′ or 3′ untranslated regions (UTR), can reciprocally influence protein fate by controlling protein translation [5]. The synergistic interplay between molecular partners often occurs through the formation of dynamic RNA and protein-containing condensates. In the nucleus, this spatial distribution offers many advantages for the processes of RNA transcription and processing since it concentrates the molecular machinery necessary for gene expression in the three-dimensional space. Notably, the importance of RNA/RBP interactions was shown to be favored by noncoding RNAs. Long non-coding RNAs (lncRNAs), in particular, can guide and influence their protein interactome in both nuclear and cytoplasmic compartments due to their structural versatility and highly context-specific expression [6–12]. Over time, many different approaches have been developed to study RNA–protein interactions [13–14]. These methodologies can be classified into two main types: protein-centric or RNA-centric. The first class is based on the possibility to precipitate a specific protein of interest with an antibody to identify its RNA interactome, while the second class uses modified oligos to precipitate an RNA of interest and then characterize the proteins bound to it. However, several examples in the literature highlight the high rate of false positives that characterize these techniques. Indeed, there is a high possibility to observe artefact interactions between RNA and proteins that can occur in vitro after cell lysis [13, 15, 16]. For this reason, one of the essential advancements made in the identification of in vivo RNA/protein interactions is to catch direct binding between molecules while they are occurring in viable cells thanks to the application of different cross-linking methods. In particular, the UV light wavelength of 254 nm is highly efficient for the cross-linking of RNA and proteins in close contact [17] during a cross-linking immunoprecipitation (CLIP) experiment. Another improvement is in the use of fresh tissues as starting substrate to capture the RNA/protein interactions in a more physiological context. However, since the purification of viable cells from tissues can be challenging, few examples of their use are available [18–20].
Here, we present an optimized protocol that, starting from fresh tissue (embryonal murine hearts), allows the preparation of in vivo cardiac extracts suitable for UV cross-linking and subsequent immunoprecipitation assays. The described steps allow the maintenance of in vivo RNA/protein interactions, which is an essential prerogative to identify functional molecular partners involved in the regulation of a biological process.
Materials and reagents
Biological materials
C57BL/10 wild-type (WT) mouse (JAX, catalog number: 000665)
Reagents
Ethanol (Sigma, catalog number: 32221)
Dulbecco’s phosphate buffered saline (PBS) (Sigma, catalog number: D8662)
Protein inhibitor complex (PIC) (Roche, catalog number: 11873580001)
Ribolock (Thermo Fisher Scientific, catalog number: EO0381)
Phenylmethylsulfonyl fluoride (PMSF) (Roche, catalog number: 10837091001)
Dynabeads (Thermo Fisher Scientific, catalog number: 10004D)
Tri reagent (Zymo Research, catalog number: R2050-1-200)
HEPES (Sigma, catalog number: H3375)
KCl (Sigma, catalog number: P9541)
EDTA (Sigma, catalog number: EDS)
NaF (Sigma, catalog number: 201154)
NP40 (Sigma, catalog number: I3021)
Tween 20 (Sigma, catalog number: P9416)
Dithiothreitol (DTT) (Roche, catalog number: 10708984001)
Tris–HCl (Sigma, catalog number: T1503)
NaCl (Sigma, catalog number: S9888)
SDS (PanReac Applichem, catalog number: A3942)
Proteinase K (Roche, catalog number: 3115828001)
Laemmli sample buffer (Bio-Rad, catalog number: 1610747)
Anti-MATR3 antibody [Bethyl, catalog number: A300-591A (western 1:500; IP 5 μg)]
Anti-IgG antibody (Immunoreagents Inc, catalog number: Rb-003-N; 5 μg)
Direct-zolTM RNA MiniPrep (Zymo Research, catalog number: R2050)
SuperScript VILO cDNA Synthesis kit (Thermo Fisher Scientific, catalog number: 11754050)
PowerUp SYBR-Green MasterMix (Thermo Fisher Scientific, catalog number: A25742)
Primers:
pCharme FW 5′-tttctgtttgccctggacac-3′
pCharme RV 5′ - gcactcttccttctctccga- 3′
mCharme FW 5′-ggcacagacaccaaggccag-3′
mCharme RV 5′ - gcactcttccttctctccga- 3′
Gapdh FW 5′-tgacgtgccgcctggagaaa-3′
Gapdh RV 5′-agtgtagcccaagatgcccttcag-3′
Solutions
PBT (PBS + 0.02% Tween 20)
Dissociation media (see Recipes)
NP40 lysis buffer (see Recipes)
HighSalt NP40 wash buffer (see Recipes)
Proteinase K buffer (see Recipes)
Recipes
Dissociation media (1 mL)
Note: This volume is intended for 10–12 embryonal hearts. Prepare fresh on the day of the experiment.
Reagent Final concentration Volume
PBS 0.983 mL
PIC (1 tablet in 500 μL of RNase/DNase-free water) 1× 10 μL
PMSF (0.1 M) 1× 10 μL
Ribolock (40 U/μL) 1:300 3.3 μL
Total (optional) 1 mL
NP40 lysis buffer (50 mL)
Note: Solution can be stored at +4 °C for up to six months. On the day of the experiment, add 0.5 mM DTT (1 M stock), 1× PIC, and 1:200/400 Ribolock to the needed volume (ideal volume of reaction 900 μL for each condition).
Reagent Final concentration Volume
HEPES (pH 7.5) (1 M) 50 mM 2.5 mL
KCl (1 M) 150 mM 7.5 mL
EDTA (0.5 M) 2 mM 200 μL
NaF (1 M) 1 mM 50 μL
NP40 (100 %) 0.5% 250 μL
H2O 39.5 mL
Total (optional) n/a 50 mL
HighSalt NP40 wash buffer (50 mL)
Note: Solution can be stored at +4 °C for up to six months.
Reagent Final concentration Volume
HEPES (pH 7.5) (1 M) 50 mM 2.5 mL
KCl (5 M) 500 mM 5 mL
NP40 (10 %) 0.05% 250 μL
H2O 42.250 mL
Total (optional) n/a 50 mL
Proteinase K buffer (50 mL)
Note: Solution can be stored at room temperature for up to six months. On the day of the experiment, add 0.5 mM DTT (1 M stock), 1× PIC, and 1:200/400 Ribolock to the needed volume.
Reagent Final concentration Volume
Tris–HCl (pH 7.5) (1 M) 100 mM 5 mL
NaCl (1 M) 150 mM 7.5 mL
EDTA (0.5 M) 12.5 mM 1.25 mL
SDS (20 %) 2% 5 mL
H2O 31.25 mL
Total (optional) n/a 50 mL
Laboratory supplies
1.5 mL tube (Sarstedt, catalog number: 72.706)
15 mL tube (Corning, catalog number: 430791)
Petri dish 100 mm, tissue culture treated (Corning, catalog number: 353003)
Equipment
Cell strainer 70 μm (Miltenyi Biotec, catalog number: 130-098-462)
Cell lifter (Biologix, catalog number: 70-2180)
Surgical scissors (F.S.T, catalog number: 14060-10)
Jewelers’ forceps, Dumont No. 5, L 4 1/4 (Sigma, catalog number: F6521)
Tweezer (Millipore, catalog number: XX6200006P)
Micropestle (Geneaid, catalog number: MP050)
Microscope (Zeiss, model: Axio Vert.A1)
UV-Crosslinker (Spectronics corporation, model: XL-1000)
Magnetic rack (Millipore, catalog number: 20-400)
Rotating wheel Lab roller (Labnet, catalog number: H5500)
Sonicator (Diagenode, model: Bioruptor plus, catalog number: B01020001)
Thermomixer (Eppendorf)
Procedure
This protocol is suitable for the preparation of cellular extract from UV-crosslinked cells isolated from embryonal hearts. Cardiac tissue can be highly heterogeneous and composed of different cell types, such as cardiomyocytes, cardiac fibroblasts, and endothelial cells. Due to their size (lower than strainer cutoff), all these cells are kept during the filtering step, which is mainly required to remove clumps and tissue debris. To maintain in vivo interactions, we recommend the use of freshly collected hearts as cells need to be cross-linked while they are still viable. The cardiac extracts can be used as input for CLIP assay. An IgG negative control should always be run in parallel to the specific IP to check the specificity of the antibody (see sections D–G and [11]). It is important to note that at least 1 mg of total extract per condition (IP-specific and IgG negative control) is necessary. In Taliani et al. (2023) [11], a total of ~60 hearts (E15.5) were collected for CLIP and yielded ~5.3 mg of protein extract (0.09 mg for each E15.5 heart).
Embryonal hearts isolation
Sacrifice the pregnant mouse by CO2 or cervical dislocation as approved by the Institutional Animal Use and Care Committee.
Wet the skin and the fur of the mouse with 70% ethanol to avoid samples contamination with mice hair.
Position the mouse under the hood. Pinch the skin with tweezers, pull up and incise and pull apart the skin to expose the abdomen. Cut the peritoneum: you will see all the embryos contained in the placenta (Figure 1A, upper panel).
Figure 1. Collection of developing hearts from mouse embryos. A. Representative image of mouse (E15.5) embryos enveloped in the placenta sack. Isolated single embryos are shown below. B. Zoom-in image of representative E15.5 mouse embryo and heart. Ruler is shown for measurement assessment.
Carefully cut the placenta to extract the embryos. Place the embryos in a Petri dish filled with PBS to wash away the blood (Figure 1A, lower panel).
Cut the heads of the embryos and open the chest cavity to collect the hearts (Figure 1B). Depending on the developmental stage (E10.5–E15.5), hearts can be very small (1–3 mm).
Transfer the hearts to a 1.5 mL tube with 1 mL of cold PBS buffer.
Tip: To facilitate manual dissociation (section B), do not store more than 5–6 hearts in a single 1.5 mL tube.
Manual dissociation
Attention: Prepare 1 mL of dissociation media. This volume is intended for 10–12 E15.5 embryonal hearts.
Carefully remove as much PBS as possible to let the hearts settle down to the bottom of the tube (Figure 2A, left).
Figure 2. Preparation of cardiac extract. A. 1.5 mL tubes containing E15.5 hearts (n = 5) resuspended in PBS before (left) and after (right) manual dissociation with a pestle. B. Graphic representation of the procedure used to speed up dissociation by trapping the embryonal hearts between pestle and tube.
Add 500 μL of cold dissociation media to each tube (5–6 hearts).
Mash the hearts with a pestle for 2–4 min on ice.
Further dissociate the tissue by pipetting the solution with a 1,000 μL tip until no big clumps of tissue can be observed (Figure 2A, right).
Tip: For a faster dissociation, trap the hearts between the pestle and the edge of the 1.5 mL tube (Figure 2B). It is possible to follow the dissociation state by looking through the tube against a source of light.
Cardiac cells preparation
Place the 70 μm strainer (Neonatal Heart Dissociation Kit) on a 15 mL tube on ice.
Slowly add the cardiac homogenates to the strainer with a P1000 pipette. The solution will be filtered with gravity (Video 1).
Video 1. Filtration of cardiac homogenates using a cell strainer
Wash the strainer with PBS until 5 mL of volume is reached in total.
Use a cell lifter to facilitate the filtration process by gently scraping the top of the strainer. Be careful not to break the filter membrane (Video 2).
Video 2. Cell lifter facilitates the filtration process
Pour the 5 mL of filtered cardiac homogenates in one Petri dish (100 mm of diameter) at room temperature and check cells’ viability.
Tip: Slightly move the plate, and consequently the media, to better visualize the translucent cells under a brightfield light microscope. Cells should stay in suspension (Figure 3). To visualize the cardiac cells, fluorescent or colorimetric dyes can also be used on a small volume of the filtered homogenates.
Figure 3. Cardiac cells preparation. Representative image of freshly isolated cells before UV-cross-linking step. Black arrows indicate examples of viable cells.
Extract preparation
Remove the plate lid and UV-crosslink the cells in a Spectrolinker UV Crosslinker at 254 nM with 4,000 μJ/cm2 on ice.
Harvest the cells using a cell strainer and transfer the cell suspension to a 15 mL tube.
Centrifuge at 600× g for 5 min at 4 °C.
Remove the supernatant very gently to avoid disturbing the cell pellet and snap-freeze the cell pellet on liquid nitrogen. Pellets can be stored at -80 °C for up to 12 months. Attention: To proceed with extract preparation, scale up the number of hearts and repeat sections A–B.
Resuspend the frozen pellets in 3 mL of NP40 lysis buffer.
Pipette up and down with a P1000 pipette until the solution becomes homogeneous. Split the volume evenly in three 1.5 mL tubes (1,000 μL each).
Incubate the tubes for 10 min at 4 °C with gentle shaking.
Perform six cycles of sonication at low intensity (see instrument manual) for 30 s at 4 °C with a Bioruptor Plus sonication device to ensure nuclear membrane lysis.
Centrifuge at 16,000× g for 5 min at 4 °C and collect the supernatant, which represents the total cellular extract. Attention: After centrifugation, the supernatant should be clear. If the sample is still turbid, perform an additional centrifugation (16,000× g for 5 min at 4 °C) and repeat steps D5–D9 using a lower quantity of NP40 lysis buffer (1 mL instead of 3 mL).
Quantify the protein extract concentration with Bradford assay. Then, dilute the sample in supplemented NP40 lysis buffer to adjust the final concentration to 1 mg/mL.
Beads loading
Attention: Prepare 1 mL of PBT supplemented with 1× PIC and 1:200/400 Ribolock. This volume is intended for the 2 h incubation of two reactions (IP and IgG). The washing volume is not included.
For each immunoprecipitation, use two 1.5 mL tubes labeled IP (protein-specific) and IgG (negative control). Gently mix the Dynabeads Protein G magnetic particles and add 30 μL of beads to each tube.
Wash the beads by adding 1 mL of PBT per tube.
Place the tubes on the magnetic rack. Let the beads settle towards the magnetic rack and carefully remove all the supernatant.
Repeat steps E2–E3. Carefully, aspirate the last PBT wash without touching the beads.
Remove tubes from the magnetic rack and resuspend the beads in 400 μL of PBT by pipetting.
Add the same amount of IP-specific or IgG antibodies to the corresponding tubes. Use antibody (Ab) amounts recommended by the manufacturer (usually 5–10 μg of Ab/condition). Attention: Use the same Ab isotype for IP and IgG in order to use the same beads in the following section (F).
Incubate at room temperature on a rotating wheel for 2 h.
Immunoprecipitation (IP)
Wash the IP-specific and IgG-loaded beads twice at room temperature with 1 mL of PBT. After each wash, place the tubes on a magnetic rack, let the beads settle, and carefully remove all the supernatant.
Take 10% volume of extract as INPUT (INP) and keep on ice.
On ice, mix 1 mL (1 mg) of the extract with the IP-loaded beads and 1 mL of the extract with the IgG-loaded beads.
Incubate at 4 °C overnight on a rotating wheel.
The day after, wash the beads three times with 1 mL of HighSalt NP40 wash buffer.
Resuspend the beads in 100 μL of NP40 lysis buffer.
Bring the INP sample to 100 μL of final volume with NP40 lysis buffer.
For each sample (IP, IgG, and INP), split the volume in two tubes for RNA and protein extraction. Attention: It is possible to split the 100 μL in different combinations. In Taliani et al. (2023) [11], a 3:1 v/v ratio was used for RNA (75 μL) and protein (25 μL) extraction.
RNA preparation
Add 125 μL of proteinase K buffer and 50 μL of proteinase K enzyme to the 75 μL of resuspended beads (RNA tube) to reach a final volume of 250 μL.
Incubate at 50 °C for 30 min with gentle shaking.
Spin the tubes and prepare the sample for RNA extraction by adding 3–5 volumes of Tri reagent.
Let the beads settle toward a magnetic rack and transfer the supernatant to a new tube. Repeat twice. Attention: Make sure that no residue of beads is left before proceeding. Bead contaminations can interfere with RNA extraction.
Extract and purify the total RNA to use in gene expression analyses (see section I). In Taliani et al. (2023) [11], the RNA was extracted using the Direct-zolTM RNA MiniPrep kit. Other methods can also be used (e.g., phenol-chloroform extraction). INP usually yields ~1 μg of RNA in a total of 30 μL of RNase/DNase free H2O.
Protein preparation
Add 4× Laemmli sample buffer (10 μL) and 50 mM DTT (0.5 M stock, 4 μL) to 25 μL of resuspended beads (protein tube) to reach a final volume of ~40 μL.
Heat at 95 °C for 5 min.
Let the beads settle toward a magnetic rack and transfer the supernatant to a new tube.
Store at -80 °C until western blot analysis (see section I).
CLIP experiment analysis
Check the efficacy of the target protein immunoprecipitation by western blot assay. Load half (20 μL) of the volume of each sample (INP, IP, and IgG) into the SDS-PAGE gel. Proceed with protein separation by gel electrophoresis, transfer, and immunodetection. Western blot analysis for MATR3 is shown in Figure 4 [upper panel, adapted from Taliani et al. (2023) [11], as an example of the expected outcome.
Figure 4. Analysis of CLIP results [adapted from Taliani et al. (2023) [11]. Upper panel, protein analysis: western blot analysis performed on protein samples from MATR3-CLIP assay. GAPDH protein serves as a loading control. INPUT (Inp) samples represent 10% of the total protein extracts. Lower panel, RNA analysis: RT-qPCR performed on RNA samples from MATR3-CLIP assay. pCharme enrichments were compared to mCharme and Gapdh transcripts, both used as negative controls [7, 11] in MATR3 Ip and IgG RNA samples. RT-qPCR quantification is expressed as percentage (%) of Input.
After confirmation of protein recovery, proceed with RNA analysis by RT-qPCR or RNA-sequencing. In Taliani et al. (2023) [11], the RNA samples were retrotranscribed using SuperScript VILO cDNA Synthesis kit and quantified by RT-PCR using PowerUp SYBR-Green MasterMix (hold stage: 2 min at 50 °C, 2 min at 95 °C; amplification: 40× (3 s at 95 °C, 30 s at 60 °C); melt curve: 15 s at 95 °C, 1 min at 60 °C, 30 s at 95 °C, 15 s at 60 °C). RNA was then sequenced on an Illumina Novaseq 6000 Sequencing system.
Tip: Check the outcome of CLIP by performing RT-qPCR analysis on RNA transcripts used as controls. Figure 4 (lower panel) [adapted from Taliani et al. (2023) [11]] shows an example of RT-qPCR on positive [pCharme, Desideri et al. (2020) [7]] and negative (mCharme and Gapdh) controls for MATR3 CLIP experiment.
Data analysis
RNA samples from CLIP assays are suitable for both RT-qPCR and RNA-sequencing. For RT-qPCR, the enrichment of target RNAs can be graphed as the ratio of IP over INP. In the case of RNA sequencing, a detailed description of the analysis is presented in the Material and Methods section of Taliani et al. (2023) [11] (MATR3 CLIP-seq analysis). For the identification of reliable RBP RNA interactors, at least two biological replicates must be processed.
Validation of protocol
This protocol (or parts of it) was used and validated in the following research article(s):
Taliani et al. (2023) [11]. The long noncoding RNA Charme supervises cardiomyocyte maturation by controlling cell differentiation programs in the developing heart. Elife (Figure 4, all panels; Figure 4–figure supplement 1, all panels).
General notes and troubleshooting
General notes
The cell composition from heart tissue dissociation is not pure but consists of different fractions of cardiomyocytes, cardiac fibroblasts, and endothelial cells. This heterogeneity could cloud the results if your protein/RNA target is lowly expressed or expressed in an underrepresented population of cells.
The protocol can be adapted to different starting substrates (e.g., neonatal hearts). Other tissues can also be used depending on the efficacy of viable cell purification.
Troubleshooting
Problem 1: The quantity of cells obtained from the hearts is low
Possible cause: Low starting material
Solution: Prepare and freeze multiple cell pellets
Problem 2: The solution does not flow through the 70 μm strainer
Possible cause: The remaining cell clumps in the sample clog the strainer
Solution: Recover the solution from the strainer, use a micropipette to further disrupt cell clumps, and add PBS to dilute the sample. Load the solution in the strainer, keeping the micropipette perpendicular to the strainer membrane, and apply some pressure on the filter without breaking the membrane.
Acknowledgments
The authors are grateful to Marco Simula and Alessandro Palma (Dept. of Biology and Biotechnologies "Charles Darwin", Sapienza University of Rome) for critical reading and Jessica Rea (Center for Life Nano- and Neuro-Science, Istituto Italiano di Tecnologia (IIT), Rome) for the technical help with the photos throughout the protocol. This work is supported by grants from Sapienza University (RM11916B7A39DCE5 and RM12117A5DE7A45B), Regione Lazio (2020-T0002E0001), Project “National Center for Gene Therapy and Drugbased on RNA Technology” (CN00000041). Financed by NextGenerationEU PNRR MUR – M4C2 – Action 1.4- Call “Potenziamento strutture di ricerca e di campioni nazionali di R&S” (CUP: B83C22002870006), PRIN 2022 - Progetti di Rilevante Interesse Nazionale (2022BYB33L), PRIN 2022 PNRR - Progetti di Rilevante Interesse Nazionale (P2022FFEWN) to MB.
The described protocol was used in Taliani et al. (2023) [11].
Competing interests
The authors declare no competing interests.
Ethical considerations
All procedures involving laboratory animals were performed according to the institutional and national guidelines and legislations of Italy and according to the guidelines of Good Laboratory Practice (GLP). All experiments were approved by the Institutional Animal Use and Care Committee and carried out in accordance with the law (Protocol number 82945.56).
References
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Dai, X., Zhang, S. and Zaleta-Rivera, K. (2020). RNA: interactions drive functionalities. Mol. Biol. Rep. 47(2): 1413–1434. doi: 10.1007/s11033-019-05230-7
Kelaini, S., Chan, C., Cornelius, V. A. and Margariti, A. (2021). RNA-Binding Proteins Hold Key Roles in Function, Dysfunction, and Disease. Biology 10(5): 366. doi: 10.3390/biology10050366
Thelen, M. P. and Kye, M. J. (2020). The Role of RNA Binding Proteins for Local mRNA Translation: Implications in Neurological Disorders. Front. Mol. Biosci. 6: e00161. doi: 10.3389/fmolb.2019.00161
Hinnebusch, A. G., Ivanov, I. P. and Sonenberg, N. (2016). Translational control by 5′-untranslated regions of eukaryotic mRNAs. Science 352(6292): 1413–1416. doi: 10.1126/science.aad9868
Yamazaki, T., Souquere, S., Chujo, T., Kobelke, S., Chong, Y. S., Fox, A. H., Bond, C. S., Nakagawa, S., Pierron, G., Hirose, T., et al. (2018). Functional Domains of NEAT1 Architectural lncRNA Induce Paraspeckle Assembly through Phase Separation. Mol. Cell 70(6): 1038–1053.e7. doi: 10.1016/j.molcel.2018.05.019
Desideri, F., Cipriano, A., Petrezselyova, S., Buonaiuto, G., Santini, T., Kasparek, P., Prochazka, J., Janson, G., Paiardini, A., Calicchio, A., et al. (2020). Intronic Determinants Coordinate Charme lncRNA Nuclear Activity through the Interaction with MATR3 and PTBP1. Cell Rep. 33(12): 108548. doi: 10.1016/j.celrep.2020.108548
Rea, J., Menci, V., Tollis, P., Santini, T., Armaos, A., Garone, M. G., Iberite, F., Cipriano, A., Tartaglia, G. G., Rosa, A., et al. (2020). HOTAIRM1 regulates neuronal differentiation by modulating NEUROGENIN 2 and the downstream neurogenic cascade. Cell Death Dis. 11(7): e1038/s41419-020-02738-w. doi: 10.1038/s41419-020-02738-w
Cipriano, A., Macino, M., Buonaiuto, G., Santini, T., Biferali, B., Peruzzi, G., Colantoni, A., Mozzetta, C. and Ballarino, M. (2021). Epigenetic regulation of Wnt7b expression by the cis-acting long noncoding RNA Lnc-Rewind in muscle stem cells. eLife 10: e54782. doi: 10.7554/elife.54782
Statello, L., Guo, C. J., Chen, L. L. and Huarte, M. (2021). Gene regulation by long non-coding RNAs and its biological functions. Nat. Rev. Mol. Cell Biol. 22(2): 96–118. doi: 10.1038/s41580-020-00315-9
Taliani, V., Buonaiuto, G., Desideri, F., Setti, A., Santini, T., Galfrè, S., Schirone, L., Mariani, D., Frati, G., Valenti, V., et al. (2023). The long noncoding RNA Charme supervises cardiomyocyte maturation by controlling cell differentiation programs in the developing heart. eLife 12: e81360. doi: 10.7554/elife.81360
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A Plate Growth Assay to Quantify Embryonic Root Development of Zea mays
JR Jason T. Roberts
TM Tyler J. McCubbin
DB David M. Braun
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4858 Views: 941
Reviewed by: Samik BhattacharyaErin SparksAntony Chettoor
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Original Research Article:
The authors used this protocol in The Plant Cell Oct 2021
Abstract
Murashige-Skoog medium solutions have been used in a variety of plant plate growth assays, yet most research uses Arabidopsis thaliana as the study organism. For larger seeds such as maize (Zea mays), most protocols employ a paper towel roll method for experiments, which often involves wrapping maize seedlings in wet, sterile germination paper. What the paper towel roll method lacks, however, is the ability to image the roots over time without risk of contamination. Here, we describe a sterile plate growth assay that contains Murashige-Skoog medium to grow seedlings starting two days after germination. This protocol uses a section of a paper towel roll method to achieve uniform germination of maize seedlings, which are sterilely transferred onto large acrylic plates for the duration of the experiment. The media can undergo modification to include an assortment of plant hormones, exogenous sugars, and other chemicals. The acrylic plates allow researchers to freely image the plate without disturbing the seedlings and control the environment in which the seedlings are grown, such as modifications in temperature and light. Additionally, the protocol is widely adaptable for use with other cereal crops.
Key features
• Builds upon plate growth methods routinely used for Arabidopsis seedlings but that are inadequate for maize.
• Real-time photographic analysis of seedlings up to two weeks following germination.
• Allows for testing of various growth conditions involving an assortment of additives and/or modification of environmental conditions.
• Samples are able to be collected for genotype screening.
Graphical overview
Keywords: Plate assay Murashige-Skoog Maize Imaging Embryonic root system Embryonic shoot system
Background
Murashige-Skoog (MS) medium solutions have been used to provide the essential nutrients for plant development in root growth assays and can readily undergo alterations to incorporate a broad range of nutrients, chemicals, exogenous sugars, and plant hormones. A variety of these assays are routinely used in growing Arabidopsis thaliana (LaMontagne et al., 2016; Collins et al., 2020; Deslandes-Hérold et al., 2022; Mathew et al., 2023; Montpetit et al., 2023) and have included additives such as sodium chloride (Zhou et al., 2018), polyethylene glycol (PEG) (Zheng et al., 2016), and gibberellins (Unterholzner et al., 2015). For crops that produce larger seeds, such as maize, few protocols have been described employing MS media for seedling development. Most protocols utilize paper towel rolls to germinate and grow seedlings (Abdel-Ghani et al., 2016; Draves et al., 2022). Similarly, other additives can be incorporated into paper towel roll methods. A disadvantage to this method, however, includes imaging. To image the roots consistently, the rolls need to be removed from the incubator and laid out on a surface long enough to collect data before re-rolling and placing them back in the incubator, risking contamination if not carefully done. Imaging the progress of root growth over designated time intervals becomes restricted (Dowd et al., 2019).
Here, we introduce a sterile plate growth assay consisting of a half-strength MS medium used to grow maize seedlings starting two days after germination (DAG). This procedure is based on a protocol in a previously published paper (Julius et al., 2021) and was modified extensively to produce an assay functional to a wide range of experiments. Like paper towel roll methods, which can control environmental conditions such as temperature and light (Hochholdinger, 2009), this procedure also grants researchers control over the environment in which the seedlings develop. Moreover, the plates can be easily removed from an incubator and imaged at any time without disturbing the seedlings. The procedure can grow seedlings under a variety of conditions and treatments in a sterile environment, allowing researchers to investigate numerous effects in the embryonic root and shoot systems of maize and other cereal crops.
Materials and reagents
Materials
Three 24.5 cm × 24.5 cm acrylic square plates (Fischer Scientific, catalog number: 12-565-224)
Four 1.0 L bottles (w/caps)
Two stir bars
One spatula
One piece of 4" × 4" weigh paper (Fisherbrand, catalog number: 09-898-12B)
One piece of 6" × 6" weigh paper (Fisherbrand, catalog number: 09-898-12C)
One 1.0 L graduated cylinder
One 2-gallon zip-loc bag
One black marker
One 1.0 L Erlenmeyer flask
One 1.0 L beaker
One piece of 38# Regular weight seed germination paper, 10" × 15" (Anchor Paper Company, catalog number: SD3815L)
Three sets of flathead forceps
One 50 mL conical tube (Fisher Scientific, catalog number: 14-955-239)
Parafilm; 15 square (or 75 cm) strips
Aluminum foil
Plastic 6" ruler
Biological materials
This protocol was initially optimized to successfully grow maize seedlings of the B73 cultivar. Subsequent studies using other chemicals, maize mutants, and cereal seeds were performed (Slewinski and Braun, 2010; Tran et al., 2019; Huang et al., 2020; Babst et al., 2021), and examples will be provided later in this protocol. However, we recommend practicing this protocol with B73 seedlings.
Maize seedlings (B73 background)
Reagents
Bleach (liquid)
Ethanol
Spectracide (Immunox multi-purpose fungicide spray concentrate; any hardware store)
Murashige-Skoog basal medium (MP Biomedicals LLC, catalog number: 2623122)
MS vitamin solution (MP Biomedicals LLC, catalog number: 2625149)
MES (Millipore Sigma, catalog number: 6110-OP)
Phytagel (Sigma-Aldrich, catalog number: P8169)
Sodium hydroxide (NaOH) (Fisher Scientific, catalog number: S318-1)
Solutions
5% bleach (see Recipes)
5.5% spectracide (see Recipes)
70% ethanol (see Recipes)
Half-strength MS media (see Recipes)
Recipes
5% bleach
Note: Use 18.2 mΩ filtered, deionized water.
Reagent Final concentration Quantity (1.0 L)
Bleach 5% 50 mL
H2O n/a 950 mL
Total n/a 1,000 mL
5.5% spectracide
Note: Use 18.2 mΩ filtered, deionized water.
Reagent Final concentration Quantity (1.0 L)
Spectracide 5.5% 55 mL
H2O n/a 945 mL
Total n/a 1,000 mL
70% ethanol
Note: Use 18.2 mΩ filtered, deionized water. The solution will be mixed in a spray bottle; pour volumes as needed.
Reagent Final concentration Quantity (1.0 L)
Ethanol (absolute) 70% 700 mL
H2O n/a 300 mL
Total n/a 1,000 mL
Half-strength MS media
Note: Use 18.2 mΩ filtered, deionized water. Preparation of 1.0 L of media makes approximately three plates; see Procedure C for preparation instructions. Any additional ingredients must be calculated and experimented with independently.
Murashige-Skoog (MS) basal salts mixture (2.2 g/L)
MS vitamins at 1× (1 mL of vitamins per 1.0 L of media)
10 mM MES (2.13 g/L)
Phytagel (12 g/L)
18.2 mΩ filtered, deionized water (autoclaved)
Optional: Other additives can be incorporated, but the volumes need to be calculated for a 1.0 L solution.
Equipment
Incubator (Lindberg/Blue M, model: GI200C) set at 28 °C
Laminar flow hood
pH meter
200–1,000 pipettor
200–1,000 pipette tips (sterile aerosol barrier tips)
Orbital shaker (VWR, model: DS-500)
Stir plates
Weigh scale
Flatbed scanner (Ricoh, model: MP C6503)
Software
ImageJ (version 1.53k) (https://imagej.nih.gov/ij/)
SmartRoot (version 4.21) (https://smartroot.github.io/)
PDF to JPG converter (https://pdftoimage.com/pdf-to-jpg)
Smartphone app to convert images to PDFs, such as Microsoft Lens-PDF Scanner
Procedure
Sterilization techniques
It is crucial to work under sterile conditions when handling autoclaved media, equipment, and sterilized seedlings. We will indicate where sterile technique is applied in the procedure. This will be noted by “Caution: Under sterile conditions” at the beginning of the step.
Tie back long hair or wear a hat to prevent hair follicles from landing in any sterilized area and wear a face mask.
Wash your hands and forearms thoroughly with soap and warm water.
Put on a new pair of nitrile gloves, spray them with 70% ethanol, and rub your hands together.
Note: We recommend spraying your gloves with 70% ethanol periodically while working.
Autoclaving materials
Autoclaving glassware and equipment
Wrap three sets of forceps and one piece of germination paper (individually) with aluminum foil and cover the openings of one 1.0 L Erlenmeyer flask and one 1.0 L beaker with aluminum foil.
Note: If needed, you can autoclave more glassware and equipment depending on the number of experiments.
Apply a piece of autoclave tape on each piece of equipment and autoclave them.
Autoclaving water
Fill two 1.0 L bottles with 18.2 mΩ filtered, deionized water and autoclave both bottles.
Note: Loosen the cap on the bottles prior to autoclaving. You can autoclave more water if necessary.
Allow the water to cool before use.
Note: This water is used in seed rinsing (step D4), wetting the germination paper (step D9), and for the incubation process (step D12).
Half-strength MS media plate preparation
Rinse two stir bars in distilled water. Add a stir bar into two 1.0 L bottles and place on the stir plates.
Note: The media will be sterilized in step C9. Steps prior to this do not require sterile conditions or equipment. One of the bottles can be set aside until after preparation of 1.0 L of media.
Weigh out the following and transfer into one bottle: 2.13 g of MES and 2.2 g of MS basal medium.
Note: This is the step at which other additives (e.g., glucose) may be incorporated into the media, but the quantities need to be calculated prior.
Using a 200–1,000 μL pipettor, add 1.0 mL (1,000 μL) of MS vitamins.
Add 800 mL of 18.2 mΩ filtered, deionized water and turn on the stir plate.
Note: You do not need to autoclave the 1.0 L graduated cylinder prior to adding the water.
Adjust the pH to 5.7 using NaOH and a pH meter.
Weigh out and add 12 g of phytagel to the media.
Note: It is critical to add the phytagel gradually while the media is stirring rapidly. If the phytagel clumps in the media, it will improperly melt during autoclaving. We recommend having the phytagel powder on 6" × 6" weigh paper folded in half. Carefully lift the weigh paper above the mouth of the bottle and slowly add a spatula’s worth at a time. It should take several minutes to add the phytagel.
Dilute the media to 1.0 L with 18.2 mΩ filtered, deionized water; keep the media on the stir plate for 5 min.
Place the second 1.0 L bottle onto the second stir plate. Pour a 500 mL aliquot of the media into the second bottle.
Note: Aliquoting the media to 500 mL prevents the media from boiling over during autoclaving.
Leave both bottles on the stir plates with the media stirring rapidly for 5 min; autoclave the media.
While the media is in the autoclave, turn the UV lights and the fan on in the laminar flow hood to sterilize the work area.
Note: The UV light should be kept on for 15 min at minimum. However, you can keep the UV light on throughout the duration of the autoclave cycle.
After autoclaving, place the media back on the stir plates stirring rapidly; allow the media to cool until it can be handled.
Note: If you cannot pour the media at the current time, you can store it in a 60 °C oven. The media cannot be remelted after it has cooled.
Caution: Under sterile conditions. While the media is cooling, turn off the UV lamp and wipe down the surface of the laminar flow hood with 70% ethanol.
Caution: Under sterile conditions. Pour 250–350 mL aliquots of media into three acrylic square plates. Immediately place the lids on after pouring and allow to cool for at least 1 h.
Note: We recommend keeping the laminar flow hood fan on while the media cools.
Caution: Under sterile conditions. With nitrile gloves on, date and label each plate with a black marker and place them in a two-gallon zip-loc bag.
Caution: Under sterile conditions. Store lid-side up in a walk-in 4 °C cold room (or store at 4 °C). After a day, flip the plates upside down until use.
Note: Plates are good for up to a month.
Seedling germination and transplantation
Count out the number of seeds that will be used for the experiment.
Note: Keep in mind that some of the seeds will not germinate, so you may need to include more seeds in this section. A maximum of 10 seeds can grow on a plate at a time.
Place the seeds in a 50 mL conical tube.
Sterilize the seeds with 5% bleach and leave on the orbital shaker set at a speed of 150 rpm for 30 min.
Pour out the bleach. Rinse the seeds by pouring 50 mL of autoclaved 18.2 mΩ filtered, deionized water into the conical tube, sealing it with the cap, and agitating the seedlings for three seconds. Pour out the water and repeat seven more times.
Treat the seeds with 5.5% spectracide and leave on the orbital shaker set at a speed of 150 for 30 min.
Pour out the spectracide into an appropriate waste container.
Turn on the fan and the UV lights in the laminar flow hood for 15 min.
Note: The UV light should be kept on for 15 min at minimum. However, you can keep the UV light on throughout the duration of the sterilization process.
Caution: Under sterile conditions. Turn off the UV light and spray the surface with 70% ethanol.
Caution: Under sterile conditions. Unwrap and spread a sheet of sterilized germination paper across the sterile laminar flow hood surface and wet it with autoclaved 18.2 mΩ filtered, deionized water.
Note: We recommend keeping the germination paper on the aluminum foil it was autoclaved in so that it can be handled more easily if it is not in a position for rolling (see step D11).
Caution: Under sterile conditions. With sterilized forceps, arrange the sterilized seeds onto the paper spaced approximately 1.0–2.0 cm apart.
Note: If more than 15 seedlings are used, create a second row of seedlings 6–10 cm below the first row to allow space for root development. We recommend having at most 30 seedlings per piece of germination paper. Any number above 30 may make it difficult to insert the germination paper roll into the Erlenmeyer flask.
Caution: Under sterile conditions. Carefully roll the germination paper up tightly.
Caution: Under sterile conditions. Unwrap an autoclaved 1.0 L Erlenmeyer flask and add 100–200 mL of autoclaved 18.2 mΩ filtered, deionized water.
Caution: Under sterile conditions. Fold approximately three inches of the bottom of the rolled-up germination paper and place it inside the 1.0 L Erlenmeyer flask.
Caution: Under sterile conditions. Cover the flask and the paper with a sterilized 1.0 L beaker.
Caution: Under sterile conditions. Move the apparatus into a 28 °C incubator and allow the seeds 2–3 days to germinate (Figure 1).
Note: You may need to unwrap the paper to see if the seeds have germinated. Do this in the laminar flow hood with sterilized surfaces, clean hands, and a new pair of nitrile gloves sprayed with 70% ethanol.
Figure 1. Example of a paper towel roll inside an Erlenmeyer flask residing within an incubator set at 28 °C. Seedlings within the germination paper have their roots growing aligned with the gravitropic vector. The germination paper will wick up water to keep the seeds saturated.
Before transplantation, turn on the laminar flow hood fan and UV lights for 15 min.
Note: The UV light should be kept on for 15 min at minimum. However, you can keep the UV light on for a longer period of time.
Caution: Under sterile conditions. Turn off the UV light and wipe down the surface with 70% ethanol.
Caution: Under sterile conditions. Bring three growth media plates from the cold room and the apparatus containing the germinated seedlings. Re-sterilize your nitrile gloves with 70% ethanol.
Caution: Under sterile conditions. Carefully remove, unroll the germination paper, and check for a primary root length of approximately 1.0–2.0 cm.
Caution: Under sterile conditions. Take out and uncover one media plate. With sterilized forceps, gently transfer the viable seedlings to the plate and re-cover it with the lid. Repeat for the remaining plates.
Note: At most, ten seedlings can fit per plate.
Caution: Under sterile conditions. Embed each seedling into the media approximately 6.0 cm from the top of the plate. Make sure that the primary roots are facing down towards the bottom of the plate (aligned with the gravity vector) during incubation.
Note: We recommend using two sets of sterilized forceps to transfer the seedlings. Using an extra set of forceps will help keep the seedling from sliding in the media. You can dig out a seed-sized section of the media to help embed the seedling into the media, but you will have to gently push the seedling to insert it. Align the primary root to be flush with the surface of the media. Be careful not to break the developing primary root and/or the root tip. The seedling will need to be disposed of if that occurs.
Caution: Under sterile conditions. Cover the completed plate with the lid and set it off to the side (on the sterilized laminar flow hood surface). If you are transplanting more seeds, repeat steps D21–D22 until all seedlings are transplanted.
Note: We recommend wiping the laminar flow hood surface with 70% ethanol and using two new sets of sterilized tweezers between plates.
Caution: Under sterile conditions. Lift each plate up and hold it vertically to make sure none of the seedlings move or fall.
Caution: Under sterile conditions. Seal the sides of the plate(s) up with two full wraps of parafilm.
Note: You can also substitute parafilm with medipore tape.
Caution: Under sterile conditions. Label the plate with experimental details and number the seedlings on the back of the plate with a black sharpie.
Caution: Under sterile conditions. Take the plate(s) out to the flatbed scanner and scan the plate. Use color settings, convert the image to jpeg, set the dpi to 300, input a custom scan size of 10.0 in. × 10.0 in., and transfer the scanned image to a computer file.
Note: The purpose of the custom scan size is to get the scan as close to the plate’s lid edge as possible. If necessary, input different custom scan sizes to achieve this.
Caution: Under sterile conditions. Store the plate(s) in the 28 °C incubator at a near vertical angle with the primary roots facing downward.
Image the plate(s) every two days until six days after transferring the seedlings onto plates like in step D26.
Data analysis
Primary root lengths are measured using a plugin in ImageJ, SmartRoot (version 4.21), on JPG images of whole plates over the course of six days; shoots are measured using ImageJ (version 1.53k) on individual JPG images taken before analysis using a black backdrop and a downloadable PDF scanner (see Software #4) of each root system following day six. Both use the Windows operating system. Here, we provide a step-by-step guide on how to use each software to measure root and shoot growth. For ImageJ, we measured a shoot as an example; for SmartRoot, we measured a primary root. We recommend confirming the results on at least one more replicate (at least an extra ten seedlings). All images need to be converted to JPG or TIF formats before opening in ImageJ. We have listed two programs to aid in this process (see Software #3 and #4), but ImageJ should work with other software.
Using ImageJ
Open your image(s) in ImageJ.
Note: There are three ways to open an image. (1) Click on File in the toolbar, Open, and search. (2) Press CTRL + O and search. (3) Save your image to your desktop and drag the file onto the ImageJ window.
The program defaults to measure images in pixels; to set the scale, click on the line tool, click, hold on the starting point of the scale, drag the cursor through the length of the scale distance (have a ruler present in the image), release, and press CTRL + M (Figure 2).
Figure 2. Obtaining the pixel value used in setting the scale. The numbers indicate the steps at which to measure. Select the straight-line tool found in the ImageJ window (step one). Left-click the area where the scale begins (in this case, the start of the 1.0 cm mark), hold down the left mouse button, drag it until the end of the desired scale, and release (step two). Press CTRL + M to get the measurement in pixels.
A new window labeled Results will open; use the pixel value under Length for the scale. Next, click on Analyze (in the toolbar) and then Set Scale (Figure 3).
Figure 3. Opening the window to set the scale. The numbers indicate the steps to open the Set Scale window. Click on Analyze in the ImageJ toolbar (step one) and then click on Set Scale (step two).
Enter the pixel value in the Distance in pixels box, enter the scale distance (for example, 1.0) in Known distance, keep the Pixel aspect ratio at 1.0, set the Unit of length (for example, cm), and click OK (Figure 4).
Figure 4. Setting the scale using the collected pixel value. The numbers indicate the steps to enter in the scale of the image in centimeters. Enter the length (denoted by the red box) into the Distance in pixels box (step one), enter the known distance of the scale you are using (1.0 if you measured 1.0 cm segment of a ruler; step two), keep the Pixel aspect ratio box 1.0 (step three), and change the Unit of length box to “cm” (step four).
To measure, right-click on the straight-line tool, choose between the segmented line and freehand line tool, and trace out the shoot (Figure 5).
Note: If using the segmented line tool, left-click on the starting point, move the cursor down the root and left-click again. Proceed to trace the root until the root tip is reached. Either double left-click or right-click to end the trace and press CTRL + M to measure. If using the freehand line tool, hold down left-click and drag the mouse until the root tip is reached.
Figure 5. Measuring the shoot. Numbers indicate the steps to measure the shoot with the segmented line tool. Right-click the straight-line tool in the ImageJ window and select Segmented Line (step one). Start by left-clicking the spot where the shoot begins, left-click to trace the shoot up to its tip, and right-click to finish the measurement (step two). Press CTRL + M to measure.
Release the left-click and press CTRL + M to measure.
Repeat steps A5–A6 (see Data analysis) until all shoots are measured.
To save the data table (i.e., “results”), go to File on the Results window, click on Save As, and save to a location.
Using SmartRoot
Open your image(s) on ImageJ.
For each image, click on Image (in the toolbar), Type, and change the image to 8-bit (Figure 6).
Figure 6. Preparing images for analysis in SmartRoot. The numbers indicate the order to convert an image to 8-bit. Click on Image in the toolbar (step one), then Type (step two), and click on 8-bit (step three) to convert the image.
Re-save the images as jpegs onto the desktop or folder of choice.
To open SmartRoot, click on Plugins (in the toolbar), SmartRoot, then SR Explorer (Figure 7).
Note: Three new windows will open on your desktop.
Figure 7. Opening the SmartRoot plugin. The numbers indicate the order to open SmartRoot. Click on Plugins in the toolbar (step one), then click on SmartRoot (step two) and on SR Explorer (step three).
In the SmartRoot window, click on Display Axis and Display Nodes under Layers (the window defaults to this tab; Figure 8).
Figure 8. Setting up SmartRoot. The red box indicates the two items that require checkmarks in their boxes: Display Axis and Display Nodes.
Under the Data Transfer tab in the SmartRoot window, uncheck Send to SQL database and check Send to CSV file. Click on Choose and input your desired location and name for the date table; click on Save (Figure 9).
Figure 9. Setting up the file path for saving the root measurements. The numbers indicate the order at which this process occurs. Click on Data transfer (step one), uncheck Send to SQL database (step two), check Send to CSV file (step three), click on Choose and pick a folder to send the data to (step four), name the file (step five), and click Save (step six).
To open an image, click on the SmartRoot Explorer window, find the image, and double-click on it.
To measure, invert the image (CTRL + Shift + I), click on the Trace Root tool (orange circle with an orange crosshair found on ImageJ’s toolbar), left-click on the bottom of the kernel, and trace the root by left-clicking until the root tip is reached. Right-click your final point, and a new window will open; name the root and click OK (Figure 10).
Note: To see the measurement, open the SmartRoot window, click on Root List in the tabs, click on Refresh in the bottom-right corner, and click on the desired root.
Figure 10. Measuring and logging the primary root length into the data table. The numbers indicate the order at which this process occurs. Click on the trace root icon in the ImageJ window (step one). Trace the root by left-clicking on the bottom edge of the kernel, then left-clicking on the root until the root tip is reached, and right-clicking to end the measurement (step two). Enter a name for the root (step three) and click OK to log the data (step four). Click Refresh while in the Root List tab in the SmartRoot window to see the measurement appear (step five).
Measure the rest of the roots, click on Data transfer in the tabs, and click Transfer tracing data; an Excel table will appear in the location inputted earlier (Figure 11).
Note: The length measurement will appear under column D in the Excel table (labeled “length”).
Figure 11. Saving the root measurements in the designated file. In the SmartRoot tab, click on Data Transfer (step one), then click on Transfer tracing data. The measurements will appear as an Excel file in the designated location.
Validation of protocol
Results from the B73 growth experiments were validated with 43 seedlings. The lengths of primary roots and shoots are represented by standard box-and-whisker plots. Primary roots were measured four days after seedling transfer onto media plates (approximately 6 DAG; Figure 12A), and shoots measured were six days after transfer (approximately 8 DAG; Figure 12B). Three B73 plates imaged at days 0, 2, and 4 are provided for illustration of how a plate could appear (Figure 13).
Figure 12. Average primary root and shoot lengths of B73 embryonic systems. (A) Primary root lengths were measured 6 days after germination (DAG) (four days after being transferred to a plate). (B) Shoot lengths were measured 8 DAG (six days after being transferred to a plate). n = 43 replicates. Averages are represented by the X. Error bars are represented by a 95% confidence interval.
Figure 13. B73 seedling growth on MS media plates. (A) B73 seedlings are imaged after transfer onto plates [2 days after germination (DAG)], (B) two days after transfer (4 DAG), and (C) four days after transfer (6 DAG).
General notes and troubleshooting
General notes
The media should be modifiable to include a broad range of treatments. Two experiments are provided to exhibit the types of material that can be added. (1) As described by Julius et al. (2021), sucrose was supplemented into the MS media to test whether decreased root growth caused by the mutant carbohydrate partitioning defective28 (cpd28) is due to a lack of carbon mobilization. (2) We incorporated PEG (Van Der Weele et al., 2000; Verslues and Bray, 2004) to test if the MS media plates can integrate chemicals and grew B73 seedlings for comparison to plates without PEG. As expected, by infusing PEG into the MS plates, B73 seedlings grown in PEG displayed shorter primary roots 6 DAG (Figure 14A) and shoots 8 DAG (Figure 14B) than B73 seedlings on basic MS plates lacking PEG.
Figure 14. Polyethylene glycol (PEG)-treated B73 seedlings display shorter primary roots and shoots. (A) Primary root lengths are measured 6 days after germination (DAG) (four days after plate transfer). (B) Shoot lengths are measured 8 DAG (six days after plate transfer). N = 43 replicates for control; N = 20 replicates for PEG-treated. Averages represented by X, * represents P < 0.001 using a Student’s t-test. Error bars are represented by a 95% confidence interval.
Tissue samples of the roots can be collected for genotyping. For example, roots of seedlings of a previously characterized mutant, Carbohydrate partitioning defective1 (Cpd1) (Julius et al., 2018) and their wild-type siblings underwent DNA extraction (Leach et al., 2016), followed by PCR and gel electrophoresis (GE). Here, we demonstrate that we can genotype Cpd1 vs. wild-type seedlings through GE using an agarose-based gel after tissue collection, DNA extraction, and PCR (Figure 15).
Figure 15. Genotyping results of a set of Cpd1/+ and wild-type seeds. A gel image of two families (N = 10 each) segregating Cpd1/+ or wild-type individuals, with B73 and Mo17 genomic DNA backgrounds as controls with 18.2 mΩ filtered, deionized water as a no template control (NTC). The solid black bands indicate the homozygous recessive wild types, and the double bands (red arrows) indicate the heterozygous mutants.
This protocol can be applied to other crop plants, such as wheat, millet, sorghum, rice, soybean, common bean, etc. (Figure 16).
Figure 16. Wheat and Millet growth on MS media plates. (A) Wheat seedlings are imaged after transfer onto plates at 2 days after germination (DAG), (B) two days after transfer (4 DAG), (C) four days after transfer (6 DAG), and (D) six days after transfer (8 DAG). (E) Millet seedlings are imaged after transfer onto plates at 2 DAG, (F) two days after transfer (4 DAG), (G) four days after transfer (6 DAG), and (H) six days after transfer (8 DAG).
Troubleshooting
If the seed stock yields a lower rate of germination, substitute the old stock out for a new, fresh stock of the desired seeds.
If fungal contamination appears on the seeds, two practices can be done to prevent future contamination. (1) Examine your seed stock and check for signs of mold or other signs of decomposition. Exchange seed stock if necessary. If it is not the seed stock, (2) make new solutions of 5% bleach and 5.5% spectracide (see Recipes).
If fungal or bacterial contamination appears on the plates during an experiment, take extra caution when pouring a new set of plates and transferring seedlings. Two practices can be implemented to prevent future contamination. (1) Reduce the time the media is exposed to the environment when working under the laminar flow hood. (2) Minimize how far your arms hang over the plate. This can be done by having the side of the plate where the seedlings are aligned closest to you, with the primary roots facing away when inserting them into the media.
Acknowledgments
We thank three anonymous reviewers for comments that improved the manuscript. This research is supported by the National Science Foundation Plant Genome Research Program grant (IOS-1444448), the Interdisciplinary Plant Group of the University of Missouri (MU), and the MU College of Arts & Science Undergraduate Research Mentorship Program. The protocol is modified from Julius et al. (2021).
Competing interests
No competing interests to declare.
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YX Ye Xia
Published: Vol 13, Iss 20, Oct 20, 2023
DOI: 10.21769/BioProtoc.4859 Views: 714
Reviewed by: Samik BhattacharyaIgnacio Lescano Anonymous reviewer(s)
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Abstract
Strawberries are delicious and nutritious fruits that are widely cultivated and consumed around the world, either fresh or in various products such as jam, juice, and ice cream. Botrytis cinerea is a fungal pathogen that causes gray mold disease on many crops, including strawberries. Disease monitoring is an important aspect for growing commercial crops like strawberry because there is an urgent need to develop effective strategies to control this destructive gray mold disease. In this protocol, we provide an important tool to monitor the gray mold fungal infection progression in different developmental stages of strawberry. There are different types of inoculation assays for B. cinerea on strawberry plants, such as in vitro (in/on a culture medium) or in vivo (in a living plant). In vivo inoculation assays can be performed at early, middle, and late stages of strawberry development. Here, we describe three methods for in vivo inoculation assays of B. cinerea on strawberry plants. For early-stage strawberry plants, we modified the traditional fungal disc inoculation method to apply to fungal infection on strawberry leaves. For middle-stage strawberry plants, we developed the flower infection assay by dropping fungal conidia onto flowers. For late-stage strawberry plants, we tracked the survival rate of strawberry fruits after fungal conidia infection. This protocol has been successfully used in both lab and greenhouse conditions. It can be applied to other flowering plants or non-model species with appropriate modifications.
Key features
• Fungal disc inoculation on early-stage strawberry leaves.
• Fungal conidia inoculation on middle-stage strawberry flowers.
• Disease rating for late-stage strawberry fruits.
• This protocol is applicable to the other flowering plants with appropriate modifications.
Graphical overview
In vivo infection progression assays of gray mold fungus Botrytis cinerea at different developmental stages of strawberry. Created with BioRender.com.
Keywords: Strawberry Leaf inoculation Flower inoculation Gray mold disease Botrytis cinerea Fungal disc inoculation Fungal conidia inoculation Flowering plants Non-model species Commercial crops
Background
Strawberries are valuable commercial fruit crops that provide critical income and food security for many farmers and consumers around the world. Strawberry plants can be affected by bacterial and fungal infections like gray mold, anthracnose, and powdery mildew, which can damage the plant, reduce fruit quality, and even lead to plant death (Yang et al., 2023a). Botrytis cinerea is the causing agent of the gray mold disease in almost all vegetable and fruit crops, including strawberry plants and post-harvest fruits (Dean et al., 2012; Yang et al., 2023b). This necrotrophic fungal phytopathogen causes an estimated $10–100 billion yearly loss (Fillinger and Elad, 2016). The airborne conidia are produced by B. cinerea in diseased plant tissues, posing a long-lasting threat as part of the secondary infection cycle. An outbreak of the gray mold disease will likely occur under low temperature and high humidity in greenhouse and field conditions. Importantly, B. cinerea is also widely accepted as the second most important fungal pathogen in the research field of molecular plant pathology (Dean et al., 2012), ranked after Magnaporthe oryzae (Chen et al., 2023) but before Fusarium graminearum (Yang et al., 2018). Although plant species such as Arabidopsis and tomato have been used as classic models to study plant response against B. cinerea (Sun et al., 2011), other economically valuable crop species, like strawberries, are not well studied (Petrasch et al., 2019), especially from the flower and fruit perspectives. This protocol provides a detailed method for assessing the response of strawberry plants to the infection by the gray mold fungus B. cinerea at different stages of plant development. It can be adapted for other research purposes (e.g., in vitro detached leaf/fruit inoculation) and other flowering plants with appropriate modifications. The main factors to consider are the inoculation method (fungal disc vs. fungal conidia depending on the plant tissues) and the inoculation dosage (different plant species have varying levels of resistance to the fungus). For example, leaves and fruits of strawberry, tomato, and tobacco can be detached from plants and kept in optimal in vitro conditions (e.g., low temperature and high humidity) for subsequent fungal disc/conidia inoculation. This protocol can also be combined with other methods in related research fields, such as plant–microbe interactions. For instance, some beneficial bacteria can induce plant defense mechanisms that make the plants more resilient to future attacks (Sahib et al., 2019; Zhao et al., 2021; Yang et al., 2022). These defense mechanisms are called induced systemic resistance (ISR) and systemic acquired resistance (SAR) (Chanda et al., 2011; Yang et al., 2023b). This protocol can be used to study ISR/SAR of flowering plants against the gray mold fungus and help develop strategies for plant disease management from the plant’s perspective. However, this protocol has not been tested yet in field conditions and should also be modified according to plant protection principles such as the plant disease triangle, the plant fitness tetrahedron, and the plant disease management hexagon (Yang et al., 2023a), which should take into account multiple factors rather than a single one.
Materials and reagents
B. cinerea strain SS2Ap1 (isolated from diseased strawberry field by IFF)
Petri dishes (90 mm × 14 mm) (VWR, catalog number: 391-0439)
2 mL microcentrifuge tubes (Fisher Scientific, catalog number: 02-681-320)
Sterile water
Lens paper (VWR, catalog number: 52846-000)
Strawberry (day-neutral variety Monterey-UC)
2-gallon pots (greenhouse megastore)
Commercial potting mix (PRO-MIX Company, PRO-MIX BX)
Ziploc sandwich bags (Ziploc, model: 664546)
Spray bottle
Micropore tape (MicroporeTM, catalog number: 1530S-1)
BactoTM agar (BD, catalog number: 214010)
Potato dextrose agar (PDA) (Sigma-Aldrich, catalog number: 70139)
Commercial V8 100% vegetable juice (V8) (Campbell, catalog number: 0067-17KN)
Half-strength PDA medium (see Recipes)
Half-strength V8 medium (see Recipes)
Recipes
Half-strength PDA medium: for culturing B. cinerea
Half-strength of PDA diluted with sterile water.
Half-strength V8 medium: for preparing conidia suspension
Half-strength of supernatant of V8, diluted with sterile water.
Equipment
Autoclave machine (Panasonic Healthcare, model: MLS-3781L)
Sterile pipettes (VWR, catalog number: VHPA23004)
Microcentrifuge (Eppendorf, model: 5415D)
Hemocytometer (Hausser Scientific, catalog number: 497559)
Digital Calipers (Fisher Scientific, catalog number: 1464817)
Light microscope (Nikon, model: E100, required objective: 10×)
Laminar flow hood (SterilGARD 3 Advance)
Freezer (-80 °C) (Panasonic VIP Plus, model: MDF-V76VC-PA)
Vortex mixer (VWR, catalog number: 58815-232)
Laboratory gloves (Fisher Scientific, catalog number: 19-130-1597B)
Forceps (Grainger, catalog number: 4CR15)
Software and datasets
Microsoft Excel Spreadsheet Software (Microsoft 365)
GraphPad Prism (v9.0.2)
All raw example datasets are included in this protocol.
Procedure
Preparation of plant materials
Purchase fresh commercial strawberry seedlings (e.g., day-neutral variety Monterey-UC) from Lassen Canyon Nursery INC (Lassen Canyon | Strawberry Plants | Lassen Canyon Nursery). We conducted a greenhouse trial to test the susceptibility of the Monterey-UC seedlings to the fungal pathogen Botrytis cinerea, which causes gray mold disease. We inoculated the seedlings with the fungus and observed infection symptoms.
Select the uniform seedlings and transplant them in a commercial potting mix in the greenhouse (temperature: 20–32 ; humidity: 40%–60%) with a 16:8 h light/darkness cycle. We selected uniform seedlings based on the following criteria: similar root length, root biomass, and crown (stem) diameter.
Fungal inoculum preparation and inoculation
Fungal inoculum preparation
Withdraw B. cinerea SS2Ap1 strain from -80 °C glycerol stock (25%) and grow for four days on half-strength PDA at room temperature (RT).
Keep the fungal agar plate for another ~10 days at RT for conidiation.
Strawberry leaf inoculation
Punch new fungal discs from the edge of the PDA agar plates using the reverse side of sterile 1 mL tips (Figure 1A).
Inoculate the fully expanded strawberry leaves with fungal discs (Figure 1B).
Secure them with sterile tape (Figure 1C).
Punch a hole on each leaf through the attached fungal disc with a sterile needle (Figure 1D).
Spray sterile water mist on the leaf surface (Figure 1E).
Place the inoculated leaves in plastic bags to maintain high humidity (Figure 1F).
For early-stage (~2-month-old) strawberry leaf inoculation, the lesion diameter was recorded at 4 days post inoculation (dpi) (Figure 1G).
Analyze the data (Figure 1H and Table 1). “Mock” stands for the inoculation with no fungal conidia on strawberry leaves. “Infected” stands for the inoculation with fungal conidia on strawberry leaves (two-tailed unpaired t-test).
Figure 1. Procedure for inoculating strawberry leaves with Botrytis cinerea at early stage. After the seedling transplant, fresh four-day-old fungal discs (A) were taken from the agar plate edge and inoculated to early-stage (~2-month-old) strawberry leaves (B). The fungal discs' mycelia side was firmly attached to the upside of strawberry leaves with micropore tapes (C). A sterilized needle (D) was used to wound the leaf and facilitate fungal penetration. Sterile water was sprayed on infected leaves (E) and enclosed in a sandwich bag to maintain relative high humidity (F). A dark-brown necrosis lesion (G) was detected on the inoculated leaves at 4 dpi, and the lesion diameter was measured for data analysis (H). “Mock” stands for the inoculation with no fungal conidia on strawberry leaves. “Infected” stands for the inoculation with fungal conidia on strawberry leaves (two-tailed unpaired t-test).
Strawberry flower inoculation
Wash fungal conidia from the agar plates with sterile water (Figure 2A).
Filter washed conidia with sterile lens paper to a 2 mL tube and centrifuge to enrich the conidia (Figure 2B).
Dilute the conidia with half-strength V8 medium to appropriate concentration by counting conidia under a light microscope (10× objective) using a hemocytometer (Figure 2C). We used the following formula to calculate the conidial concentration using the hemocytometer: conidia concentration = N × 104 conidia/mL, where N is the number of conidia counted under the 10× objective (when 10 μL of conidial solution was applied to the hemocytometer).
Adjust the conidial suspension to the final inoculum concentration as needed. For example, we used 100 conidia/mL suspension (20 μL drop) to inoculate middle-stage (~3-month-old) strawberry flowers (Figure 2D).
Spray mint water to the inside of a plastic bag (e.g., sandwich bag) and use it to cover the inoculated flower carefully (Figure 2E).
Record the disease symptom of inoculated flowers with a 0–4 scoring system (Figure 2F).
Analyze the data (Figure 2G and Table 2). “Mock” stands for the inoculation with no fungal conidia on strawberry flowers. “Infected” stands for the inoculation with fungal conidia on strawberry flowers (two-tailed unpaired t-test).
Figure 2. Procedure for inoculating strawberry flowers with Botrytis cinerea at middle stage. Fungal conidia were washed from potato dextrose agar (PDA) plates (A), enriched by centrifuge (B), and calculated using a hemocytometer (C). A 20 μL droplet of conidia suspension was inoculated in the middle-stage (~3-month-old) newly opened strawberry flower (D) and enclosed into a sandwich bag pre-sprayed with sterile water to maintain high relative humidity (E). Four days later, disease symptoms of inoculated flowers were scored on a 0–4 scale (F) and further analyzed (G). “Mock” stands for the inoculation with no fungal conidia on strawberry flowers. “Infected” stands for the inoculation with fungal conidia on strawberry flowers (two-tailed unpaired t-test).
Strawberry fruit inoculation
Prepare the conidia inoculum as above in steps B3a–B3e, with the exception that newly emerging fruits (~3.5-month-old) are inoculated instead of flowers.
Record the disease development at 4, 8, and 12 dpi (Figure 3A). We counted the number of healthy and diseased fruits at 12 dpi.
Analyze the data (Figure 3B and Table 3). We calculated the survival rate using this formula: survival rate (%) = the number of healthy fruits/the number of total fruits × 100. “LD” stands for inoculation with low dosage of fungal conidia (100 conidia/mL). “HD” stands for inoculation with fungal high dosage of fungal conidia (5,000 conidia/mL) (two-tailed unpaired t-test).
Figure 3. Procedure of evaluating fruit survival rate to Botrytis cinerea infection at the late-flowering to early-fruiting stage. Inoculated newly emerging fruits (~3.5-month-old) were monitored until 12 dpi to evaluate fruit development (A) and its survival rate to overcome the gray mold disease (B). “LD” stands for inoculation with low dosage of fungal conidia. “HD” stands for inoculation with fungal high dosage of fungal conidia (two-tailed unpaired t-test).
Data analysis
The strawberry lesion diameter was measured using a vernier caliper/ruler.
Strawberry flower symptoms were scored on a 0–4 scale. Symptom score: 0, no symptom; 1, < 50% of the flower area shows dark brown necrosis; 2, > 50% of the flower area shows dark brown necrosis; 3, flower area is heavily covered with fungal hyphae, with additional chlorosis spreading to stem; 4, the whole flower structure collapses (see Figure 2F).
Strawberry fruit disease symptoms were assayed regarding fruit survival rate (%), overcoming the fungal infection.
All statistical analyses were performed using the GraphPad Prism v9.0.2 software. Data were presented as box & whiskers (min to max, show all points). Exact p value (two-tailed unpaired t-test) was shown in each graph.
All raw example datasets were provided in this protocol.
Validation of protocol
B. cinerea, a fungal pathogen, is known to cause gray mold disease in strawberries and many other crops (Dean et al., 2012; Yang et al., 2023b). In vivo inoculation assays have been developed in this study to assess the resistance or susceptibility of in planta strawberry leaf and flower samples to B. cinerea. The idea of infecting strawberry flowers with fungal conidia was inspired by the research of Yang et al. (2018). This protocol outlines the procedure for conducting in vivo inoculation assays at different stages of strawberry development, utilizing either fungal discs or fungal conidia suspension under realistic conditions.
The protocol has demonstrated successful implementation in both strictly controlled laboratory settings and less-controlled greenhouse environments, yielding highly effective results (see Figures 1–3). Its robustness and reproducibility are ensured through consistent employment of standardized fungal disc sizes or conidia concentrations, uniform inoculation techniques, and the use of controlled sandwich bags that mimic optimal growth conditions for B. cinerea. Additionally, the protocol incorporates an ample number of replicates (at least 24 leaves/flowers/fruits per inoculation) and appropriate statistical analysis (two-tailed unpaired t-test) to validate the outcomes and account for potential sources of variation.
The outcome of the protocol is measured by assessing lesion diameter (for leaf infections), symptom score (for flower infections), or survival rate (%, for fruit infections) after 4–12 days of incubation. These parameters reflect the degree of resistance or susceptibility exhibited by each strawberry sample to B. cinerea infection.
Below is the raw dataset as Table 1 presented in Figure 1H.
Table 1. Raw dataset presented in Figure 1H
Mock Infected
2 20
4 25
2 30
1 25
1 24
2 23
0 23
0 23
0 21
2 23
0 26
1 26
0 20
2 21
1 29
0 29
0 21
0 26
2 25
2 23
0 25
0 30
0 27
2 23
Unpaired t-test
p value < 0.0001
p value summary ****
Significantly different (p < 0.05)? Yes
One- or two-tailed p value? Two-tailed
t, df T = 36.09, df = 46
Below is the raw dataset presented in Figure 2G (Table 2).
Table 2. Raw dataset presented in Figure 2G
Mock Infected
0 2
1 3
0 4
0 4
0 4
0 4
0 3
1 3
0 3
0 3
0 4
1 4
1 3
1 3
1 2
0 3
0 3
0 3
0 2
0 4
0 4
1 2
1 2
0 3
Unpaired t-test
p value < 0.0001
p value summary ****
Significantly different (p < 0.05)? Yes
One- or two-tailed p value? Two-tailed
t, df t = 15.48, df = 46
Below is the raw dataset presented in Figure 3B (Table 3).
Table 3. Raw dataset presented in Figure 3B
LD HD
75 50
75 25
75 25
100 0
50 25
75 25
Unpaired t-test
p value 0.0003
p value summary ***
Significantly different (p < 0.05)? Yes
One- or two-tailed p value? Two-tailed
t, df t = 5.477, df = 10
General notes and troubleshooting
General notes
B. cinerea SS2Ap1 was isolated from a commercially grown and harvested strawberry in California and tends to produce mass conidia at the edge of the agar plate under darkness at RT.
Miracloth might be a better alternative to lens paper due to its fixed pore size. We used lens paper for its lower cost and higher availability.
Fungal conidia were diluted in half-strength V8 medium to facilitate reproducible pathogenesis.
The high relative humidity is critical for reproducible disease symptom development.
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: The fungal conidia production is low.
Possible cause: The culture condition is not optimal.
Solution: You can try to incubate the fungal plate under darker and cooler conditions, as this might affect its growth. Another option is to reduce the nutrient strength of the agar media, for instance, from half to 1/10.
Problem 2: The symptoms of the disease are uneven.
Possible cause: The optimal dose for inoculation has not been determined.
Solution: To find the optimal dose of inoculum, you should consider the differences in plant tissues, species, and size. It is important to note that high relative humidity is critical for the reproducibility of disease symptoms.
Acknowledgments
We want to thank the support from International Flavors & Fragrances Inc (IFF), USDA Hatch Project, and OSU-OARDC Start-Up Fund.
Competing interests
The authors declare no competing interests.
References
Chanda, B., Xia, Y., Mandal, M. K., Yu, K., Sekine, K., Gao, Q. m., Selote, D., Hu, Y., Stromberg, A., Navarre, D., et al. (2011). Glycerol-3-phosphate is a critical mobile inducer of systemic immunity in plants. Nat. Genet. 43(5): 421–427.
Chen, X., Selvaraj, P., Lin, L., Fang, W., Wu, C., Yang, P., Zhang, J., Abubakar, Y. S., Yang, F., Lu, G., et al. (2023). Rab7/Retromer‐based endolysosomal trafficking is essential for proper host invasion in rice blast. New Phytol. 239(4): 1384–1403.
Dean, R., Kan, J. a. L. V., Pretorius, Z. A., Hammond‐Kosack, K. E., Pietro, A. D., Spanu, P. D., Rudd, J. J., Dickman, M., Kahmann, R., Ellis, J., et al. (2012). The Top 10 fungal pathogens in molecular plant pathology.Mol. Plant Pathol. 13(4): 414–430.
Fillinger, S. and Elad, Y. (Eds.). (2016). Botrytis – the Fungus, the Pathogen and its Management in Agricultural Systems. Cham: Springer International Publishing.
Petrasch, S., Knapp, S. J., van Kan, J. A. L. and Blanco-Ulate, B. (2019). Grey mould of strawberry, a devastating disease caused by the ubiquitous necrotrophic fungal pathogen Botrytis cinerea. Mol. Plant Pathol 20(6): 877–892.
Sahib, M. R., Yang, P., Bokros, N., Shapiro, N., Woyke, T., Kyrpides, N. C., Xia, Y. and DeBolt, S. (2019). Improved Draft Genome Sequence of Microbacterium sp. Strain LKL04, a Bacterial Endophyte Associated with Switchgrass Plants. Microbiol. Resour. Announce. 8(45): e00927–19.
Sun, J. Q., Jiang, H. L. and Li, C. Y. (2011). Systemin/Jasmonate-Mediated Systemic Defense Signaling in Tomato. Mol. Plant 4(4): 607–615.
Yang, P., Bokros, N., Debolt, S., Zhao, Z. and Xia, Y. (2022). Genome Sequence Source of Bacillus amyloliquefaciens Strain GD4a, a Bacterial Endophyte Associated with Switchgrass Plants. Phytobiomes Journal 6(4): 354–357.
Yang, P., Chen, Y., Wu, H., Fang, W., Liang, Q., Zheng, Y., Olsson, S., Zhang, D., Zhou, J., Wang, Z., et al. (2018). The 5-oxoprolinase is required for conidiation, sexual reproduction, virulence and deoxynivalenol production of Fusarium graminearum. Curr. Genet 64(1): 285–301.
Yang, P., Zhao, L., Gao, Y. G. and Xia, Y. (2023a). Detection, Diagnosis, and Preventive Management of the Bacterial Plant Pathogen Pseudomonas syringae. Plants 12(9): 1765.
Yang, P., Zhao, Z., Fan, J., Liang, Y., Bernier, M. C., Gao, Y., Zhao, L., Opiyo, S. O. and Xia, Y. (2023b). Bacillus proteolyticus OSUB18 triggers induced systemic resistance against bacterial and fungal pathogens in Arabidopsis. Front. Plant Sci. 14: e1078100.
Zhao, Z., Bokros, N., DeBolt, S., Yang, P. and Xia, Y. (2021). Genome Sequence Resource of Bacillus sp. RRD69, a Beneficial Bacterial Endophyte Isolated from Switchgrass Plants. Mol. Plant Microbe Interact. 34(11): 1320–1323.
Article Information
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Plant Science > Plant immunity > Disease bioassay
Plant Science > Plant physiology > Biotic stress
Biological Sciences > Biological techniques > Microbiology techniques
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Detection and Cloning of Spliced Transcripts by RT-PCR
Bianca Hoffmann
Bastian Grewe
Published: Vol 3, Iss 8, Apr 20, 2013
DOI: 10.21769/BioProtoc.486 Views: 13019
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Original Research Article:
The authors used this protocol in PLOS ONE Nov 2012
Abstract
Using a Reverse Transcriptase-PCR approach spliced transcripts can be converted to cDNA, amplified and cloned into an expression plasmid. Sequencing of the obtained cDNA allows identification of the splicing events that generated the detected RNA (Grewe et al., 2012).
Keywords: Pre-mRNA Splicing Transcription RT-PCR Retrovirus
Materials and Reagents
I. RT-PCR
QuantiTect Probe RT-PCR Kit (QIAGEN, catalog number: 204443 )
Primers (sense and antisense) (Biomers)
Isolated RNA
PCR tubes (Starlab)
Nuclease-free water (B Braun)
II. Cloning
Agarose (Roth, catalog number: 2267.4 )
Ethidium bromide (AppliChem)
DNA ladders (1 kb and 100 bp) (Life Technologies, InvitrogenTM)
Geneclean III Kit (Qbiogene, catalog number: 1001-600 )
Example of an acceptor plasmid: pcDNA3.1 (Life Technologies, InvitrogenTM, catalog number: V790-20 )
Restriction endonucleases with reaction buffers (New England Biolabs)
Antarctic phosphatase with reaction buffer (New England Biolabs, catalog number: M0289L )
TaKaRa-DNA-Ligation-Kit version 2.1 (TaKaRa, catalog number: 6022 )
Ultracompetent XL2-blue bacteria (Stratagene, catalog number: 200150 )
Carbenicillin (AppliChem, catalog number: A1491 )
Bacteria cell culture tubes (Sarstedt)
Plasmid preparation Kits (QIAGEN, catalog number: 27104 , 12362 )
Tris-HCl (pH 7.8)
Sodium acetate
EDTA
NaCl
KCl
MgCl2
MgSO4
LB medium (see Recipes)
TAE buffer (see Recipes)
6x loading dye (see Recipes)
SOC medium (see Recipes)
LB agar (see Recipes)
Equipment
PCR cycler (Mastercycler gradient 5331, Eppendorf, catalog number: 5331000.010 )
Microwave oven (Alaska, model: M 1001 )
Gel electrophoresis chamber (Perfect Blue electrophoresis system 40-1214, PeqLab) and power supply (Power supply E844, Consort)
UV light box (UV-Transilluminator) (Vilber)
Incubators (Thermomixer comfort, Eppendorf, catalog number: 5355000.011 ; 37 °C shaker for bacterial cell cultures Novotron, Infors HT)
Scalpel (B Braun)
Scalpel (B Braun)
Bacteria cell culture tubes (Sarstedt)
Procedure
Depending on the experiment isolation of cytoplasmic or total RNA should be done. In general, cytoplasmic RNA is enriched in spliced RNA lacking all introns whereas total RNA contains unspliced, partially-spliced and fully-spliced RNA. A protocol for the isolation of cytoplasmic RNA from mammalian cells can be obtained from the first part of the bio-protocol "Packaging of retroviral RNA into viral particles analyzed by quantitative Reverse Transcriptase-PCR". Sense and antisense primers should hybridize upstream of the splice donor and downstream of the splice acceptor sequence, respectively (Figure 1A). It is also possible that one primer overlaps the exon-exon junction ensuring perfect hybridization to spliced RNA only. For subsequent cloning, both primers should contain recognition sites for restriction endonucleases (cloning via restriction enzymes) or sequences identical to the acceptor plasmid (cloning via homologous sequences) at their 5' ends. A cDNA containing the whole open reading frame (ORF) of a gene can be generated when sense and antisense primers hybridize to the beginning of the ORF sequence or to the 5' untranslated region and to the end of the ORF sequence or the 3' untranslated region, respectively (Figure 1B).
Set up RT-PCR reaction with the QuantiTect Probe RT-PCR Kit (QIAGEN) as follows:
PCR mix (2x, contains Taq, dNTPs and buffer)
10 μl
Reverse Transcriptase enzyme
0.2 μl
Primer sense (10 μM)
0.4 μl
Primer antisense (10 μM)
0.4 μl
Isolated RNA
0.5 to 1 μg template RNA in up to 5 μl
Add nuclease-free water to 20 μl.
Also prepare a control without addition of the Reverse Transcriptase (RT) in order to control for (genomic or plasmid) DNA contamination.
When the cDNA obtained in the RT-PCR will be inserted in an acceptor plasmid via restriction enzyme digestion it must carry the desired restriction enzyme recognition sites which can be added to the 5' ends of the primers. Be aware that approximately 4 additional random nucleotides should be added upstream of the recognition sites in the primers to allow efficient cleavage to take place.
RT-PCR reaction:
Note: The QuantiTect Probe RT-PCR Kit (Qiagen) allows the RT reaction and the PCR to be performed in one run without interruption.
RT step at 50 °C for 30 min followed by 15 min of 95 °C. Subsequently, amplify the cDNA by 30 PCR cycles starting at 95 °C for 30 sec, 50 to 60 °C depending on the melting temperature of the primers for 30 sec and 72 °C for 30 sec per 500 bp of amplicon length. It is possible to add a final extension step at 72 °C for up to 7 min to allow the DNA polymerase to elongate remaining single stranded DNA. Keep the PCR at 4 °C until agarose gel electrophoresis.
Short elongation steps are necessary to preferentially detect spliced versus unspliced/partially-spliced transcripts. Sometimes it may be necessary to perform a nested-PCR to preamplify the cDNA. Perform the first RT-PCR with primers surrounding the sequence that is amplified in the second PCR. Primers for the second PCR must bind the fragment amplified in the first RT-PCR. One microliter directly from the first RT-PCR (without any purification) should be sufficient as template for the second PCR. However, the first amplicon can also be purified by a PCR clean-up kit and up to 200 ng of DNA per 20 μl reaction can be used as template for the second PCR which can be performed with the QuantiTect Probe RT-PCR Kit omitting the RT step and starting with 15 min at 95 °C.
Prepare 1.5 % agarose gels in TAE buffer containing ethidium bromide (see recipes).
Add 4 μl of 6x loading dye to the 20 μl PCR reaction and transfer the solution to the agarose gel.
Agarose gel electrophoresis is done for approximately 30 min at 100 to 120 V (5 to 6 V per cm distance between electrodes). Analyze DNA ladder(s) in parallel to be able to determine the DNA fragment sizes. Please note that Ethidium bromide molecules are positively charged and move in the opposite direction in the agarose gel as the DNA molecules. Analyze the gel when the loading dye band reaches approximately the middle of the agarose gel (see Figure 1D).
Document the result under a UV lamp gel documentation system. The intercalated Ethidium bromide allows detection of the amplified cDNA fragments (Figure 1D). Since UV light can damage the DNA, reduce the exposure time to a minimum.
Cut out small gel pieces containing the desired DNA fragments at the UV table with a clean scalpel. Please wear laboratory safety glasses and reduce the exposure time to the UV light to a minimum.
Extract the DNA from the gel pieces with the Geneclean III Kit. Elute the DNA from the glass beads with 10 μl of nuclease-free water.
Prepare a restriction enzyme digestion of the isolated cDNA (the entire 10 μl) and of approximately 3 μg acceptor plasmid DNA in parallel:
DNA solution
10 μl
Restriction buffer (10x)
2 μl
Bovine serum albumin solution (10x)
2 μl
Restriction enzyme(s) (20 U/μl)
1 μl
Add water to 20 μl
Incubate for 1 to 3 h at 37 °C. When more than one restriction enzyme is used or when restriction enzymes are used that have star activity please increase the reaction volume to 30 μl. The enzyme(s) should not constitute more than 10 % (v/v) of the reaction volume. Star activity can also be reduced by decreasing the digestion time. However, complete digestion is critical for successful ligation and transformation. Extended incubation times up to overnight should only be performed with restriction enzymes without star activity or in increased reaction volumes (up to 50 μl).
When compatible cohesive ends are generated during digestion (e.g. by using just one restriction enzyme) the linearized acceptor plasmid must be dephosphorylated to prevent self-ligation. After restriction enzyme digestion add Antarctic phosphatase together with its buffer directly to the reaction mix. After 30 min at 37 °C the enzyme should be inactivated by incubation for 5 min at 65 °C followed by agarose gel electrophoresis.
For transient expression of genes in mammalian cells the plasmid pcDNA3.1 (Life Technologies, InvitrogenTM) (http://tools.invitrogen.com/content/sfs/vectors/pcdna3.1+.pdf) can be used as an acceptor plasmid. Due to the immediate early Cytomegalovirus (CMV) enhancer/promoter and the Bovine Growth Hormone (BGH) polyadenylation sequence it allows high expression levels. The cDNA from the RT-PCR must be inserted into the multiple cloning site. In addition, an efficient Kozak sequence (A/GCCATGG) should be added surrounding the start codon of the open reading frame which can be implemented into the sense primer during RT-PCR.
Repeat steps 3 to 8 with the digested DNA. Cut out small gel slices containing only the linearized acceptor plasmid. Contamination with still circular plasmid DNA will lead to efficient re-transformation of the original plasmid and strongly reduce the amount of bacterial colonies carrying the insert-containing plasmid.
Mix 2 μl of the digested and purified insert, 3 μl of the digested and purified acceptor plasmid DNA and 5 μl of solution I of the Takara Ligation Kit. Incubate at 16 °C for overnight. Alternatively, the DNA concentration can be measured and a vector: Insert DNA ratio of 0.03 pmol: 0.03 - 0.3 pmol can be used for ligation (manufacturer's instructions from the TaKaRa-DNA-Ligation-Kit version 2.1).
A detailed protocol for transformation of DNA and subsequent amplification of plasmids in bacteria can be found in the bio-protocol "Standard DNA Cloning" (He, 2011b) and "Plasmid DNA Extraction from E. coli Using Alkaline Lysis Method" (He, 2011a) (please generate hyperlink).
In brief, the following steps have to be performed:
Transform ultracompetent XL-2 bacteria with 1.5 μl of the ligation reaction by heat shock.
Let transformed bacteria grow for 1 h at 37 °C in SOC medium without antibiotics.
Cultivate bacteria on LB agar plates containing an appropriate antibiotic at 30 to 37 °C overnight.
Pick single colonies and let them growth overnight at 30 to 37 °C in 3 to 6 ml LB medium containing an appropriate antibiotic.
Extract plasmid DNA from 1.5 to 3 ml of the bacterial culture. Keep the rest of the cultures at 4 °C.
Digest the plasmid DNA with appropriate restriction enzymes to identify bacteria colonies containing successfully generated plasmids. In addition, sequence those plasmids that show fitting restriction enzyme digestion patterns. This sequencing reaction allows identifying the splicing event that actually generated the detected and cloned RNA.
Prepare Midi, Maxi or Giga plasmid preparations from the identified bacteria colonies harboring the successfully generated plasmid. Endotoxin-free plasmid preparation kits avoid the presence of lipopolysaccharide in the plasmid DNA solution that will be used for transfections or to treat animals.
Figure 1. Principle of the RT-PCR approach for the detection and cloning of spliced RNA. A. Sense and antisense primers (horizontal arrows) hybridize adjacent to the exon-exon boundary generated by fusion of splice donor 1 (SD1) and splice acceptor 1 (SA1) sequences during splicing. Restriction enzyme recognition sites (RE1 and RE2, vertical arrows) are present in the plasmid DNA and in the RT-PCR-amplified DNA generated from the partially-spliced RNA which can be used to insert the PCR fragment into the plasmid. B. Sense and antisense primers (horizontal arrows) contain restriction enzyme recognition sites (RE1 and RE2, vertical arrows) at their 5' ends. They hybridize in the 5' and 3' untranslated regions (UTR). Amplification of the fully-spliced RNA by RT-PCR allows insertion of the DNA into a plasmid (e.g. pcDNA3.1) using restriction enzymes 1 and 2. C. Depicted is a plasmid used to transfect HEK293T cells and its transcripts. RNAs generated by splicing between SD1 and SA5 or after an additional splicing event between SD4 and SA7 were reinserted into VHgenomic generating the plasmids VHenv or VHnef, respectively (Grewe et al., 2012). D. Shown is the agarose gel electrophoresis result obtained after RT-PCR amplification of spliced RNA from RNA isolated from cells transfected with the plasmid VHgenomic using primer set 1 containing forward primer and antisense primer 1 or primer set 2 containing forward primer and antisense primer 2 (see horizontal arrows in C) (Grewe et al., 2012). SD1-SA5 or SD1-SA5, SD4-SA7 indicates the splicing events detected after sequencing of the cDNA.
UTR, untranslated region; RE, restriction enzyme recognition site; white bars, introns; dark grey bars, open reading frame; polyA, polyadenylation signal; SD, splice donor; SA, splice acceptor; horizontal arrows, primers; vertical arrows, location of restriction enzyme recognition sites; black dot, RNA 5' CAP; black oval; 3' polyA tail; LTR, lentiviral long terminal repeats; CMV, cytomegalovirus immediate early enhancer/promoter; GFP, green fluorescence protein.
Recipes
1x TAE buffer
40 mM Tris-HCl (pH 7.8)
5 mM Sodium acetate
1 mM EDTA
Agarose
1.5 % (w/v) agarose in 1x TAE buffer
Cook for 1 min in microwave oven and shake carefully
Add 0.7 μg/ml Ethidium bromide [e.g. 7 µl of 1 % ethidium bromide solution (10 mg/ml) per 100 ml] after the solution is cooled down but still liquid and shake carefully.
6x loading dye
30 mM Tris-HCl (pH 7.6)
1 mM EDTA
35 % (v/v) glycerine
0.35 % (w/v) bromphenol blue
SOC medium
2 % (w/v) tryptone
0.5 % (w/v) yeast extract
20 mM glucose
10 mM NaCl
2.5 mM KCl
10 mM MgCl2
10 mM MgSO4
Sterilize by autoclaving.
LB agar
1.5 % (w/v) Bacto-Agar
In LB-Medium
Sterilize by autoclaving.
LB medium
1 % (w/v) tryptone
0.5 % (w/v) yeast extract
171 mM NaCl
pH 7.4
Sterilize by autoclaving.
Acknowledgments
The protocol was adapted from our paper Grewe et al. (2012). This work was funded by a grant from the German Research Foundation (DFG) to Klaus Überla (Ue45/11-1). Bianca Hoffmann is and Bastian Grewe was supported by a fellowship from the DFG graduate school (GRK 1045). Beside Bianca Hoffmann and Bastian Grewe, Katrin Ehrhardt, Maik Blissenbach, Sabine Brandt, Klaus Überla, Alexander Stang, Thomas Grunwald, Klaus Sure, and Bettina Tippler were part of the team which established the methods described.
References
Grewe, B., Ehrhardt, K., Hoffmann, B., Blissenbach, M., Brandt, S., and Uberla, K. (2012). The HIV-1 Rev protein enhances encapsidation of unspliced and spliced, RRE-containing lentiviral vector RNA. PloS One 7(11), e48688.
He F. (2011a). Plasmid DNA extraction from E. coli using alkaline lysis method. Bio-protocol 1(3): e30.
He F. (2011b). Standard DNA cloning. Bio-protocol 1(7): e52.
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4,860 | https://bio-protocol.org/en/bpdetail?id=4860&type=0 | # Bio-Protocol Content
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This is a correction notice. See the corrected protocol.
Peer-reviewed
Update Notice: Site-specific Incorporation of Phosphoserine into Recombinant Proteins in Escherichia coli
PZ Phillip Zhu
RM Ryan A. Mehl
RC Richard B. Cooley
Published: Sep 5, 2023
DOI: 10.21769/BioProtoc.4860 Views: 346
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We’d like to make a minor edit to our protocol, “Site-specific Incorporation of Phosphoserine into Recombinant Proteins in Escherichia coli”, 10.21769/BioProtoc.4541.
In the Materials and Reagents Section, in Expression strain options we list:
B95(DE3) ∆A ∆fabR ∆serB (Zhu et al., 2019). Available upon request.
We now have this strain publicly available on Addgene.
It should be changed to:
B95(DE3) ∆A ∆fabR ∆serB (Zhu et al., 2019). Available from Addgene #197655.
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4,861 | https://bio-protocol.org/en/bpdetail?id=4861&type=0 | # Bio-Protocol Content
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Peer-reviewed
Biosynthesis and Genetic Encoding of Non-hydrolyzable Phosphoserine into Recombinant Proteins in Escherichia coli
PZ Philip Zhu
RM Ryan A. Mehl
RC Richard B. Cooley
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4861 Views: 822
Reviewed by: Willy R Carrasquel-UrsulaezBhanu Jagilinki Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in ACS Central Science Apr 2023
Abstract
While site-specific translational encoding of phosphoserine (pSer) into proteins in Escherichia coli via genetic code expansion (GCE) technologies has transformed our ability to study phospho-protein structure and function, recombinant phospho-proteins can be dephosphorylated during expression/purification, and their exposure to cellular-like environments such as cell lysates results in rapid reversion back to the non-phosphorylated form. To help overcome these challenges, we developed an efficient and scalable E. coli GCE expression system enabling site-specific incorporation of a non-hydrolyzable phosphoserine (nhpSer) mimic into proteins of interest. This nhpSer mimic, with the γ-oxygen of phosphoserine replaced by a methylene (CH2) group, is impervious to hydrolysis and recapitulates phosphoserine function even when phosphomimetics aspartate and glutamate do not. Key to this expression system is the co-expression of a Streptomyces biosynthetic pathway that converts the central metabolite phosphoenolpyruvate into non-hydrolyzable phosphoserine (nhpSer) amino acid, which provides a > 40-fold improvement in expression yields compared to media supplementation by increasing bioavailability of nhpSer and enables scalability of expressions. This “PermaPhos” expression system uses the E. coli BL21(DE3) ∆serC strain and three plasmids that express (i) the protein of interest, (ii) the GCE machinery for translational installation of nhpSer at UAG amber stop codons, and (iii) the Streptomyces nhpSer biosynthetic pathway. Successful expression requires efficient transformation of all three plasmids simultaneously into the expression host, and IPTG is used to induce expression of all components. Permanently phosphorylated proteins made in E. coli are particularly useful for discovering phosphorylation-dependent protein–protein interaction networks from cell lysates or transfected cells.
Key features
• Protocol builds on the nhpSer GCE system by Rogerson et al. (2015), but with a > 40-fold improvement in yields enabled by the nhpSer biosynthetic pathway.
• Protein expression uses standard Terrific Broth (TB) media and requires three days to complete.
• C-terminal purification tags on target protein are recommended to avoid co-purification of prematurely truncated protein with full-length nhpSer-containing protein.
• Phos-tag gel electrophoresis provides a convenient method to confirm accurate nhpSer encoding, as it can distinguish between non-phosphorylated, pSer- and nhpSer-containing variants.
Graphical overview
Keywords: Phosphoserine Non-hydrolyzable phosphoserine Genetic code expansion Amber suppression Biosynthetic pathway Recombinant protein expression PermaPhos
Background
Discerning molecular mechanisms by which phosphorylation alters protein structure and function requires efficient methods to make homogenously and site-specifically phosphorylated proteins in milligram quantities for in vitro characterization. Kinases can be used to install phospho-groups onto target proteins, but the complexities of kinase specificity and their activation hinder broad application of these approaches. Genetic code expansion (GCE), on the other hand, enables production of site-specifically phosphorylated proteins by installing phospho-amino acids into proteins during translation in response to UAG (amber) stop codons. Escherichia coli GCE systems for expressing proteins homogenously modified with one or more phosphoserine (pSer) groups are well developed and becoming more widely used (Rogerson et al., 2015; Zhu et al., 2019). However, pSer proteins produced in E. coli via these GCE encoding systems can become partially dephosphorylated during expression by a few phosphatases that E. coli does harbor, frustrating downstream applications. Further, experiments requiring exposure of purified phospho-proteins to environments with phosphatases will compromise their phosphorylation status. As one example where this is an issue, “pulldowns” are a time-tested technique to discover new protein–protein interactions whereby bait proteins are mixed into cell lysates and then retrieved along with any bound prey proteins, which can then be identified by mass spectrometry or immunological assays. A phosphorylated bait protein will be hydrolyzed back to the wild-type form by phosphatases when added to eukaryotic lysates, even in the presence of phosphatase inhibitors, so that the retrieved pool of prey proteins will be a mixture of wild-type and phospho-protein interactors (Zhu et al., 2023). Consequently, it is challenging (if not impossible) to distinguish between proteins that preferentially interact with the phosphorylated or the wild-type form of the prey protein.
A GCE system to encode a stable, non-hydrolyzable mimic of pSer called phosphono-methyl-alanine (also known as 2-amino-4-phosphono-butryate and here referred to as non-hydrolyzable pSer or “nhpSer” for simplicity) was developed by Chin and colleagues in 2015 (Rogerson et al., 2015). This system relies on adding exogenous nhpSer to the culture media while expressing the same amino-acyl tRNA synthetase (RS)/tRNA pair and EF-Tu variant (EF-Sep) used for pSer incorporation. To ensure nhpSer is encoded over endogenous pSer, an E. coli ΔserC mutant is used in combination with overexpression of SerB (Figure 1). The negatively charged nhpSer does not traverse the cellular membrane effectively when exogenously added to the culture media, and so we developed a 6-step biosynthetic pathway that converts phosphoenolpyruvate into nhpSer inside the cell (Figure 2) to increase bioavailability of nhpSer for subsequent encoding into a protein, providing > 40-fold improvement in nhpSer-protein production and enabling scalability of expressions (Zhu et al., 2023). This “PermaPhos” expression system provided us with sufficient quantities of nhpSer-containing proteins to confirm that nhpSer mimics pSer function in several cases where aspartate/glutamate do not (Zhu et al., 2023). We outline here the general workflow for expressing a control protein (super folder GFP, sfGFP) containing nhpSer at site N150, and confirming correct encoding. Strategies to adopt this protocol for biologically relevant proteins are discussed.
Figure 1. Biosynthesis of serine in E. coli. In this protocol, a ∆serC mutant expression host is used as it lacks the ability to biosynthesize pSer amino acid, which would compete with nhpSer encoding. SerB is also overexpressed to hydrolyze any free pSer amino acid that might enter the cell from the media or be formed by promiscuous transaminases that can substitute for SerC function. For GCE systems that encode authentic phosphoserine, ∆serB expression hosts are used in order to build up intracellular pSer concentrations.
Figure 2. Biosynthesis of nhpSer amino acid by Streptomyces rubellomurinus FrbA, B, C, D, and E proteins. In this protocol, the pCDF-Frb-v1 plasmid expresses five Frb enzymes that, along with an unknown transaminase in E. coli, convert the central metabolite phosphoenolpyruvate into nhpSer, which is then encoded into target proteins using the GCE machinery.
Materials and reagents
Biological materials
Strains
BL21(DE3) ∆serC (Addgene, #197656). This BL21(DE3) strain of E. coli has the serC gene knocked out to prevent biosynthesis of phosphoserine, which would compete with nhpSer for incorporation into proteins (Figure 1). This strain contains Release Factor 1 (RF1), the protein responsible for terminating translation at TAG amber codons, so truncated protein will be produced along with full-length nhpSer-protein. To avoid co-purification of truncated protein with full-length protein, C-terminal purification tags are recommended. For proteins that self-assemble into homo-multimers (dimers, trimers, etc.), purification can be challenging due to the possible co-purification of truncated forms that are incorporated as subunits in the assembly. If using N-terminal purification/solubilization tags, additional purifications steps may be needed to remove truncated protein species. See General note 1 for more discussion.
DH10b (Thermo Fisher, catalog number: EC0113). This strain can be used for faithful propagation of plasmids and for cloning needs when users wish to clone their genes of interest into the pRBC plasmid (see Plasmids below). Do not use for protein expression. Though we have not explicitly tested all of them, other classical cloning strains of E. coli can be used including NEB 10-beta (New England BioLabs, catalog number: C3019H), DH5α (e.g., Thermo Fisher, catalog number: 18258012), NEB 5-alpha (New England BioLabs, catalog number: C2987H), or TOP10 (Thermo Fisher, catalog number: C404010).
Plasmids
pERM2-nhpSer (Addgene, #201922): machinery plasmid for nhpSer incorporation, kanamycin resistance with pUC origin of replication. This plasmid expresses the same tRNA synthetase and EF-Tu used for pSer incorporation (Rogerson et al., 2015). The amber codon suppressing Sep-tRNACUA is v2.0 developed by Chin and colleagues to minimize mis-aminoacylation (Zhang et al., 2017). The Sep-tRNA and tRNA-synthetase are constitutively expressed via lpp and GlnS promoters, respectively. The EF-Tu is under control of an IPTG inducible tac promoter. Also constitutively expressed via an OXB20 promoter is the E. coli serB protein to further eliminate (hydrolyze) free pSer amino acid that may come from the media or produced by promiscuous transaminases.
pCDF-Frb-v1 (Addgene, #201923): expresses Streptomyces rubellomurinus FrbA, FrbB, FrbC, FrbD, and FrbE enzymes required for the biosynthesis of nhpSer. See Zhu et al. (2023) for a detailed description of this biosynthetic pathway, which is also summarized in Figure 2. Each Frb protein is expressed under the control of an engineered T7 transcriptional promoter variant and so all Frb enzyme are expressed by the addition of IPTG. This plasmid confers spectinomycin resistance and contains the CloDF origin of replication. This plasmid is large (~11 kb) and contains repetitive elements; while we have not observed instability of this plasmid, given its large size it is good practice to minimize propagations. We recommend that when you receive the DH10b cells with the pCDF-Frb-v1 plasmid from Addgene, grow up a few individual colonies overnight in liquid media [e.g., 2× YT media supplemented with spectinomycin at 100 μg/mL (see Recipes)] and make frozen glycerol stocks of the cells for long-term storage. Glycerol stocks can be made by mixing 600 μL of overnight culture with 400 μL of sterile 50% (v/v) glycerol. Place culture tube(s) in -80 °C freezer for storage. These stocks serve as a permanent, long-term source of plasmid, which can be prepared by inoculating cultures with cells from a glob of the frozen glycerol stock. Do not thaw the glycerol stock(s) once frozen.
pRBC-sfGFP wild-type (Addgene, #174075): expresses wild-type sfGFP control protein with C-terminal His6 tag, under a T7 transcriptional promoter, ampicillin resistance/p15a origin of replication. sfGFP is expressed by the addition of IPTG.
pRBC-sfGFP 150TAG (Addgene, #174076): same as above except the sfGFP gene contains a TAG amber stop codon at site N150. The TAG codon is used to direct the translational encoding of nhpSer. Note that this is also the same plasmid used for pSer encoding (Zhu et al., 2022); any protein cloned into this vector for pSer encoding will also work for nhpSer encoding. See General note 2 for more discussions.
pRBC-[x] wild type (you must create): expresses your wild-type protein of interest (POI). You can clone your POI into pRBC by removing the sfGFP gene from the pRBC-sfGFP wt plasmid by restriction digest with NdeI and XhoI enzymes and replacing it with your POI gene using standard cloning techniques (e.g., ligation, Gibson Assembly, or SLiCE). For reasons mentioned above, a C-terminal purification tag is preferred. For helpful tips on construct design, see Troubleshooting 1.
pRBC-[x] TAG (you must create): expresses your POI with nhpSer encoded at a TAG codon. You can clone your POI into pRBC by removing the sfGFP gene by restriction digest with NdeI and XhoI and replacing with your POI gene using standard cloning techniques (e.g., ligation, Gibson Assembly, or SLiCE). Using site-directed mutagenesis, change the codon to TAG (the amber stop codon) where you intend to encode nhpSer into your POI.
Reagents
Tryptone (e.g., VWR, catalog number: 97063-386)
Yeast Extract (e.g., VWR, catalog number: 97064-368)
NaCl (e.g., VWR, catalog number: 97061-274)
Agar (e.g., VWR, catalog number: 97064-336)
MgSO4·7H2O (e.g., VWR, catalog number: 97062-134)
K2HPO4 (potassium dibasic) (e.g., VWR, catalog number: 97062-234)
KH2HPO4 (potassium monobasic) (e.g., VWR, catalog number: BDH9268)
α-D-glucose (e.g., VWR, catalog number: 97061-168)
Glycerol (e.g., VWR, catalog number: BDH24388.320)
Isopropyl b-D-1-thiogalactopyranoside (IPTG) (e.g., Anatrace, catalog number: I1003)
Ampicillin (e.g., VWR, catalog number: 97061-442)
Kanamycin (e.g., VWR, catalog number: 75856-684)
Spectinomycin (e.g., VWR, catalog number: 89156-368)
Antifoam B (e.g., J.T. Baker, catalog number: B531-05)
Phos-tag acrylamide (e.g., VWR, catalog number: 101974-086)
Dry ice
Ethanol (e.g., Sigma, catalog number: 65348-M)
Acrylamide/Bio-acrylamide solution (e.g., Bio-Rad, catalog number: 1610146)
Solutions
Phosphate Buffered Saline (PBS) (e.g., VWR, catalog number: 75800-982)
LB/agar (see Recipes)
2× YT media (see Recipes)
50% (v/v) glycerol (see Recipes)
10% (v/v) glycerol (see Recipes)
SOC media (see Recipes)
Starter culture media (see Recipes)
1.1× Terrific Broth media (see Recipes)
10× TB potassium phosphate buffer (see Recipes)
Ampicillin stock (see Recipes)
Kanamycin stock (see Recipes)
Spectinomycin stock (see Recipes)
0.5 M IPTG (see Recipes)
Recipes
LB/agar media (0.5 L)
Reagent Final concentration Quantity
Tryptone 1% (w/v) 5 g
Yeast extract 0.5% (w/v) 2.5 g
NaCl 1.0% (w/v) 5 g
Agar 1.5% (w/v) 7.5 g
H2O n/a To 500 mL
Total n/a 500 mL
After mixing reagents thoroughly, autoclave on standard liquid setting to sterilize. Note the agar will not go into solution until autoclaved.
After autoclaving, gently swirl the bottle to ensure melted agar is evenly mixed.
Notes:
i. Store LB/agar bottle in a 55 °C oven and pour plates on an as needed basis. LB/agar can be stored in molten form for ~2 weeks if sterility is maintained.
ii. If an oven is not available, plates can be poured with antibiotics once LB/agar is sufficiently cooled to touch. Plates can be stored at 4 °C for up to a week.
2× YT media (1 L)
Reagent Final concentration Quantity
Tryptone 1.6% (w/v) 16 g
Yeast extract 1.0% (w/v) 10 g
NaCl 0.5% (w/v) 5 g
H2O n/a To 1,000 mL
Total n/a 1 L
After mixing reagents thoroughly, autoclave on the standard liquid setting to sterilize.
After autoclaving, allow to cool to room temperature before use.
50% (v/v) glycerol (500 mL)
Reagent Final concentration Quantity
Glycerol (100%) 50% (v/v) 250 mL
H2O n/a 250 mL
Total n/a 500 mL
Mix by placing a suitable magnetic stir bar in a 500 mL graduated cylinder and add 250 mL of water to graduated cylinder. While stirring, pour pure glycerol to the 500 mL mark on graduated cylinder. Stir for 5 min and then transfer to a 0.5 L bottle. Sterilize by autoclaving on liquid setting.
10% (v/v) glycerol (1 L)
Reagent Final concentration Quantity
Glycerol (100%) 10% (v/v) 100 mL
H2O n/a 900 mL
Total n/a 1 L
Mix by placing a suitable magnetic stir bar in a 1,000 mL graduated cylinder and add 900 mL of water to graduated cylinder. While stirring, pour pure glycerol to the 1,000 mL mark on graduated cylinder. Stir for 5 min and then transfer to a 1 L bottle. Sterilize by autoclaving on liquid setting.
SOC media, 50 mL
Reagent Final concentration Quantity
2× YT media n/a 49 mL
1 M MgSO4 10 mM 0.5 mL
40% (w/v) α-D-glucose 0.4% (w/v) or ~20 mM 0.5 mL
Total n/a 50 mL
1 M MgSO4 can be made by mixing 12.3 g of MgSO4·7H2O in water up to 50 mL of total volume. Adjust mass of MgSO4 if using the salt with different hydration status.
40% (w/v) α-D-glucose can be made by mixing 20 g of α-D-glucose with water up to 50 mL of total volume. Mix thoroughly until glucose is dissolved. Gentle heating in a microwave may facilitate dissolution of glucose.
Sterilize MgSO4, glucose, and 2× YT solutions individually by autoclaving. Allow each component to cool to room temperature and mix as indicated above. Maintain sterility while adding components together.
It is easy to contaminate SOC. We suggest breaking this into 5 × 10 mL aliquots before use or making smaller batches. If sterility is maintained, SOC can be stored at room temperature indefinitely. It can also be stored at -20 °C but avoid repeated freeze/thaw.
Starter culture media: 2× YT + 0.5% (v/v) glycerol (50 mL)
Reagent Final concentration Quantity
2× YT media (sterile) n/a 49.5 mL
50% (v/v) glycerol (sterile) 0.5% (v/v) 0.5 mL
Total n/a 50 mL
Prepare immediately before use.
1.1× Terrific Broth media (0.9 L)
Reagent Final concentration (of 1×) Quantity
Tryptone 1.2% (w/v) 12 g
Yeast extract 2.4% (w/v) 24 g
Glycerol (50% [v/v]) 0.5% (v/v) 10 mL
H2O n/a To 900 mL
Total n/a 900 mL
Sterilize by autoclaving on liquid setting.
Do not mix TB phosphate buffer (below) until all solutions are cool and immediately before use.
10× TB potassium phosphate buffer (1 L)
Reagent Final concentration Quantity
KH2PO4 (monobasic, anhydrous) 0.17 M 23.1 g
K2HPO4 (dibasic, anhydrous) 0.72 M 125.4 g
H2O n/a To 1 L
Total n/a 1 L
The masses provided for the potassium phosphate salts are for anhydrous formulations. Hydrate forms of these salts can be used, but masses must be adjusted to maintain indicated molarity.
Sterilize by autoclaving on liquid setting.
Do not mix TB phosphate buffer to 1.1× TB media until all solutions are sterilized and cooled to room temperature, and immediately before use.
Ampicillin stock (10 mL)
Reagent Final concentration Quantity
Ampicillin 100 mg/mL 1 g
H2O n/a To 10 mL
Total n/a 10 mL
Sterilize by filtering with a 0.2 μm syringe-end filter.
Store in 1 mL aliquots at -20 °C.
Kanamycin stock (10 mL)
Reagent Final concentration Quantity
Kanamycin 50 mg/mL 0.5 g
H2O n/a To 10 mL
Total n/a 10 mL
Sterilize by filtering with a 0.2 μm syringe-end filter.
Store in 1 mL aliquots at -20 °C.
Spectinomycin stock (10 mL)
Reagent Final concentration Quantity
Spectinomycin 100 mg/mL 1 g
H2O n/a To 10 mL
Total n/a 10 mL
Sterilize by filtering with a 0.2 μm syringe-end filter.
Store in 1 mL aliquots at -20 °C.
Note: Do not confuse spectinomycin with streptomycin. These antibiotics are not interchangeable.
0.5 M IPTG (10 mL)
Reagent Final concentration Quantity
IPTG 0.5 M 1.19 g
H2O n/a To 10 mL
Total n/a 10 mL
Sterilize by filtering with a 0.2 μm syringe-end filter.
Store in 1 mL aliquots at -20 °C.
Laboratory supplies
1.7 mL Eppendorf tubes (e.g., VWR, catalog number: 87003-294)
0.6 mL Eppendorf tubes (e.g., VWR, catalog number: 89000-010)
100 mm plates (e.g., VWR, catalog number: 470210-568)
0.5 L centrifuge bottles (must fit available centrifuges/rotor)
500 mL graduate cylinder
15 mL conical tubes (e.g., VWR, catalog number: 89126-798)
50 mL conical tubes (e.g., VWR, catalog number: 89039-656)
14 mL sterile culture tubes (e.g., VWR, catalog number: 60818-689)
250 mL baffled flasks (e.g., VWR, catalog number: 89095-266)
2.8 L baffled Fernbach flasks (e.g., VWR, catalog number: 22877-168)
Micro pipette tips 10 μL (e.g., VWR, catalog number: 76323-394)
Micro pipette tips 200 μL (e.g., VWR, catalog number: 76323-390)
Micro pipette tips 1,000 μL (e.g., VWR, catalog number: 76323-454)
0.2 μm syringe end filter (e.g., VWR, catalog number: 28145-477)
10 mL syringes (e.g., VWR, catalog number: 76124-664)
Equipment
Autoclave (any capable of sterilizing liquid media and culturing materials at 121 °C, and of a saturated steam pressure of 15 PSI)
Expression equipment:
Static incubator for growing LB/agar plates (set to 37 °C) (e.g., VWR, catalog number: 97025-630)
Shaker incubator for growing liquid cultures (e.g., New Brunswick I26R, Eppendorf, catalog number: M1324-0004)
i. Shaker should be able to rotate at 200–250 rpm
ii. Refrigeration is necessary for expressions below room temperature (< 25 °C)
iii. Shaker deck should have clamps to hold 250 mL and 2.8 L Fernbach flasks
Optical density 600 nm spectrophotometer (e.g., Ultrospec 10, Biochrome, catalog number: 80-2116-30)
Competent cell preparation and cell harvesting
Centrifuge capable of speeds of at least 5,000× g and able to hold volumes commensurate with culture sizes (e.g., Beckman Coulter, model: Allegra 25R)
Centrifuge bottles that can withstand centrifugal forces and hold the volume of cultures grown for harvesting cells. The type of bottles depends on the centrifuge and rotor used
Freezer (-80 °C), if cells are to be stored before protein is purified (e.g., VIP ECO ULT freezer, VWR, catalog number: 76305-596)
Electroporator (e.g., Eppendorf Eporator, Fisher Scientific, catalog number: E4309000027)
0.1 cm electro-cuvettes (e.g., Bio-Rad, catalog number:1652089)
Fluorometer capable of reading sfGFP fluorescence (excitation 488 nm/emission 512 nm). Handheld fluorometers work well for routine fluorescence reads (e.g., Turner Biosystems, model: PicoFluor)
-20 °C freezer for storing plasmids and antibiotics (e.g., Fisher Scientific, catalog number: 10-549-264)
Ice machine (e.g., Fisher Scientific, catalog number: 09-540-003)
Procedure
In Part A, you first prepare electro-competent BL21(DE3) ∆serC cells before expressing protein in Part B. These electro-competent cells need to be highly efficient for expressions to be successful (see Troubleshooting 2). The process outlined below is sufficient to produce enough of electro-competent cells for ~20–30 transformations and expressions.
Preparation of electro-competent BL21(DE3) ∆serC cells
Day 1
Prepare LB/agar plate (no antibiotics) and cool to room temperature to solidify.
Streak out BL21(DE3) ∆serC cells onto LB/agar plate from a small glob of cells taken from a frozen glycerol stock.
Place LB/agar plate with streaked cells upside down in a static 37 °C incubator and grow overnight (14–20 h).
Day 2
After overnight growth, place LB/agar plate with BL21(DE3) ∆serC cells in a refrigerator until the end of the day.
Prepare and autoclave 1 L of 2× YT media in a 2.8 L baffled Fernbach flask.
Prepare 1 L of 10% (v/v) glycerol and sterilize by autoclaving along with the following:
i. 500 mL graduated cylinder.
ii. 2 × 0.5 L centrifuge bottles (or bottles suitable for centrifuging 1 L of culture at 4,000× g). Centrifuge bottles should be thoroughly cleaned to remove any residual DNA or cells that could contaminate electrocompetent cells prior to autoclaving.
iii. A box each of 1,000, 100, and 10 μL pipette tips.
iv. Approximately fifty 0.6 mL Eppendorf tubes.
After autoclaving, place the 10% (v/v) glycerol solution in a refrigerator or cold room to chill overnight.
At the end of the day, add 5 mL of 2× YT media to a 14 mL culture tube and inoculate with a single colony of BL21(DE3) ∆serC cells grown the prior night on the LB/agar plate. Grow liquid culture at 37 °C overnight with shaking at 250 rpm.
Day 3
Inoculate 1 L of 2× YT media (in a 2.8 L baffled Fernbach flask) with the 5 mL of overnight starter culture.
Add ~5–6 drops of anti-foam to the culture flask. Excessive foaming of the media caused by the flask baffles with inhibit air exchange and result in slower cell growth.
Grow the 1 L culture at 37 °C with shaking at 200–250 rpm until OD600 ~0.4. This should take approximately 2–3 h. Do not let culture grow above OD600 ~0.5. It is a good idea to check the OD600 approximately 1.5 h after inoculation to verify if cells are growing.
While cells are growing, pre-chill centrifuge and rotor to 4 °C and place centrifuge bottle(s) and graduated cylinder in refrigerator to chill as well.
Once cells reach OD600 ~0.4, immediately place culture flask in an ice/water bath for 15 min. Mix culture occasionally to ensure efficient cooling.
Note: From this point forward, work quickly and keep cells ice-cold at all times. Never let cells warm and work in a cold room if necessary. Maintain sterility.
Pour chilled culture into two sterilized 0.5 L centrifuge bottles and centrifuge cells at 4,000× g for 15 min at 4 °C.
Pour supernatant off without disturbing cell pellet. Wipe any residual media supernatant around the lip of the centrifuge bottle if necessary. Place centrifuge bottle with cell pellet immediately on ice.
Resuspend each cell pellet with 250 mL each of ice-cold 10% (v/v) glycerol solution (can be measured with the sterile graduated cylinder) by gentle swirling on ice. Be gentle with the cells, and DO NOT VORTEX. Keep cells cold at all times.
Once cells are evenly resuspended in 10% (v/v) glycerol, combine into one 0.5 L centrifuge bottle and centrifuge again at 4,000× g for 15 min at 4 °C.
After centrifuging, pour supernatant off without disturbing cell pellet. Place centrifuge bottle with cell pellet on ice.
Measure 250 mL of 10% (v/v) ice-cold glycerol in the same pre-chilled graduated cylinder. Resuspend cell pellet with this 250 mL of ice-cold glycerol solution by gentle swirling on ice. Again, DO NOT VORTEX and keep cells and centrifuge bottle cold at all times during the resuspension process.
Once cells are evenly resuspended in 10% (v/v) glycerol, centrifuge again at 4,000× g for 15 min at 4 °C.
Pour supernatant off without disturbing cell pellet. Place centrifuge bottle with cell pellet on ice.
Resuspend cell pellet with 30 mL of ice-cold glycerol solution by gentle swirling on ice. Again, DO NOT VORTEX and keep cells and centrifuge bottle cold at all times during the resuspension process. Transfer resuspended cells to a sterile 50 mL conical tube and top off conical tube to 50 mL total volume with ice cold 10% (v/v) glycerol.
Centrifuge again at 4,000× g for 15 min at 4 °C.
During this final centrifuge run, prepare a dry ice/ethanol bath as follows:
In a small Styrofoam box, a pan, or dish-like container, add dry ice pellets and pour enough ethanol (or isopropanol) to cover the bed of dry ice. Place an empty 1 mL pipette tip rack on the bed of dry ice; the base of the pipette rack should be touching the ethanol but should not be submerged, such that when you place a 0.6 mL Eppendorf tube in the rack, the bottom half of the Eppendorf tube is suspended in the ethanol bath while the cap of the tube stays well above the ethanol.
When the final centrifuge run is complete, pour off or aspirate the supernatant. Resuspend the cell pellet gently with ~0.3 mL of 10% (v/v) glycerol into an even suspension and transfer to a sterile 15 mL conical tube. To resuspend the cells, you can use a 1,000 μL pipettor equipped with a pipette tip that has had ~3 mm cut off the end with a sterile razor blade. The reason for cutting off the end of the pipette tip is to widen the hole at the end to minimize shearing forces on your cells as you are pipetting them up and down to resuspend.
Once resuspended into an even cell suspension, measure OD600.
Note: You will need make a ~1:500 dilution of the cells to get an accurate OD600 reading, e.g., 998 μL of water + 2 μL of cell suspension. Measure the OD600 of these diluted cells and then calculate OD600 of your actual cell suspension based on the dilution factor. For example, if your OD600 reading of your 1:500 dilution = 0.5, then the cell suspension is 0.5 × 500 = 250.
The OD600 of your resuspended cells should be between 200 and 300 to achieve sufficient competency for triple plasmid transformations and expressions. If the OD600 is above 300, dilute the cells with cold 10% glycerol so that the final density lies between 200 and 300. If the OD600 is below 200, then centrifuge the cells again and resuspend in a smaller volume of 10% (v/v) glycerol, e.g., 0.1 mL instead of 0.3 mL, re-measure OD600, and adjust volumes as needed.
Pipette ~35 μL of cell suspension into a 0.6 mL Eppendorf tube and quickly place tube in the dry-ice/ethanol bath rack to freeze cells.
The bottom of the Eppendorf tube should be submerged in the dry ice/ethanol bath, but do not let the cap of the tube come in contact with the ethanol; the alcohol will wick its way into the tube, ruining the cells.
Once cells are frozen (~2 min in the dry ice/ethanol bath), remove tube from bath, wipe the tube free of ethanol with a Kimwipe, and place the tube with frozen cells in a cryogenic freezer box containing dry ice.
Repeat until all cells have been aliquoted and frozen. Once all aliquots are frozen and in the box with dry ice, place box at -80 °C for long-term storage. You should obtain ~20–30 aliquots of competent cells from this procedure.
When done, place dry-ice/ethanol bath in a fume hood to allow dry ice to sublime. The ethanol/isopropanol can be poured into a bottle for re-use. Caution: the ethanol/isopropanol will off-gas CO2 for many hours, so if you pour it into a bottle be sure to leave cap lose/off to allow gas to escape, ensuring pressure does not build up inside the bottle.
Expression of protein with site-specifically encoded nhpSer
Fresh triple plasmid transformations must be performed for each expression. Never freeze expression cells containing plasmids; if expression cells are frozen with expression plasmids in them, the cells will grow in the presence of the correct antibiotics, but they will not express protein. Do not sequentially transform plasmids.
To standardize this protocol in your lab, first run control expressions with sfGFP wt and sfGFP-150TAG to verify you can achieve the reference expression benchmarks (see Table 1 below) and ensure that nhpSer encoding at the TAG site is functioning as expected with a model protein. The protocol below outlines the protocol for a standard 50 mL test expression to encode nhpSer into sfGFP at position N150 (i.e., sfGFP-150TAG), as well as wild-type sfGFP. Pointers on how to scale up to a 1 L expression, as is often used for target proteins, are also included.
Day 1: transformations
Prepare two LB/agar plates, one for sfGFP-wt and one for sfGFP-150TAG expressions, with antibiotics as follows:
i. Sterilize LB/agar as described above. After autoclaving, allow to cool sufficiently to touch while still remaining liquid.
ii. Pour 50 mL of LB/agar into a sterile 50 mL conical tube. Add 35 μL each of ampicillin, kanamycin, and spectinomycin stock solutions. Mix thoroughly and pour ~20–25 mL into each 100 mm plate. Final working concentrations of antibiotics should be 70 μg/mL ampicillin (for the pRBC plasmid), 35 μg/mL kanamycin (for the pERM2-nhpSer plasmid), and 70 μg/mL spectinomycin (for the pCDF-Frb-v1 plasmid).
iii. Allow LB/agar to cool and solidify beside a flame with plate lid slightly ajar for 30 min.
iv. Label the two plates accordingly, e.g., “sfGFP-WT” and “sfGFP-150TAG.”
Place two 1 mM electro-cuvettes on ice for 15 min to pre-chill.
Place two 0.6 mL Eppendorf tubes on ice and label, e.g., “sfGFP-WT” and “sfGFP-150TAG.”
i. To the “sfGFP-WT” tube, add 2 μL each of pRBC-sfGFP WT, pERM2-nhpSer, and pCDF-Frb-v1 plasmids (~100–500 ng of each plasmid).
ii. To the “sfGFP-150TAG” tube, add 2 μL each of pRBC-sfGFP-150TAG, pERM2-nhpSer, and pCDF-Frb-v1 plasmids.
iii. Allow tubes to chill on ice for 5–10 min.
Thaw two aliquots of electrocompetent BL21(DE3) ∆serC cells. Cells can be thawed rapidly with the warmth of your fingers, but immediately place tube on ice once thawed.
Transfer 30 μL of electrocompetent BL21(DE3) ∆serC cells to each tube with plasmids and gently mix cells with plasmids by pipetting up and down 2–3 times. Cells should not appear stringy or behave like “snot,” which would indicate cells have lysed.
Transfer the “sfGFP-WT” electro-competent cell/plasmid mix into a pre-chilled electro-cuvette. Wipe metal electrode plates dry with a Kimwipe. Gently flick the cuvette to ensure cells are sitting in the bottom and stretch the full width of the cuvette.
Place electro-cuvette into the electroporator and electroporate according to instrument instructions for 1 mM cuvettes (e.g., 1,500 V for the Eppendorf Eporator).
Note: The time constant should be 4–6 ms. If arc’ing occurs, discard cells and cuvette. Arc’ing occurs when DNA and/or cells have too much salt. Re-attempt electroporation by diluting a fresh cell/plasmid mix with equal volume of cold MQ water.
Immediately add 1 mL of SOC media to electro-cuvette and resuspend cells. Transfer cells to a sterile 1.7 mL Eppendorf tube.
Repeat steps f–h for the “sfGFP-150TAG” electro-competent cell/plasmid mix.
Place both Eppendorf tubes in a shaker (250 rpm) and recover cells at 37 °C for 90 min.
Note: Electro-cuvettes can be reused at least 10 times if washed shortly after use. Wash cuvettes as follows while cells are recovering:
i. MQ H2O.
ii. 0.1 M HCl (dilute acid helps to break down residual DNA in the cuvette).
iii. MQ H2O.
iv. 70% (v/v) ethanol.
v. Let dry upside down on a Kimwipe. Once dry, store with cap to maintain sterility.
After 90 min of recovery, plate all the cells from the “sfGFP-WT” recovery culture onto the LB/agar/Amp/Kan/Spec plate labeled “sfGFP-WT.”
To plate all the transformed cells, you cannot put the entire 1 mL of recovery culture on the agar plate as this is too much liquid and it will not dry properly. Instead, pellet the cells by centrifugation at 3,000× g for 3 min. Remove the top 900 μL of media and gently resuspend the pellet in the remaining ~100 μL. Spread this cell suspension evenly onto the agar plate.
Repeat plating process for the “sfGFP-150TAG” culture.
Let plates dry with lid partially open for ~20 min near a flame (maintaining sterility) and then incubate plates upside down overnight (~16 h) at 37 °C.
Day 2: expressions
This protocol is intentionally written to avoid overnight liquid starter cultures that would reach stationary phase. Rather, in one day, you will use freshly grown colonies on an agar plate to grow a starter liquid culture in the morning, after which you will inoculate expression culture in the early afternoon. By the end of the day the cultures will be induced and allowed to express for the next 20–24 h. Having a large number (> 500) of colonies on each LB/agar plate is necessary to execute this one-day expression protocol.
All liquid cultures must contain 70 μg/mL ampicillin, 35 μg/mL kanamycin, and 70 μg/mL spectinomycin.
Important: Baffled culture flasks are critical to ensure adequate aeration of expression cultures.
Remove LB/agar plates from 37 °C incubator. There should be several hundred or thousands of colonies (examples of successful triple plasmid transformations are shown in Figure 3).
Figure 3. Representative LB/agar plates after triple plasmid co-transformation of pRBC-sfGFP WT (left) or pRBC-sfGFP-150TAG (right) with pERM2-nhpSer and pCDF-Frb-v1 plasmids and growth at 37 °C overnight (~18 h). The sfGFP-WT plates on the left are fluorescent due to leaky expression of the T7 expression system in BL21(DE3) cells. Note the size homogeneity of colonies and that all colonies on the pRBC-sfGFP WT plate are green and no green colonies appear on the pRBC-sfGFP 150TAG plate.
Prepare starter cultures
i. Starter cultures are grown in starter culture media as defined above (2× YT + 0.5% (v/v) glycerol) with the same antibiotics and concentrations used in the LB/agar plates.
ii. For small-scale (e.g., 50 mL) sfGFP control expressions, prepare two starter cultures by adding 5 mL of starter culture media (with ampicillin, kanamycin, and spectinomycin antibiotics) to 14 mL sterile culture tubes.
iii. To inoculate these 5 mL cultures, scrape a “glob” of cells constituting several dozen to hundreds of colonies from overnight LB/agar plate with a sterile pipette tip, shake the glob off into the culture media, and break apart by gentle pipetting. Enough cells should be transferred to the 5 mL starter culture such that it is visibly turbid upon inoculation.
iv. For larger scale expressions (e.g., > 1 L), prepare 50 mL of starter culture media (with antibiotics) in a baffled 250 mL culture flask. Transfer 5 mL of starter culture media onto the LB/agar plate containing transformed cells. Gently scrape all colonies with a sterile glass spreader to resuspend them into the liquid media. Transfer media with suspended cells into the 250 mL baffled culture flask containing ~45 mL of starter culture media. Add 1–2 drops of antifoam.
Grow starter cultures at 37 °C with shaking at 250 rpm for ~3–4 h until OD600 > 1. Do not grow overnight.
While starter cultures are growing, prepare expression media:
i. For each 50 mL sfGFP test expression, add 5 mL of (room temperature) sterile 10× potassium phosphate buffer to 45 mL of 1.1× TB media in a 250 mL baffled culture flask. Add antibiotics and ~1–2 drops of antifoam.
ii. For 1 L expressions, add 100 mL of sterile 10× potassium phosphate buffer to 900 mL of 1.1× TB media in a 2.8 L baffled Fernbach flask. Add antibiotics and ~5–6 drops of antifoam.
After ~3–4 h of starter culture growth, inoculate expression media as follows:
i. For 50 mL of sfGFP test expressions, add 1 mL of the sfGFP-WT starter culture to 50 mL of TB expression media (in a baffled 250 mL culture flask). Repeat for the sfGFP-150TAG starter culture.
ii. For a 1 L expression, add 10–20 mL of starter culture to each baffled Fernbach flask containing 1 L of TB expression media.
Grow expression cultures at 37 °C with shaking (200–250 rpm) until OD600 = 1.0. This should take approximately 3–5 h.
When cultures reach OD600 = 1, add IPTG to a final concentration of 0.5 mM.
i. For 50 mL cultures, this corresponds to 50 μL of 0.5 M IPTG stock solution.
ii. For 1 L cultures, this corresponds to 1 mL of 0.5 M IPTG stock solution.
Reduce temperature to 30 °C for sfGFP control expression cultures. For other target proteins, temperature can be reduced to as low as 20 °C. The Frb biosynthetic pathway functions optimally between 20 and 30 °C. Expression temperatures below 20 °C may be possible but expression times will need to be extended and overall yields will be diminished. Do not express target proteins above 30 °C since the Frb biosynthetic pathway does not function above this temperature.
For 50 mL expression cultures, add 1–2 more drops of antifoam. For 1 L expressions, add 5–6 more drops of antifoam.
Continue shaking cultures at 200–250 rpm for 20–24 h at the desired temperature (between 20 and 30 °C; optimal for sfGFP is 30 °C). During this expression period, the density of the culture (OD600) should increase significantly (up to 20).
Day 3: Evaluating expression results and harvesting cells
sfGFP control protein expression analysis
Approximately 20–24 h after induction, measure OD600 and fluorescence of both sfGFP and sfGFP-150TAG cultures using a fluorometer. Since sfGFP chromophore formation requires synthesis of full-length protein, fluorescence of whole cells provides a convenient strategy to evaluate the efficiency of sfGFP TAG codon suppression, and therefore nhpSer incorporation. Fluorescence can be measured with any fluorimeter capable of detecting sfGFP fluorescence (ex/em: 488/510 nm). Diluting the culture 1:10 to 1:100 in a buffer (e.g., 100 μL of culture + 1,900 μL of PBS for a 1:20 dilution) prior to fluorescence measurements may be necessary to obtain a signal within the linear range of the fluorometer.
Test expression benchmarks:
In general, the sfGFP-150TAG culture fluorescence should be approximately 20%–30% that of the sfGFP wild-type culture, corresponding to 50–150 mg of sfGFP-150nhpSer per liter of culture (Table 1).
The OD600 values will vary depending on target protein. Normal values will range from ~10 to 20 for sfGFP expressions. Final OD600 values below 5 are indicative of poor cell growth or toxicity due to target protein expression.
Table 1. Benchmarks for sfGFP-WT and 150TAG protein expression yields, based on culture fluorescence
Fluorescence of culture# mg of sfGFP per liter culture* OD600
wt sfGFP 8,030 (6,000–12,000) 458 (340–600) 10.0 (5–15)
sfGFP 150TAG
(with Frb-v1 pathway)
2,550 (1,500–3,000) 120 (50–150) 17.0 (10–20)
#The range indicated in parenthesis is considered normal depending on the day and reagent preparation. Fluorescence values reported here were obtained on a hand-held PicoFluor fluorometer (Turner Biosystems) by diluting cells directly from the culture into PBS (1:20). Fluorescence values are arbitrary and will depend on the fluorometer used. It is important that the relative ratio of sfGFP-150TAG and sfGFP-WT culture fluorescence is consistent with the above values. For reference, background cultures not expressing sfGFP have auto-fluorescence values of ~150–250.
*Yield of sfGFP in milligrams per liter was determined using a fluorescence standard curve of purified sfGFP.
Harvesting cells:
Harvest cells by centrifugation at 5,000× g for 15 min at 4 °C. Pour off culture supernatant and resuspend cells in an appropriate buffer.
The choice of buffer depends on the protein of interest, the downstream purification strategy, and the application, and should be determined by the user.
Adding a cryoprotectant [e.g., 10% (v/v) glycerol] to this buffer can help minimize adverse effects associated with freezing sensitive or unstable proteins. Cells can be flash frozen in liquid nitrogen and stored at -80 °C or you can proceed with purification.
For His6-tagged proteins to be purified via TALON resin, a recommended resuspension/lysis buffer would be 50 mM Tris pH 7.5, 500 mM NaCl, 10% (v/v) glycerol, 5 mM imidazole. Phosphatase inhibitors are not required since nhpSer is not hydrolyzed by phosphatases.
Validation of protocol
For each expression, it is important to confirm faithful incorporation of nhpSer into the target protein. Several methods can be used to evaluate the phosphorylation status of the target protein.
Phos-tag electrophoresis: Phos-tag gel electrophoresis is a modified form of SDS-PAGE, in which phosphorylated proteins migrate with attenuated electrophoretic mobility compared to their non-phosphorylated counterparts. The degree to which protein migration is attenuated increases with sequential addition of phosphoryl groups, allowing one to distinguish between no, single-, and multi-pSer containing proteins (Kinoshita et al., 2006). Serendipitously, proteins with nhpSer migrate slower on Phos-tag electrophoresis than the same proteins with authentic pSer, allowing easy discrimination between the two [for examples, please see (Zhu et al., 2023) as well as Figure 4 below]. By measuring the density of the non-phosphorylated vs. pSer vs. nhpSer protein bands, one can estimate the percentage of expressed/purified protein that contains nhpSer. Important to note is that wild-type (non-phosphorylated) protein must be run side-by-side with the phosphorylated protein to observe relative shifts in electrophoretic mobility. Similarly, it is also helpful to express your target proteins with pSer as well so that you can confirm if the electrophoretic shifts of your nhpSer-containing protein are due to nhpSer encoding and not pSer [see our previous protocol for expressing proteins with pSer: (Zhu et al., 2022)]. Other advantages of Phos-tag electrophoresis include the ability to evaluate multiple samples at once, requiring only a standard SDS-PAGE electrophoresis setup, and being economical. Do not run molecular weight markers on Phos-tag gels, as they often contain EDTA, which is not compatible with Phos-tag. See Figure 4 below for an example of SDS-PAGE and Phos-tag gels of nhpSer and pSer containing sfGFP.
Note: The attenuated electrophoretic mobility of sfGFP-150pser/nhpSer proteins in Phos-tag gels are particularly pronounced and easily discernable; however, the degree to which other phosphorylated proteins migrate slower than their non-phosphorylated counterparts depends on the protein and the site of encoding. If you do not see a convincing electrophoretic shift between your phosphorylated and non-phosphorylated proteins, consider optimizing the Phos-tag electrophoresis procedure to improve resolving power by (i) increasing concentration of the Phos-tag acrylamide reagent from 25 or 50 μM to 100 μM or higher, (ii) decreasing the amount of protein loaded for narrower bands and enhanced resolution between similarly migrating protein bands, (iii) decrease the total acrylamide concentration [e.g., from 15% to 10% (w/v) acrylamide] so that target proteins bands travel at least half-way through the gel thereby increasing separation, and (iv) run sfGFP WT/150nhpSer control proteins alongside the proteins of interest to ensure the Phos-tag gel is run correctly. If still no shift is observed on Phos-tag electrophoresis, you will have to confirm incorporation via mass spectrometry (see below).
Figure 4. Assessing nhpSer encoding into sfGFP by Phos-tag gel electrophoresis. SDS-PAGE (top) and Phos-tag (bottom) gels of purified WT sfGFP, sfGFP-150pSer, and sfGFP-150nhpSer proteins. The sfGFP-150pSer protein was produced following our prior protocol (Zhu et al., 2022) and the sfGFP-150nhpSer protein was produced using the protocol described here. Because they migrate differently on Phos-tag gels, non-phosphorylated, pSer-containing, and nhpSer-containing proteins can be resolved from one another, and the fraction of each form can be easily evaluated in each sample. In this example, the sfGFP-150pSer protein sample is > 90% phosphorylated with pSer, with only trace amounts of non-phosphorylated protein that migrates identically as the wild-type sfGFP protein, while the sfGFP-150nhpSer protein is > 95% phosphorylated with nhpSer, with only trace amounts of pSer-containing contaminant protein and no detectable non-phosphorylated protein. Note that all proteins migrate identically on SDS-PAGE, indicating the electrophoretic shifts observed in Phos-tag are due to phosphorylation status and not differences in protein size. These samples were incubated at 95 °C for 5 min in standard Laemmli buffer prior to their loading on gel. The Phos-tag gel contained 50 μM Phos-tag acrylamide, 100 μM MnCl2, and a 29:1 acrylamide:bis-acrylamide ratio and was run at 120 V for 1.5 h at room temperature.
Mass spectrometry: whole-protein mass spectrometry (MS) is the most convincing methodology for confirming nhpSer encoding, if facilities are available and the target protein is amenable to whole-protein MS analysis. Note that some mass spectrometers may not have sufficient resolution to distinguish masses of whole proteins with pSer and nhpSer (∆ = 1.98 Da), in which case Phos-tag electrophoresis or tryptic-digests followed by MS/MS fragmentation (i.e., bottom-up analysis) can be used. For an example of high-resolution whole-protein MS data distinguishing between sfGFP with pSer and nhpSer, please see Figure 3 in Zhu et al. (2023). When analyzing whole-protein MS data of sfGFP proteins, be sure to account for chromophore maturation, which results in the loss of a water molecule and two protons; therefore, the mass will be exactly 20 Da less than that predicted by amino acid sequence.
Important note: Tryptic-digest followed by MS fragmentation analysis (i.e., “bottom-up” MS/MS analysis) of nhpSer-proteins is useful for confirming nhpSer incorporation and the site of incorporation, but it should not be used to make conclusions regarding the degree of nhpSer encoding.
Western blotting: a variety of antibodies are available to detect specific pSer-containing proteins via western blotting. We have not yet tested if such pSer-specific antibodies also cross-react with the same proteins containing nhpSer but expect this to be dependent on the protein, site of phosphorylation, and the antibody. Note that while western blotting with a phosphorylation-sensitive antibody may be useful to confirm target protein phosphorylation, it cannot be used to evaluate the extent of phosphorylation without running alongside a standard curve of homogenously phosphorylated protein standard. Alternatively, it is possible to image Phos-tag gels via western blotting to evaluate the degree of wild-type, pSer, and nhpSer-containing target protein in a mixture of other proteins [e.g., see Supporting Information Figure S26 in Zhu et al. (2023)]. In these cases, using an antibody that binds to its epitope irrespective of protein phosphorylation status is important (e.g., FLAG or MYC tags).
General notes and troubleshooting
General notes
Truncation. Like most standard E. coli expression hosts, the BL21(DE3) ∆serC strains contains Release Factor-1 (RF1), the protein responsible for terminating translation at TAG codons. Consequently, nhpSer encoding at TAG codons is in constant competition with RF1 and so a mixture of full-length protein containing nhpSer and prematurely truncated protein will be produced. Keeping this in mind when designing the expression construct of the target protein is important. Appending a C-terminal purification tag ensures one purifies only full-length protein, though not all proteins express well with C-terminal tags. If an N-terminal purification tag is used, the truncated protein may likely co-purify with full-length protein, and could be the dominant purified protein species. In these cases, you will need to develop additional strategies to separate the truncated protein. Note that if the target protein is a homo-oligomer, truncated protein may be difficult or impossible to purify away when the site of encoding is C-terminal to the oligomerization domain, as both truncated and full-length proteins will oligomerize and co-purify regardless of where the purification tag is placed.
Compatible plasmids for expressing target protein. In this protocol, three plasmids must be present in the cell in order to encode nhpSer into a target protein. For multiple plasmids to be faithfully maintained in E. coli, each plasmid must have an origin-of-replication from a different compatibility group, as well as confer different antibiotic resistances. We engineered the pRBC vector (ampicillin resistance, p15a origin-of-replication) to meet these requirements when being propagated with the nhpSer machinery vector (pERM2-nhpSer, kanamycin resistance with pUC origin-of-replication) and the nhpSer biosynthetic pathway vector (pCDF-Frb-v1, spectinomycin resistance with CloDF origin-of-replication). Critical to note is that because the pERM2-nhpSer vector harbors a pUC origin of replication, you cannot use traditional pET, pGEX, or pBAD-like vectors to express target proteins because these vectors harbor an origin-of-replication from the same compatibility group as the pERM2-nhpSer vector (the pUC origin of replication is a derivative of pBR322/pMB1 origins and is therefore in the same compatibility group). The pRBC vector was also engineered to be compatible with established pSer GCE encoding systems, which use the pKW2-EFSep machinery plasmid (chloramphenicol resistance and pBR322 origin-of-replication) enabling the flexibility to encode either pSer or nhpSer with the same pRBC vector depending on downstream applications. If you wish to use a vector other than pRBC to express target proteins with pSer or nhpSer, ensure that it has a p15a or RSF origin-of-replication and that it confers resistance to an antibiotic other than kanamycin, spectinomycin, and chloramphenicol (e.g., ampicillin, tetracycline).
Troubleshooting
Construct design. There are a variety of reasons why a target protein may not express when encoding nhpSer (or pSer), even though your sfGFP-150TAG controls are working well and your equivalent wild-type target protein construct expresses at high yields. In these situations, it is important to consider that phosphorylation is a post-translational modification, whereas with GCE systems the phospho-amino acids are installed during translation, and we therefore require the target protein to fold properly with the phospho-amino acid present and while isolated from any potential interacting/stabilizing partner proteins. So, even when using highly efficient pSer/nhpSer GCE systems to encode phospho-amino acids into otherwise well-behaved, highly expressing and soluble native proteins, there may be difficult-to-anticipate challenges that will require clever strategies to coax target phospho-proteins to express stably and efficiently. Strategies to improve phospho-protein expression, folding and stability include (i) fusing solubility tags to the N- and/or C-terminus of the target protein (e.g., SUMO, GST, MBP), (ii) co-expressing folding chaperone proteins (e.g., GroEL/ES, Trigger Factor), and (iii) co-expressing binding partner proteins, if they are known. Similarly, fusing a cleavable fluorescent reporter protein (e.g., super-folder GFP) to the C-terminus of your target protein provides a convenient strategy to screen for optimal expression conditions by measuring cell fluorescence. In this case, only when nhpSer is encoded and full-length target protein is produced will translation continue into the sfGFP causing the cells to fluoresce. Including a cleavage site between target protein and sfGFP (e.g., TEV, 3C) allows the sfGFP to be removed should it interfere with function or downstream applications.
Transformations. This protocol requires three plasmids to be transformed into E. coli cells simultaneously for every expression, and with an efficiency that allows you to obtain enough cells to inoculate a starter culture that only takes ~1/2 a day to grow to a density sufficient to inoculate expression cultures. If only a few colonies are obtained after the transformation, you will not have enough cells to inoculate your starter culture, and then this starter culture will take too long to grow. Overnight growth of the starter culture would seem an obvious strategy to overcome this issue; however, we have observed poor expression of nhpSer proteins when starter cultures are allowed to reach stationary phase. Protein also will not express if expression cells containing plasmids were grown from a frozen glycerol stock as opposed to from freshly transformed cells. If you are experiencing low transformation efficiencies, this may be caused by (i) poorly prepared electrocompetent cells, (ii) too low plasmid concentrations, (iii) degraded, compromised, or salty plasmids (see note below) and/or (iv) improper electroporator parameters. To troubleshoot the source of the inefficiency, transform each plasmid individually and plate on LB/agar plates with a single antibiotic (ampicillin, kanamycin, and spectinomycin). You should obtain a “lawn” of cells for each individual plasmid transformation. If one of the plasmids transforms notably worse than another, there may be integrity issues associated with that particular plasmid. If you get few colonies on all three single plasmid transformations, either the electrocompetent cells were prepared incorrectly or the electroporator is not functioning properly.
Plasmid integrity. We have observed in rare cases the pERM2-nhpSer plasmid recombining when propagated in E. coli cells to render it non-functional. We recommend sequencing the plasmid when you receive it from Addgene and after propagation to ensure integrity. Economical, whole-plasmid sequencing is available from a variety of companies, such as Plasmidsaurus. Sequencing the pCDF-Frb-v1 and pRBC-sfGFP-150TAG plasmids are also recommended, though we have not observed any stability concerns with these plasmids when propagated in DH10b E. coli cells.
Acknowledgments
The original research paper in which this protocol was described and validated is Zhu et al. (2023). This work was supported by the GCE4All Biomedical Technology Development and Dissemination Center supported by National Institute of General Medical Science grant RM1-GM144227. This work was also supported in part by the National Institute of Health [5R01GM131168-02 to R.A.M.], the Medical Research Foundation at Oregon Health Sciences University [to R.B.C.], and the Collins Medical Trust [to R.B.C.].
Competing interests
The authors declare no competing interests.
Ethical considerations
No human or animal use is described in this protocol.
References
Kinoshita, E., Kinoshita-Kikuta, E., Takiyama, K. and Koike, T. (2006). Phosphate-binding Tag, a New Tool to Visualize Phosphorylated Proteins. Mol. Cell. Proteomics 5(4): 749–757.
Rogerson, D. T., Sachdeva, A., Wang, K., Haq, T., Kazlauskaite, A., Hancock, S. M., Huguenin-Dezot, N., Muqit, M. M. K., Fry, A. M., Bayliss, R., et al. (2015). Efficient genetic encoding of phosphoserine and its nonhydrolyzable analog. Nat. Chem. Biol. 11(7): 496–503.
Zhang, M. S., Brunner, S. F., Huguenin-Dezot, N., Liang, A. D., Schmied, W. H., Rogerson, D. T. and Chin, J. W. (2017). Biosynthesis and genetic encoding of phosphothreonine through parallel selection and deep sequencing. Nat. Methods 14(7): 729–736.
Zhu, P., Gafken, P. R., Mehl, R. A. and Cooley, R. B. (2019). A Highly Versatile Expression System for the Production of Multiply Phosphorylated Proteins. ACS Chem. Biol. 14(7): 1564–1572.
Zhu, P., Mehl, R. and Cooley, R. (2022). Site-specific Incorporation of Phosphoserine into Recombinant Proteins in Escherichia coli. Bio Protoc 12(21): e4541.
Zhu, P., Stanisheuski, S., Franklin, R., Vogel, A., Vesely, C. H., Reardon, P., Sluchanko, N. N., Beckman, J. S., Karplus, P. A., Mehl, R. A., et al. (2023). Autonomous Synthesis of Functional, Permanently Phosphorylated Proteins for Defining the Interactome of Monomeric 14-3-3ζ. ACS Cent. Sci. 9(4): 816–835.
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Workflow for High-throughput Screening of Enzyme Mutant Libraries Using Matrix-assisted Laser Desorption/Ionization Mass Spectrometry Analysis of Escherichia coli Colonies
KC Kisurb Choe
JS Jonathan V. Sweedler
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4862 Views: 653
Reviewed by: Neha NandwaniRama Reddy GoluguriAmit Kumar Dey
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Original Research Article:
The authors used this protocol in Metabolic Engineering May 2023
Abstract
High-throughput molecular screening of microbial colonies and DNA libraries are critical procedures that enable applications such as directed evolution, functional genomics, microbial identification, and creation of engineered microbial strains to produce high-value molecules. A promising chemical screening approach is the measurement of products directly from microbial colonies via optically guided matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS). Measuring the compounds from microbial colonies bypasses liquid culture with a screen that takes approximately 5 s per sample. We describe a protocol combining a dedicated informatics pipeline and sample preparation method that can prepare up to 3,000 colonies in under 3 h. The screening protocol starts from colonies grown on Petri dishes and then transferred onto MALDI plates via imprinting. The target plate with the colonies is imaged by a flatbed scanner and the colonies are located via custom software. The target plate is coated with MALDI matrix, MALDI-MS analyzes the colony locations, and data analysis enables the determination of colonies with the desired biochemical properties. This workflow screens thousands of colonies per day without requiring additional automation. The wide chemical coverage and the high sensitivity of MALDI-MS enable diverse screening projects such as modifying enzymes and functional genomics surveys of gene activation/inhibition libraries.
Key features
• Mass spectrometry analyzes a range of compounds from E. coli colonies as a proxy for liquid culture testing enzyme mutant libraries.
• Colonies are transferred to a MALDI target plate by a simple imprinting method.
• The screen compares the ratio among several products or searches for the qualitative presence of specific compounds.
• The protocol requires a MALDI mass spectrometer.
Graphical overview
Overview of the MALDI-MS analysis of microbial colonies for screening mutant libraries. Microbial cells containing a mutant library for enzymes/metabolic pathways are first grown in agar. The colonies are then imprinted onto a MALDI target plate using a filter paper intermediate. An optical image of the MALDI target plate is analyzed by custom software to find the locations of individual colonies and direct subsequent MALDI-MS analyses to the selected colonies. After applying MALDI matrix onto the target plate, MALDI-MS analysis of the colonies is performed. Colonies showing the desired product profiles are found by data analysis via the software, and the colonies are picked for downstream analysis.
Keywords: Screening Protein engineering Mass spectrometry Microbial colonies MALDI-MS
Background
Directed evolution is an iterative process of mutagenesis and phenotype screening to achieve incremental improvements towards desired characteristics (Arnold, 2018). The method is used widely for creating efficient biocatalysts producing drugs, fuels, food, and industrial products (Turner, 2009; Dietrich et al., 2010; Wang et al., 2021). Lack of high-throughput methods for phenotype screening can be a bottleneck for directed evolution (Zeymer and Hilvert, 2018). Chromogenic conversion, pH change, and fluorescence readings can be used for phenotype screening; however, they are often indirect measurements of the desired phenotype (Körfer et al., 2018; Alfaro-Chávez et al., 2019; Minges et al., 2020). A promising approach is mass spectrometry (MS), which provides label-free measurement of a wide variety of analytes. Acoustic ejection mass spectrometry is the state-of-the-art high-throughput screening technology that applies an acoustic pulse to transfer liquid samples to an MS device, achieving a subsecond analytical cycle (Liu et al., 2020; Zhang et al., 2021). However, it requires extensive automation systems for sample creation such as biochemical reactions, cell culture, liquid transfer, and storage (Simon et al., 2021). Such robotic systems are not commonly available to many labs. Alternatively, matrix-assisted laser desorption/ionization MS (MALDI-MS) is a widely available MS system that boasts high throughput, relatively simple sample preparation, high salt tolerance, and wide chemical coverage (Fournier et al., 2008; Grove et al., 2011; Singhal et al., 2015). Previously, the optically guided MALDI-MS method was applied to perform high-throughput screening of microbial colonies for engineering multistep enzyme reactions (Si et al., 2017). In this workflow, hundreds of agar colonies are transferred to a MALDI plate by imprinting, and MALDI-MS of the scattered colonies is performed using the coordinates reported by microMS image analysis software. This work successfully enables individual colonies to serve as individual reaction vessels for phenotyping, which is a promising alternative to the complex automation system needed for performing and processing large numbers of liquid cultures. For routine analysis of a larger number of microbial colonies, the current protocol simplifies steps for imaging, instrument setup, software operation, and downstream data analysis.
The simplification of the workflow is enabled by macroMS, a newly created software package that is freely available. The macroMS is the adaptation of a single-cell microMS package and it was created for high-throughput screening of macroscopic samples larger than 200 μm (Comi et al., 2017; Choe et al., 2021). The macroMS package enables rapid imaging by flatbed scanner and camera, user-friendly tools for image analysis and optical correction, simple MS instrument training, and data analysis/visualization for isolating colonies of interest. Based on macroMS, this protocol describes a simplified end-to-end colony screening workflow by MALDI-MS, including growing and preparing colonies for the MALDI-MS screen, using the macroMS software to set up and perform MALDI-MS analysis, performing data analysis, and picking colonies of interest. This workflow can be used to efficiently screen mutant libraries for modifying a wide variety of enzymes or metabolic pathways that form compounds on colonies that are detectable by MALDI-MS. While the workflow has been developed using the Bruker ultraflextreme mass spectrometer, it can be adapted to other instruments, although this would require adapting several of the software and instrument operation steps. The limitations of the workflow are: 1) it works with nonvolatile compounds that do not evaporate in the vacuum environment of MALDI-MS, 2) it is a semiquantitative MALDI-MS screen that uses the ratio between observed MALDI-MS peaks, and 3) the macroMS data analysis tool limits the width of the mass range of the mass spectrum to be under m/z 1,000.
As the equipment to spray matrix onto the slides is not commonly available in many laboratories, the end of the protocol includes the construction of a spraying device for applying MALDI matrix onto four extra-large MALDI target plates (i.e., 105 mm × 75 mm). MALDI matrix application is a commonly performed step for MALDI imaging, where airbrush, sublimation chamber, and commercial automatic sprayer systems have been used to coat samples with the compounds (Hankin et al., 2007; Ye et al., 2013; Andersen et al., 2020). However, few devices have the capacity for processing wide target plates holding the large number of colonies needed for high-throughput screening. Also, while MALDI-MS has become commonly available in universities and industrial settings, facilities often lack the capability and experience with MALDI imaging and therefore do not have devices for MALDI matrix coating. If other alternatives are available, they can be used; otherwise, the system described here provides the capability to simultaneously prepare four large MALDI plates as needed for high-throughput screens and reducing variability.
The workflow was successfully used to screen mutant libraries for an acyl-ACP thioesterase and an acyl-ACP desaturase. Acyl-ACP thioesterase cleaves the thioester bond in acyl-ACPs in the type II fatty acid synthesis pathway, deciding the chain length of the resulting free fatty acids. Acyl-ACP thioesterase from the California bay laurel (UcFatB2) expressed in E. coli produces dodecanoic acid as a primary product and tetradecanoic acid as a secondary product. The workflow compared the ratio between dodecanoic acid and tetradecanoic acid measured from colonies that express UcFatB2 random mutant library and found variants that have higher ratio for dodecanoic acid (Jindra et al., 2023). Acyl-ACP desaturase removes two hydrogens from a single bond between two carbons (C — C) resulting in double bond (C = C) in fatty acyl-ACP, deciding the location of the double bond in unsaturated fatty acid. Acyl-ACP desaturase from the Black-eyed Susan vine (TaDes) expressed in E. coli produces C16:1 Δ6 as a primary product and C16:1 Δ8 acid as a secondary product. By combining with a simple ozone treatment step, the workflow compared the ratio between the ozonolysis products of C16:1 Δ6 and C16:1 Δ8 measured from colonies that express TaDes random mutant library. The screen found variants that produce C16:1 Δ8 as a primary product (Choe et al., 2023). The protocol covers the workflow demonstrated using the thioesterase screen.
Materials and reagents
Prespotted AnchorChip adapter (Bruker Daltonics, catalog number: 221598), four custom built 1.2 mm × 105 mm × 75 mm target plates, and copper conducting tape (3M, catalog number: 05012A-AB) (Note 1)
MTP target frame (Bruker Daltonics, catalog number: 8074115) and MTP 384 target plate polished steel (Bruker Daltonics, catalog number: 8280781)
Vacuum desiccator (SP Bel-Art, catalog number: F42020-0000)
5′ of 1/2″ tubing (McMaster-Carr, catalog number: 5648K33)
N-Phenyl-2-naphthylamine (PNA) (Sigma-Aldrich, catalog number: 178055)
7.25″ × 5″ × 1.875″ HDX hydrophilic sponge (QEP, catalog number: 70005-24)
6″ × 6″ × 3/8″ metal plate (McMaster-Carr, catalog number: 8983K212)
6″ × 6″ × 0.12″ metal plate (McMaster-Carr, catalog number: 8983K118)
A4 papers
Standard lab tape
Lighter
Push pins
Duct tape
Sharpie pen
Power strip
Isopropyl alcohol
Methanol
Glycerol
Delicate task wipes (Kimberly-Clark Professional, catalog number: 34155)
10% bleach
150 mm × 15 mm bacteriological Petri dish (Corning, catalog number: 351058)
Syringe filter, 0.2 μm Nylon (Thermo Scientific, catalog number: 726-2520)
90 mm filter unit, 500 mL (Thermo Scientific, catalog number: 569-0020)
Sterile 10 mL syringe, single use (BD, catalog number: 302995)
Sterile 1.5, 15, and 50 mL tubes and corresponding tube racks
Sterile 2, 20, and 1,000 μL pipette tips and corresponding adjustable volume pipettes
Sterile 5 and 50 mL serological pipettes and a pipetaid
Carbenicillin (Gold Biotechnology, catalog number: C-103-5)
Autoclaved 90 mm membrane filter, 0.22 μm (Millipore, catalog number: GVWP09050)
14 mL cell culture tubes (Greiner Bio-One, catalog number: 187262)
Sterile 250 and 1,000 mL autoclavable flasks (Pyrex, catalog number: 4980)
L-shaped cell spreader (VWR, catalog number: 76207-748)
Inoculating turntable
Ice bucket
4″ × 125′ Parafilm-M film (Bemis Company Inc, catalog number: PM996)
Gallon storage double zip bags, 26.8 cm × 27.3 cm (Walmart)
Plastic box for transporting MALDI target adapter
Aluminum foil
Tweezer
Spectrophotometer cuvette
10 nm gold colloid solution (BBI Solutions, catalog number: EM.GC10/4)
LB powder (Fisher Scientific, catalog number: BP9722-2)
Agar (BD, catalog number: 214030)
Isopropyl β-D-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich, catalog number: IPTG-RO)
Glucose (Fisher Scientific, catalog number: D1610)
Autoclave tapes
Clock
Paper towel
Miniprep kit (QIAGEN, catalog number: 27104)
Chemical waste bottle
Cuffed sleeve lab coat
Disposable laboratory exam gloves
Biohazard trash can
Electrocompetent cells (i.e., RL08ara strain)
Mutant library plasmid (i.e., pTrc99a-*BTE)
Wild-type control plasmid (i.e., pTrc99a-WT BTE)
Acetonitrile (Thermo Fisher Scientific, catalog number: 51101) (Note 2)
Acetone (Fisher Scientific, catalog number: A929-1)
LB agar plates (see Recipes) (Note 3)
Liquid LB media (see Recipes)
PNA solution (per 20 mL) (see Recipes) (Note 2)
(Optional) materials for building MALDI matrix spray station
CNC machine (Huizhou Bachin Electronic Technology, catalog number: T-A4)
Windows PC compatible with the CNC machine
Chemical safety hood with working ports for nitrogen, air, vacuum, and electricity. The plastic barbed connector for the nitrogen port must be detachable by unscrewing, exposing the female opening for the pipe
Syringe pump (KDS Scientific, catalog number: KDS-100-CE)
Lab stand with a clamp
Two adjustable wrenches
Caliper
25′ of 1/4″ nylon tubing (McMaster-Carr, catalog number: 5548K75)
Two 50 mL syringes with Luer tapering (BD, catalog number: 309653)
1/4″ × 1/16″ stainless steel reducing union (Swagelok, catalog number: SS-400-6-1)
1/16″ OD × .020″ ID stainless steel tubing, 5 cm (IDEX, catalog number: U-101)
1/16″ OD × .030″ ID stainless steel tubing, 5 cm (IDEX, catalog number: U-115)
1/16″ Union Tee (Swagelok, catalog number: SS-100-3)
Two 1/16″ OD × .0155″ ID × 1.6″ NanoTightTM tube sleeves (IDEX, catalog number: F-242)
350 μm OD × 250 μm ID fused silica capillary (Polymicro, catalog number: 1068150026)
10-32 female-to-female Luer adapter (IDEX, catalog number: P-659)
1/16″ OD one-piece finger-tight fitting, 10-32 coned (IDEX, catalog number: F-120Y)
Gorilla glue (The Gorilla Glue Company)
(Optional) HTS MALDI adapter 1.0 mm (Bruker Daltonics, catalog number: 1847571)
(Optional) HTS MALDI plates (Bruker Daltonics, catalog number: 1833280)
(Optional) MTP target frame (Bruker Daltonics, catalog number: 8074115)
(Optional) MTP 384 target plate polished steel (Bruker Daltonics, catalog number: 8280781)
(Optional) 1/4″ tube OD × 3/8″ ID-18 NPTF male connector (Parker, catalog number: 68C-4-6)
(Optional) 1 mL glass syringe (Hamilton, catalog number: Gastight #1001)
Recipes
LB agar plates
Make 500 mL of 2× concentrated LB media (20 g of LB powder per 500 mL media).
Perform filter sterilization of the solution.
Add 250 mL of deionized water into a 1 L flask.
Add 7.5 g of agar into the flask.
Autoclave the flask for 20 min at 15 psi and 121 .
Immediately after completing the autoclave sterilization, transfer the flask to a biosafety hood.
Add 250 mL of the 2× LB media at room temperature into the flask, while swirling the flask.
Add carbenicillin (100 µg/mL), IPTG (0.1 mM), and glucose (1% w/v).
Thoroughly mix the media by swirling the flask.
Pour 40 mL of the media into each 150 mm Petri dish.
Leave the dishes completely open and dry for 30 min in the biosafety hood.
Put the dishes into plastic bags and store at 4 °C. During storage, the dishes should face upside down.
Liquid LB media (500 mL)
After dissolving 10 g of LB powder into deionized water (500 mL), perform filter sterilization of the solution using the vacuum filter unit.
PNA solution (per 20 mL)
Reagent Quantity
PNA 100 mg
Acetonitrile (Note 2) 20 mL
Equipment
PC with internet connectivity and Firefox or Chrome web browser
Flatbed scanner (EPSON, catalog number: Perfection V300 Photo)
Biosafety hood
Chemical safety hood
-20 °C freezer
-70 °C freezer
4 °C refrigerator
Autoclave
Cell culture incubator (Fisher Scientific, catalog number: 1600D)
Electroporator (Bio-Rad, catalog number: Gene Pulser Xcell)
Refrigerated centrifuge (Eppendorf, catalog number: 5810 R)
UV-Vis spectrophotometer (Thermo Scientific, catalog number: Genesys 10S)
Cell culture shaker (New Brunswick, catalog number: I 24) with clamps for 250 mL flasks
MALDI-ToF-MS (Bruker Daltonics, catalog number: ultrafleXtreme)
Ultrasonic bath sonicator (Cole Parmer, catalog number: 8891R)
Solid insert tray for sonicator bath (Branson, catalog number: 100410174)
Milli-Q water supply (ELGA LabWater, catalog number: PURELAB flex)
Ice supply
Electronic scale (Mettler Toledo, catalog number: PB4002-S)
(Optional) Acrylic nitrogen purge cabinets with nitrogen flow (Cleatech, catalog number: 1500-1-A)
(Optional) IR Thermometer (FLUKE, catalog number: 62MAX+) (Note 4)
Software
flexControl 3.4 (Build 135.12, Compass for flexSeries 1.4, Bruker Daltonics)
Bachin Draw (Huizhou Bachin Electronic Technology, http://www.bachinmaker.com)
Inkscape 1.0 (Inkscape, https://inkscape.org)
macroMS (Lab software, https://macroms.scs.illinois.edu/)
flexAnalysis 3 (Bruker Daltonics)
Control software for scanner
Procedure
Sample preparation (Note 3)
Colony culture
Transform electrocompetent cells with > 50 ng of plasmid mutant library (Note 5).
Resuspend the bacterial cells using 1 mL of LB media and transfer the cells into a cell culture tube holding 2 mL of LB. Grow the cells within a shaking incubator for 1 h at 37 °C.
Add antibiotics to the 3 mL of LB solution (100 μg/mL carbenicillin) and grow overnight at 37 °C in a shaker incubator.
Prepare four microcentrifuge tubes holding 990 μL of LB and perform three 100-fold dilutions of the overnight culture by serially transferring 10 μL of sample to the next tube three times. Mix the tubes thoroughly in each dilution step. For the fourth tube, transfer 10 μL from the second tube (10,000-fold diluted) resulting in two tubes having 1,000,000-fold diluted samples.
Using an inoculating turntable and a spreader stick, spread 500 μL of the diluted culture solution onto a prewarmed 150 mm plate in the biosafety hood. The liquid should be spread over an area like the 90 mm filter paper.
Immediately after, use the tip top of the shorter leg of the L-spreader stick to uniformly spread the liquid layer while slowly rolling the turntable. Do not spread until the liquid becomes dry.
Dry the liquid layer in the hood. Seed three more plates in the same manner.
Seal the plates with parafilm-M (Note 6).
Label the sample name at the top corner of the bottom dish, including the plate number. The up-down orientation of the label decides the sample orientation throughout the sample preparation workflow (Figure 1).
Figure 1. Maintaining a constant up-down orientation of the sample slide throughout the sample preparation workflow. Up-down orientation of the target plate should be equal to the up-down orientation of (A) Petri dish when transferring colonies, (B) flatbed scanner glass when imaging, (C) the image of the target plate, and (D) MALDI target adapter when mounting the target plate.
Incubate the Petri dishes in the cell culture incubator without water bath. Do not stack the plates in the incubator and set the plates facing upside down.
In ~12–18 h, collect the plates when the colonies are ~1–2 mm.
Imprinting colonies
Immediately after the completion of the cell culture step, seal the plates by an additional layer of parafilm and put the plates into a sealed plastic bag. To prevent drying of the colonies, all subsequent steps until the imprinting step should be completed in 2 h.
Attach two layers of tape on the left and right edge of the MALDI target plate, while avoiding covering the fiducial marks at the corner edges. This step, step B1b, and step D6z are not necessary if the HTS MALDI plates are used. Write the sample name on the tape.
Label the sample name on the backside of the MALDI target plate and on the tape at the frontside. The up-down orientation of the labels should be used for preventing placing the slide upside down in the scanner and on the MALDI target adapter (Figure 1).
Get an equal number of A4 papers as the Petri dishes and tightly fold individual papers into half.
Using your hand, remove particulate matter on the frontside of the protective steel plate (0.012″ thickness) and on the backside of the MALDI target plate.
Set the MALDI target on the protective steel plate, with the mirror polished side facing up. Check the rotational orientation of the labeling letters of the target plate (Figure 1).
Place the Petri dish next to the MALDI target, with the colonies facing up (Figure 2). Orient the Petri dish so that the labels are placed on top. The subsequent steps until the end of step A2m should be completed rapidly to minimize drying of colonies.
Figure 2. Setup for colony transfer. The Petri dish with colonies (left) and the MALDI plate sitting on the protective steel plate (right). The filter paper for colony transfer (top) is placed next to the plate.
Open the cover of the Petri dish.
Immediately set the sterile filter paper onto the colonies and close the cover of the Petri dish. Wait at least 30 s for the complete attachment of the colonies onto the paper.
Using a tweezer, grab the bottom side of the filter paper and gently pull off the filter paper from the agar plate at roughly 10 cm/s speed while avoiding lateral movement of the filter paper on the agar surface. Immediately lay the top, middle, and then the bottom side of the filter paper onto the MALDI target, having the colonies directly touch the target plate. It is important not to change the rotational orientation of the filter paper, meaning the tweezer should still be grabbing the bottom side of the filter paper when the filter paper is placed onto the MALDI target.
Quickly move the steel protector (with the MALDI target plate and the filter paper) onto the bottom surface of the floor.
Quickly set an unused folded A4 paper onto the filter paper, set the sponge on top of the paper, and set the 3/8″ thick steel plate on top of the sponge (Figure 3). Avoid lateral movement of the filter paper while placing the paper, the sponge, and the plate. The center of these items should be roughly aligned as they are placed.
Figure 3. Colony imprinting tool. The imprinting tool consists of a 3/8″ steel plate, flat sponge, folded A4 paper, filter paper with colonies, MALDI target plate, and protective steel plate, from top to bottom.
Immediately step onto the 3/8″ steel plate using your full body weight (> 50 kg). Set the center of the foot roughly near the center of the plate and press using both upside and downside of the foot; avoid pressing only one area at the periphery of the plate. Add extra weight by pushing up the edge of the table or hood using hands (> 1 s).
Remove the steel plate, the sponge, and the paper sequentially.
Place the MALDI target plate on the table and wait 30 s for drying.
Using the tweezer, remove the filter paper from the MALDI target plate (Figure 4). Put the used filter paper and the A4 paper into the biohazard trash can.
Figure 4. Example of a MALDI target plate with imprinted E. coli colonies
Store the target plates in the vacuum desiccator from step D5 for 5 min and remove weakly attached colonies by gentle airflow using the air tube (Note 7).
If no visible biomass is left on the agar dish after removing the colonies, incubate the plates in the cell culture incubator until a visible amount of biomass reappears. If the cells do not regrow, try reducing the initial cell culture time (step A1k).
Seal the agar plates with parafilm-M and store them in a 4 °C refrigerator.
Imaging using flatbed scanner
Open the cover of the scanner and leave the cover lifted.
Clean up the scanner glass with ethanol wipe, removing dust and particulate matter.
Set the target sample slides onto the scanner glass with the sample side facing down.
Check the rotational orientation of the letters on the back of the target plate to avoid rotationally flipping the plate upside down (Figure 1).
Orient the target plate so that the top and bottom edge of the target plate are perpendicular to the side edges of the frame of the scanner glass.
If the target plate is an ITO slide, turn off the room light. If the target plate is a nontransparent metal plate, close the cover.
Open the control software for the scanner.
Set the resolution to 800 dpi.
Set the scanned area so that the target plate(s) is included while minimizing the off-target area. Up to four target plates can be imaged together.
Obtain the width and the height of the scan area in pixel number. Calculate the total pixel count by multiplying these values and check if the total pixel is smaller than the max pixels allowable by the macroMS software, which is 30,000,000. In other words, macroMS does not accept an image larger than ~300 cm2 at 800 dpi.
Check if all the fiducial points are visible in the image (Figure 5).
Figure 5. Example of a fiducial mark. The point highlighted by the yellow arrow is a laser etch mark that is created during building the custom MALDI target plate. This point should be visible both in the image and in the video feed of the MALDI-MS instrument. This etch mark is ~200 μm.
Optionally, the image thresholds can be adjusted to increase the contrast between the colonies and the background. The background should be either clearly darker or clearly brighter than the colonies. The background should have uniform darkness.
Perform scanning. For mirror polished steel plates, if white vertical stripes appear due to the LED light bulbs, rotate the plates 90° and scan again.
Confirm that the total pixel count for the image is less than 30,000,000. For this, right-click the image file and click properties. Find the vertical and horizonal dimension in pixel counts and obtain the total count by multiplying the two values (i.e., 4,000,000 for a 2,000 × 2,000 image). If the number is larger than the limit, use lower dpi resolution.
Confirm that the fiducial points for each of the plates are visible in the image.
Confirm that the up-down orientation of the sample plate is correct (Figure 1).
Name the filename informatively.
Clean up the scanner glass with ethanol wipe.
macroMS for image analysis and target recognition
Confirm that the total pixel count for the image is less than 30,000,000. Refer to step A3n.
Using either Firefox or Chrome web browser, visit macroMS web tool at https://macroms.scs.illinois.edu. (Video walkthrough for operating macroMS can be found at https://www.youtube.com/watch?v=OckpuZsUfSg. More in-depth instructions can be found at https://kisurbc2.gitbook.io/macroms).
Submit the image file.
Using the Edit fiducials tool, put three fiducial marks at the fiducial points at the three corners of the target plate in the image. Use the max zoom to set the fiducial mark at the precise center of the fiducial point.
Using the Set ROI tool, set the region to perform target finding. Select the area in the plate where the colonies are located.
Select the image threshold type in the dropdown menu. Gray is the most frequently used.
Using the scroll bar, adjust the threshold value. The image panel immediately displays the image that is thresholded at the current value. Using this feedback, decide the threshold that results in more visible colonies while minimizing the background noise.
Set the minimum and maximum size for target finding. For 2 mm colonies in 800 dpi image, 200 and 1,000 can be example values for minimum and maximum size.
Set the circularity limit to approximately 0.3.
Perform target finding (Figure 6A). Several trials can be done to find the parameters increasing true positive identification and reducing false negative identification. The optimized values for the parameters are reusable as long as the next images have the same dpi resolution and similar colony sizes.
Figure 6. Overview of the functions of macroMS software. A. Finding colonies by image analysis on a scanned photo of a 50 mm × 75 mm ITO slide (Choe et al., 2021). B. Data analysis output listing signal strength for peaks of interest for all screened colonies. C, D. Data visualization tool that matches the data analysis results to individual colonies. E, F. Finding a colony from a Petri dish using the colony-hopping functionality of the data visualization tool.
If areas around the colonies are to be sampled, perform circular packing with offset value of 1.
Using the filter tools and the target editing tools, remove target boxes from false positives (i.e., merged colonies and particles) and add target boxes to false negatives. An option for addressing high false negative rate is screening more colonies from additional agar plates.
Select the target plate type and download the geometry file. Rename the geometry file informatively.
Move the file to the folder containing the geometry files for the flexControl software (i.e., D:\Methods\GeometryFiles). Never edit the content of the geometry file manually.
Spraying MALDI matrix
Spray approximately 0.1–0.3 mg/cm2 of PNA matrix onto samples using the sprayer owned by the lab. If the lab does not have a sprayer, an option is to build a sprayer device and perform the steps as described in section D. For peak alignment, putting one or two internal mass calibrants into the solution is recommended (Note 8).
Remove the tapes on the target plate from step A2b.
Using methanol wipes, remove the matrix from the fiducial points.
At the fiducial point near the top left corner edge of the target plate, spot 0.5 μL of the gold colloid solution and dry the droplet.
MALDI-MS analysis (Note 9)
Mounting target plate onto adapter (Note 1)
Take the adapter for MALDI-MS target plate into the chemical safety hood where the target plates are located.
Attaching target slide or ITO glass onto PACII adapter:
i. Remove particulate matters on the MALDI target adapter and on the backside of the target plate.
ii. Soak a paper towel with ethanol and thoroughly wipe the area on the adapter and the MALDI target where copper tapes will attach to. Pressing down with some force while wiping is recommended.
iii. Mount the MALDI target onto adapter. The up-down orientation of the target plate should be the same as the orientation of the MALDI adapter, so that the letters behind the target plate are not flipped upside down (Figure 1).
iv. Tape the plate onto the MALDI adapter using the one-sided copper tape (Figure 7). Flatten the tape as much as possible; there is little tolerance for the height differences within the instrument. The copper tape should not touch more than one surface of the adapter.
Figure 7. Example for placement of a copper tape. The copper tape does not reach above the upper edge boundary of the MALDI adapter.
Attaching target slide or ITO glass onto adapters other than PAC II adapter.
Mount target plate onto MALDI adapter, following the product manual. Check the up-down orientation of the target plate.
Put the adapter inside the transport box.
Change to new gloves and carry the box to the MALDI-MS instrument.
Setting up optically guided MALDI-MS
Insert the MALDI target adapter with the plate into the instrument.
Press the Load/Eject button. From this point, a video walkthrough for setting up the instrument can be found at https://www.youtube.com/watch?v=OckpuZsUfSg.
In the Geometry dropdown menu, select the geometry file downloaded from macroMS.
Click the Sample Carrier tab and click the Teach button.
From the drop-down menu, find the entries that are labeled as “fiducial_X_Y_null.” For example, “top_left” in “fiducial_top_left_null” means the corresponding fiducial point is located near the top left corner edge of the target plate (Figure 8). After selecting one fiducial point, click Go. After the automated movement, move the target mark in the video feed to the center of the fiducial point by clicking in the video feed. When the target mark is centered at the fiducial point, click Reached (Note 10).
Figure 8. Fiducial training in flexControl
Repeat the above procedure for the second and the third fiducial point.
Click the OK button.
Click several target points in the target map underneath the video feed. Check if the target mark in the video feed moves to the area within the targets. If not, check if the correct geometry file is loaded or if fiducial training is done correctly.
Select the desired MALDI-MS method file.
Perform mass calibration using the mass calibrant added at the top left fiducial point. For gold nanoparticles, peaks from ~1 to 7 Au can be used for calibration in positive or negative ionization mode.
Sacrificing a few colonies, adjust the following parameters that result in good intensity for peaks of interest (Note 11):
i. Laser intensity
ii. Number of laser shots
iii. Diameter of random walk
Click Detection tab and adjust the mass range. For data analysis by macroMS, the width of mass range should not be above 1,000 Da.
Save the method file.
Click the AutoXecute tab, select the desired AutoXecute method file, and set the laser settings in the editor as described in Note 12.
On another colony, check the peaks based on AutoXecute method by clicking Run method on current spot.
Click the AutoXecute tab.
Click the New button for the run and select the current geometry file.
Click Next twice.
At the sample selection page, mouse drag and drop select all the target positions (Figure 9). Avoid selecting fiducial points at the corner edges of the map.
Figure 9. Selecting colonies as targets for MALDI-MS. By mouse click-and-drag, the targets generated by macroMS can be selected for MALDI-MS analysis.
Click Next, choose the AutoXecute method, select the path for the output files, and type in the sample name. Click Next and click Finish.
Check the information listed in the window and click OK.
Type in the name for the sequence file and save.
In the mass spectrum window, zoom into one dominant peak of interest by Zoom X axis only and click Auto Vertical Scale checkbox.
In the AutoXecute tab, click Start automatic Run. Check the intensity of the peak for the first ~10 acquisitions.
Cleaning reusable MALDI targets
Squeeze the wash bottle for isopropyl alcohol and rinse off the MALDI matrix from the target plates. Collect the waste in a 10% bleach solution. During the cleaning steps, be careful not to create scratch marks on the mirror polished surface.
Remove the colonies on the plates using Kimwipes soaked with water, collecting the waste in 10% bleach solution.
Put the plates and waste into the 10% bleach solution and incubate for 30 min.
Put the plates into 100% isopropyl alcohol and sonicate for 10 min.
Dry the plate using Kimwipes.
Store the plates in a closed container.
Data analysis
Refer to the Data Analysis section for identifying colonies to isolate in the following steps.
Downstream testing
Colony picking
Open the data visualization map that is included in the data analysis email.
In the map, type in the colony ID of interest into the input box. This highlights the colony.
Obtain the corresponding Petri dish from the 4 °C refrigerator from step A2s.
Rotate the Petri dish so that the labels are at the top (See steps A1i and A2c). This means the up-down orientation of the colonies in the image and in the Petri dish are the same (Figure 1). Also, the colonies side of the Petri dish should be facing towards the observer.
Find any group of colonies that are identifiable from both the image and the plate. They are arranged in the shape of letters like I, O, and U. These colonies can be at the edge of the image.
Perform colony-hopping to gradually move towards the desired colony (Figure 6E, 6F).
i. Using a pen, circle the spot on the Petri dish where the colony from step C1e is located.
ii. By mouse drag and drop, highlight the same colony on the image page.
iii. Find the next colony that is closer to the colony of interest and is identifiable both in the plate and in the image panel.
iv. Mark the colony on the Petri dish and highlight the same colony on the image page.
v. Repeat finding and labeling a colony that is incrementally closer to the colony of interest until reaching the destination.
vi. Circle the spot on the Petri dish where the colony of interest is located.
Touch the colony with the tip of a sterile P1000 tip and circle the colony several times. If the desired colony is too small or too close to the adjacent colonies, a colony can be picked using a sterile pin following the steps below.
i. Tape a pin onto a pen.
ii. Tape the pen onto the head of a dissecting microscope.
iii. Set the Petri dish onto the stage of the microscope and turn on the background light.
iv. Using a lighter, burn the pin several seconds and wait 30 s for cooldown.
v. Move the pin towards the colony and set the tip of the pin ~2 mm directly above the colony. Use the coarse adjustment knob of the microscope for movement in Z-axis and move the Petri dish by hand for movement in the X-Y axis (Figure 10).
Figure 10. Colony picking by a flame sterilized pin. The coarse adjustment knob of the dissecting microscope is used for the precise up-down movement of the colony picking pin.
vi. Use the coarse adjustment knob again to lower the pin and make physical contact between the tip of the pin and the colony. Optionally, maximize the transfer of cells onto the pin by moving the plate a little (< 0.5 mm) while making the contact.
vii. Lift the pin away from the colony.
viii. Remove the pen.
From the isolated cells, start 5 mL overnight cultures.
Purify plasmid DNA from the overnight cultures using the plasmid extraction kit following the manual included in the kit. Save the plasmid DNA at -20 °C.
Freshly transform electrocompetent cells with the purified plasmids and the control plasmid. The electrocompetent cells should be from the same batch.
Seed the isolates and the control into different sections of the same 150 mm agar plate. The plate should contain the same media that was used to form the products of enzyme activity (step A1e). For seeding, spot 50 μL of the cell media onto agar Petri dish and streak the liquid using a sterile pipette tip.
MALDI-MS confirmation of the regrown colonies (Note 13)
Using P200 pipette tips, pick a colony on Petri dish and thinly spread onto a well of MTP 384 target plate (Figure 11). Due to possible spread of the MALDI solution, leave an empty well between the samples. Transfer three colonies from each isolate and control strain.
Figure 11. Example of spreading microbial colony onto a MALDI target well
Spot 1 μL of MALDI solution (10 mg/mL dissolved in acetone) to each well and dry the solution by evaporation (Note 14).
Perform MALDI-MS (4,000 shots, complete random walk, and ~40%–60% laser power). Refer to the data analysis section for the isolate analysis.
Building spray station and applying MALDI matrix (Figure 12) (Note 15)
Figure 12. Overview of the MALDI matrix spray station in a chemical safety hood. A sprayer head is mounted at the penholder of the CNC drawing machine. When the matrix solution and the nitrogen gas are supplied, the sprayer head nebulizes the matrix solution into aerosol. While spraying, the drawing machine moves the sprayer head in step motion, uniformly coating MALDI plates with matrix.
Nitrogen gas circuit
Using an adjustable wrench, unscrew the barbed connector for the nitrogen outlet of the chemical safety hood, exposing the female port.
Using the inside measuring jaws of a caliper, measure the diameter of the female port (distance between one inside edge and the opposite inside edge) and obtain the nearest fractional inch (Note 16).
Obtain a 1/4″ tube × D″ NPTF male connector where D is the measured fractional inch.
Screw the connector into the female port.
Cut 6′ of the 1/4″ nylon tube.
Connect the 6′ nylon tube to the 1/4″ side of the connector. To connect, untighten the hex nut for the 1/4″ side of the connector and insert the nylon tube into the hole. Using two wrenches, strongly tightening the hex nut will firmly fix the nylon tube onto the connector. Confirm the connection by pulling the tube from the connector. This method, untightening the hex nut using the adjustable wrenches, inserting the tube, and tightening the hex nut again for affixing the tube, applies to all connections mentioned below.
Connect the other side of the 6′ nylon tube into the 1/4″ × 1/16″ reducing union.
Connect a 1/16″ OD × .030″ ID tubing to the 1/16″ opening of the reducing union.
Connect the other side of the 1/16″ stainless steel tubing to the middle opening of the 1/16″ union tee. Bend the 1/16″ steel tubing by force like in Figure 13.
Connect the second opening of the 1/16″ union tee to the 1/16″ OD × .020″ ID tubing.
Connect the third opening of the 1/16″ union tee to the NanoTightTM tube sleeve. The constructed sprayer head is shown in Figure 13.
Figure 13. Completed sprayer head. The capillary tubing is inserted at step D2g.
Liquid circuit
Screw the Luer adapter into the Luer lock of the 50 mL syringe.
Partially screw the one-piece finger-tight screw fitting into the Luer adapter.
Fully insert the green tube sleeve into the screw fitting.
Cut 3′ of the capillary fused silica capillary.
Insert capillary through the tube sleeve until approximately 0.2″ of the capillary enters the syringe.
Strongly tighten the screw fitting. Check if the capillary moves by slightly pulling by hand (roughly 0.1 N force). If so, tighten the screw fitting stronger. The completed assembly is shown in Figure 14.
Figure 14. Syringe head
Connect the other tip of the capillary to the sprayer head:
i. Slightly open the hex nut for the green tube sleeve.
ii. Insert the capillary through the tube sleeve.
iii. Push the capillary until approximately 0.5 mm of the tip of the capillary goes outside of the end of the 1/16 stainless tube (Note 17). Avoid pulling the capillary from the sprayer head mistakenly in the subsequent steps.
iv. Tightly close the hex nut.
Setting up the CNC machine
Using the online manual, set up the Bachin Draw CNC machine and the control PC (Note 18). While doing so, set the Windows PC right adjacent to the side of the chemical safety hood where the CNC machine will be located. Place the CNC machine inside the hood (Note 19)
Cover the bottom surface of the CNC with aluminum foil. Attach the foil using tape.
(Optional) Place the pen holder at the home position (top left corner edge). On the aluminum foil, mark a dot at 2 cm towards the 4:30 direction from the pen holder.
(Optional) Draw a horizontal line and a vertical line crossing at the dot. The lines should be parallel to the horizontal and vertical edges of the rectangular surface of the bottom plate of the machine. Record these lines by attaching six pieces of 2-inch tape along these lines.
Position the syringe pump next to the CNC machine.
Glue the sprayer head onto the pen holder following the steps below.
i. Completely tighten the hex nuts of the sprayer head. Disconnect the 6′ nylon tube from the 1/4″ × 1/16″ reducing union.
ii. Wet the locations of the pen holder that will make physical contact with the sprayer head with water (Figure 15).
Figure 15. Locations for applying the Gorilla glue to attach the sprayer head to the pen holder of the CNC machine
iii. Apply Gorilla glue.
iv. Wet the surfaces of the Neodymium magnets provided by the CNC device and apply the Gorilla glue onto the surface of the magnets.
v. Mount the magnets on the steel area that is opposite to the blue screw.
vi. Affix the sprayer head onto the pen holder by strongly tightening the blue screw, pressing the bottom hex nut against the magnets (Figure 15). Check the glued positions make contact between the sprayer head and the penholder.
vii. Apply the glue on the same locations of the penholder again.
viii. Wait 24 h until the glue solidifies completely.
ix. Connect the 6′ nylon tube back into the 1/4″ × 1/16″ reducing union.
Stabilize the sprayer by inserting its nylon tube into the clamp of the lab stand (Figure 12). The distance between the clamp and the sprayer head should be at least 4′ to allow flexibility during the spray.
Setting up the control software for the CNC machine
Open the Inkscape software. (Instead of drawing the spray path, the provided drawing files can be used. See Note 20 for dimensions. Jump to step D4r if the provided files are used.)
In millimeters, calculate the horizontal and vertical size of the area that MALDI matrix solution will be sprayed. The horizontal and vertical size of the area should be 40 mm larger than the MALDI target plate (i.e., for a 110 mm × 75 mm plate, the spray area will be 150 mm × 115 mm).
Click the Draw Bezier curves and straight lines button at the left sidebar.
Click at point where X = 1.00 and Y = 1.00. The coordinate of the mouse can be obtained at the right bottom corner of the window.
Click the second point where X = 1.00 + the horizontal size of the spray area and Y = 1.00.
Click the third point where X = 1.00 + the horizontal size of the spray area and Y = 3.00.
Click the fourth point where X = 1.00 and Y = 3.00.
Click the fifth point where X = 1.00 and Y = 5.00. Terminate the drawing by pressing the enter key. The completed single step of the spray path is shown in Figure 16.
Figure 16. Drawing for a single step of the spray path
Select the drawing by clicking. Copy and paste the drawing.
Join the copy and the original by moving the copy so that the fifth point of the original drawing touches the first point of the copy. The touching is indicated by the message “Cusp node to cusp node.”
Zoom into the spot where the two points of the drawings are joined (Figure 17). Click Edit paths by nodes (N) button at the left sidebar.
At the zoomed view, by hold shift + clicking, select both the line from the original drawing and the line from the copied drawing.
By holding click + dragging, select the spot where the two lines meet. Then, click Join selected nodes button at the top menu. This results in one continuous joined line (Figure 17).
Figure 17. Joining two nodes to create a continuous single spray path
Repeat copying the drawing followed by joining the nodes until the vertical size of the resulting drawing grows to the desired vertical size of the spray area (Figure 18).
Figure 18. Path for one round of spray coating. Multiple steps in Figure 16 are joined by node-to-node joining indicated in Figure 17. The area of the spraying should be 4 cm larger than the size of the sample.
By hold-click + dragging, select the entire spray path. Then, copy and paste the path into the same canvas. Then, move the copied content precisely above the original, so that the copied content hides the original completely. While moving the copied content towards the original content, the software assists with the complete overlapping.
Repeat the above step, which results in four overlapping paths. This means four repeats of the spraying process.
Save the file in SVG format.
Open the SVG file in Bachin Draw software and move the top left corner edge of the spray area to the origin point of the canvas. The origin is where the two pink lines cross (Figure 19).
Figure 19. Spray path loaded into the CNC control software. The spray path indicated in red is the path drawn by the Inkscape software.
Without liquid or gas, press the start button of Bachin Draw to complete the full path defined by the SVG file. During the mock spraying process, find and remove obstacles blocking the path of the spray head, the capillary, and the gas tube. Also, check if the gas tube touches the aluminum foil or creates too much strain on the 1/16″ × .020″ tubing of the sprayer head. Avoid these by adjusting the length of the nylon tube or the location of the lab stand or by creating rotational strain on the tube by twisting. For creating the rotational strain, unscrew the hex nut holding the nylon tube, twist the tube, and tightly screw the hex nut again while holding the twisted tube by hand.
Setting up vacuum desiccator
Cut the 5′ polyurethane rubber tubing into half.
Set the vacuum desiccator inside the hood.
Using the polyurethane rubber tubing, connect the barbed connector on the desiccator to the barbed connector for the vacuum outlet of the safety hood.
Connect the barbed connector for the airflow outlet of the safety hood to polyurethane rubber tubing.
Spraying MALDI matrix
Wear a cuffed sleeve lab coat that fully covers the wrist.
Prepare 20 mL of PNA matrix solution at 5 mg/mL in acetonitrile (Note 2) and put into the 60 mL syringe. For peak alignment, putting one or two internal mass calibrants into the solution is recommended. See Notes 8, 17, and 21.
Put 50 mL of methanol into another 60 mL syringe. Put ~3 mL of air into the syringe.
Prepare a 50 mL tube containing acetonitrile.
Check if the PC communicates to the CNC by pressing the down button of the Bachin Draw software. The CNC should respond by moving the sprayer head (Note 22). Then, press the home button.
Place a plastic cap of a 50 mL tube under the nozzle of the sprayer head.
Input the width of the syringe and the flowrate into the syringe pump (i.e., 24.5 mm and 25 mL/h for the 60 mL syringe).
Connect the methanol wash syringe to the sprayer head through the capillary.
Purge the capillary with ~1 mL of the wash solution by manually pushing the plunger.
Empty the capillary by putting air from the wash syringe.
Remove the Luer adapter and the capillary from the wash syringe (non-syringe components of Figure 14) and connect them to the syringe holding the matrix solution.
Mount the syringe for the matrix solution onto the syringe pump.
Disconnect the gas line from the 1/4″ to 1/16″ connector by unscrewing the hex nut of the gas line. Purge the gas circuit of the sprayer head by pipetting 2 mL of the acetone solution into the opening of the 1/4″ to 1/16″ connector (Figure 20). Remove the solvent in the tube by pipetting air through the tube. Connect the gas line back into the connector again. Avoid bending the 1/16″ tube in the process.
Figure 20. Cleaning of the sprayer head for removing solid residue in the sprayer
In Bachin draw, open the appropriate SVG file defining the spray area for the target plate(s). At 25 mL/h flow rate, files with 4 sweeps should be selected. Make sure the top-left corner edge of the draw path is placed at the origin point.
Place the target plate(s) inside the spray area on the aluminum foil.
Close the stash of the chemical safety hood as much as possible.
Turn on the nitrogen gas line for the sprayer head.
Turn on the syringe pump.
Wait 30 s.
Check if the spray is started by appearance of the spray under the spray nozzle (Note 23). Completely shut the stash of the safety hood.
Press the start button on the CNC control software. After checking for 1 min, evacuate from the room.
After spraying, open the safety hood minimally and stop the syringe pump first.
Wait 30 s and close the nitrogen gas line.
Wait 10 min for the target plate to dry and for the aerosols from the spray to fully disappear.
Connect the capillary to the wash syringe and purge the capillary with ~1 mL of methanol.
Using methanol-soaked Kimwipes, remove the matrix from the fiducial marks on the plates.
Spot 0.5 μL mass calibrant at the fiducial mark near the top left corner of the plates.
Dry the calibrant solution.
Remove the tapes on the target plates and proceed to step B2.
If the MALDI-MS screen is performed on a separate day and if a nitrogen cabinet is available, put the plates in a clean plastic container, seal the box using parafilm-M, and put the box into the cabinet. Wipe the outside of the box with ethanol before putting into the nitrogen cabinet.
Data analysis
(A video walkthrough for the data analysis is available at the “Submitting data analysis” and “Opening processed data” section at https://www.youtube.com/watch?v=OckpuZsUfSg) Using flexAnalysis, confirm that the width of the mass range of the acquired mass spectra is below m/z 1,000. For example, if the acquired mass spectrum ranges from m/z 300 to m/z 1,400, the data file cannot be analyzed by macroMS. Do not manually modify the mass spectra data files including file names.
Compress the mass spectrum data into a zipped folder. It is important to compress the folder containing the individual data files and not compress the individual data files selected by Ctrl + A key. For example, if the path to the folder holding mass spectra data is C:\Data_313\BTE6 such that the path to individual mass spectrum data is C:\Data_313\BTE6\0_0_366.64_152.14_5 and C:\Data_313\BTE6\0_0_902.34_857.42_3, compress the BTE6 folder, not the Data_313 folder or 0_0_366.64_152.14_5_1.0. folder.
Obtain the image file used to generate the target list. Check that the name of the image file is informative enough to identify the sample, since this name will be used as the title for all data outputs. If not, rename it more informatively.
Create a text file containing m/z ranges for up to 10 peaks of interest. In flexAnalysis, load 10 randomly selected mass spectra in overlaid mode. Navigate to a peak of interest, then find two m/z values that can span all the overlaid peaks. Input the values in the text file (Figure 21). The text file is tab-delimited, meaning the tab key must be used to separate the values instead of space bar. Alternatively, cutting and pasting from two columns of Excel spreadsheet results in tab-delimited entries in the text file.
Figure 21. Defining a peak of interest. If a peak of interest appears at approximately 218.17, the m/z range values (218.02 and 218.33) that can contain this peak in all the 10 overlaid mass spectra should be inserted into the text file. In the file, the two values should be separated by a tab key.
Optionally, peak alignment can be done by adding exact mass for one or two internal calibrants that were added into the MALDI matrix solution (step D6b). The exact mass for an internal calibrant with adducts can be obtained from the chemical formula using online exact mass calculators such as the tool from sisweb.com. In flexAnalysis, load approximately 10 mass spectra in overlaid mode. Confirm the overlaid peaks corresponding to the internal calibrants appear in the mass spectra. Also, confirm there is no larger peak appearing within m/z 0.5 from the mass calibrants. Put in the exact mass as shown in Figure 22. When peak alignment is used, the format for inputting the mass range for peaks of interest changes as well. The first number is the exact mass for the compound and the second number is 70% of the width of the overlaid peaks, again tab-delimited (Figure 22) (Note 24).
Figure 22. Defining mass calibrant for peak alignment and peaks of interest. To define a mass calibrant, the exact mass of the mass calibrant (285.279 in this case) is put into the text file. To define a peak of interest, the exact mass of the compound with an adduct and the 70% of peak width that spans all 10 overlaid mass spectra (i.e., 339.73 and 0.21) should be inserted. The two values (339.73 and 0.21) should be separated by a tab key.
Submit the email address, the zipped data file, the image, and the text file into macroMS using the input fields of the main page. Do not upload multiple files simultaneously from multiple windows or tabs, which can crash the file uploading. The speed of data processing is roughly 3 s per mass spectrum (i.e., 1 h per one colony screen generating 1,200 mass spectra).
Open the email listing data analysis results and save the Excel output file.
Open the Excel spreadsheet that lists ion counts for the peaks of interest (Figure 6B).
For a screen of an enzyme mutant library searching for an isolate with modified ratio between two products, sort samples by the sum of the two peaks and remove the bottom 70% inactive variants.
Add a column that lists the ratio between the two peaks for the remaining samples. Using the larger peak as the denominator for the ratio calculation can reduce the variability caused by signal noise.
Divide the colonies into two equally sized groups by the X coordinate and Y coordinate. Using statistics such as Student’s t-test, check if there is a significant difference in the peak ratios between the groups. If there is a significant difference, options can be multivariate outlier detection (Mahalanobis distance calculation) using the X, Y coordinate and the peak ratio value in Excel, performing outlier detection by the group, or seeing if the change observed in the outlier is bigger than the difference between the group.
Sort the rows by the ratio values of all active variants. Obtain the average and the standard deviation value of all the peak ratios. Find the values that are one or two standard deviations away from the average. For data generated from macroMS circular packing, an extra sheet lists the averaged value per colony. The average of the group should be used for the data processing.
Copy the Blob ID or Group ID for the colonies of interest, open the data visualization map from the email, and put the IDs into the input box. Click Show samples to locate the samples in the image (Figure 6C).
To plot mass spectrum of the peaks of interest for the selected colonies, select the checkboxes for the peak(s) and click Plot selected peak(s). The peaks should be within the greyed area of the shown mass spectra, and the peak intensity should be preferably larger than 4,000 (Figure 6D). Then, the colony can be isolated for downstream tests.
For analysis of MALDI-MS data acquired for regrown isolates (step C1k), load the data in flexAnalysis. Click the checkboxes corresponding to each of the files. Then, highlight all the files in blue by clicking the top file and Shift + clicking the bottom file.
Click the icon button for Find Mass List.
Click the top file and open the window for Mass List.
By Ctrl + clicking, select peak picking results for all compound of interest. If peaks are missing, go to Methods, select Edit Processing Parameters, change Peak Detection Algorithm to Centroid, and run Find Mass List on all the files again.
Copy and paste the data into an Excel file. Likewise, transfer the data for all samples.
Compare the isolates against the control based on the ratio of peak areas using a Student’s t-test or create a scatterplot (Figure 23B) visualizing the change.
Figure 23. Example datasets for the MALDI-MS screening of E. coli colonies expressing UcFatB2 random library. A. Per colony ratio between C12 free fatty acid and C14 free fatty acid from a MALDI screen, as measured by analyzing imprinted colonies coated with MALDI sprayer. Two colonies (red arrows) showing the lowest C12/C14 FFA ratio were isolated for retest. B. C12/C14 FFA ratio of the regrown isolates and the wildtype control, as measured by analyzing manually spread colonies treated with dried droplet MALDI solution.
Validation of protocol
This protocol was validated by a work modifying substrate specificity of thioesterase (Jindra et al., 2023) and another work modifying regioselectivity of desaturase (Choe et al., 2023).
Notes
From a machine shop facility, build the following metallic target plates (Figure 24) using #8 mirror polished steel (9785K14, McMaster-Carr). Mirror polishing steel surface by manufacturers leaves small but visible lines on the surface, and the angle of cutting the plate should not be diagonal but parallel/perpendicular to these lines. Diagonal lines can cause image artifacts during scanner imaging. The four dots are created by laser etching, and they serve as fiducial points. Alternatively, the following adapter-target pairs also work: 1) HTS MALDI adapter 1.0 mm and HTS MALDI plates and 2) MTP target frame and MTP 384 target plate polished steel. The HTS plate and adapter are highly recommended because the plate mounts to the adapter by magnet, eliminating the need to use copper tapes, a unique serial number written at the bottom-right corner of the plate can eliminate the need to label the plate by pen, its internal fiducial marks are highly visible in the instrument video feed even after matrix spray coating, and taping at the side edges to enable attachment of copper tape is not needed. Use of metallic MALDI target instead of glass ITO slide is recommended for improved sensitivity and better attachment of the E. coli colonies. For ITO slides, the fiducial points can be the corner edge point of the glass or the center of the cross mark created by a glass engraving pen.
Figure 24. Drawing of the custom MALDI target plate
Different MALDI matrices result in varying degrees of MALDI sensitivity from the imprinted colonies. While PNA and 1,5-Diaminonaphthalene (DAN) enable measurement of compounds directly from colonies, 2,5-Dihydroxybenzoic acid (DHB) and alpha-Cyano-4-hydroxycinnamic acid (CHCA) may not. For better measurement by DHB and CHCA, use small colonies (~0.5 mm) to reduce sample thickness or measure from the area around the colonies to sample compounds that are spread away from the colonies during the MALDI matrix spraying. In the second case, the density of colonies must be reduced below 250 per MALDI target plate (75 mm × 110 mm) and the offset value of 1 should be used for the circular packing tool of macroMS. Alternatively, DAN with salt could be used as an alternative to DHB or CHCA for positive ionization directly from colonies. Also, always read Material Safety Data Sheet (MSDS) to check if the MALDI matrix and the solvent used for making MALDI matrix solution are toxic (i.e., 9-aminoacridine and acetonitrile) or safer [i.e., DHB, CHCA, and N-(1-naphthyl) ethylenediamine dihydrochloride (NEDC)]. Even if the spraying is performed inside a chemical safety hood, spraying the chemical solution into a fine mist increases risk of exposure. Moreover, while transferring the sample plate to the MALDI-MS instrument, the sample plates should be in a clean container that is held wearing clean gloves. Wearing a respirator may be needed when toxic compounds are involved. Gloves should be changed frequently. For safety, acetone can replace acetonitrile when making MALDI solution. Lastly, avoid nanoparticles and carbon-based matrices (i.e., graphite, graphene, carbon nanotubes, etc.) for large MALDI screens due to their potential impact on the instrument.
Before the colony screen for an enzyme other than this thioesterase, the duration, temperature, and media composition of cell culture method for growing colonies should be decided to improve the product formation of the enzyme activity or the biochemical activity. Plasmids with different copy numbers may be tried. Also, the strain producing improved products while maintaining a good attachment to the MALDI target plate without flaking off should be chosen (i.e., K-12 strain for E. coli). Larger peak area and signal-to-noise ratio in MALDI-MS measurement results in more reliable detection of change in product profile. For E. coli, a starting point can be growing cells on LB plate made from filter sterilized media containing 100 μM IPTG, 100 μg/mL carbenicillin, and 1% glucose for 12 h at 37 °C. Optionally, colonies can form in growth plates first, and then be transferred to an inducing plate for enzyme production (Si et al., 2017). For measuring products from the colonies grown at trial experiments, use a pipette tip to transfer the biomass of the colonies onto a well of MALDI plate and thinly spread it (Figure 11). Spot 1 μL of the MALDI solution for analysis. More accurate test for peak intensity is MALDI-MS analysis of imprinted colonies that are coated with MALDI matrix by the standard spraying procedure. In addition, the workflow works with non-volatile compounds. Check the loss of the compound in the high vacuum environment of the instrument before developing a screen. First, make a mixture of MALDI matrix solution and the standards and spot onto 12 different wells in an MTP 384 plate. When the MALDI target plate is in the high vacuum environment, perform MALDI-MS analysis of the first three wells. Repeat the measurement on a different set of three wells every 30 min. Compare the differences in peak intensity for the standards. For MALDI analysis, use 5,000 laser shots, 1,000 Hz for laser frequency, and complete random walk. Lastly, for relative quantitation of a single compound by the ratio-based colony screen, due to inconsistent compound extraction and ionization that are inherent in the MALDI analysis of colonies, an ideal internal calibrant for the ratio-based screen can be a compound that is produced at the same cellular location where the compound of interest is produced at (i.e., cytoplasm vs. membrane).
The IR thermometer can be used to check the temperature of the cooling agar solution before adding in temperature-sensitive antibiotics. Also, it can be used to check the temperature of the warmed agar plate before seeding with cells.
Using electrocompetent cells grown from richer media such as filter-sterilized LB solution can result in better production of compounds. Minimize the generational gap between the electrocompetent cells and the source strain vials. Also, seeding plates with freshly transformed cells without the overnight culture step can potentially result in better product formation.
The purpose of using parafilm-M is to reduce moisture loss to enable the colony transfer steps. To authors’ experience, increasing the humidity of the cell culture incubator itself resulted in the colonies flaking off from the metal plate when imprinted. Increasing the humidity of colonies by parafilm sealing helped resolve the flaking problem.
If significant flaking happens with E. coli: 1) perform incubation in dryer incubator; 2) for growing colonies, use agar plates that were dried longer than 40 min in the biosafety hood; 3) use smaller colonies (1 mm or less) by decreasing the growth time or increasing colony density; 4) perform deep cleaning of the mirror-polished surface of the MALDI target plate using a plasma cleaner or glow discharge system; 5) consider finding a sticky host strain (Neve et al., 2020) and avoid DH5α; 6) try room-temperature growth of the colonies; 7) add IPTG/arabinose into the growth media to induce protein expression; 8) after step A2i, soak filter paper and colonies with MS-compatible adhesive such as DHB (Jens et al., 2022) and glycerol by setting onto agar gel containing these compounds, and then imprint.
Mass range between the two internal calibrants should include all compounds of interest, and the mass calibrants should not be chosen if the peak for the calibrant appears within 0.5 m/z from a larger peak due to the algorithm used by macroMS peak alignment. Amino acids and odd chain fatty acids at ~1 mg/mL could be used as internal mass calibrants.
Prior to the screen, the MALDI-MS instrument parameters and MALDI matrix should be optimized for sensitivity and peak resolution. Also, the method should be verified for analyte detection. Instrument manuals should be consulted for tuning laser power, laser beam width, number and frequency of laser shots, grid voltage, pulse delay, random walk, polarity of ionization mode, pulsed ion extraction time, accelerating voltage, extraction voltage, lens voltage, reflector voltage, and the choice of reflectron/linear mode. These parameters should be developed based on measuring the compounds of interest from the colonies grown and prepared by the same protocol. Working parameters for ultrafleXtreme include 50% laser power, Ultra laser width, reflectron mode, 4,000 laser shots, and 800 μm complete random walk. For targets generated from macroMS circular packing, 1,500 laser shots with 600 μm complete random walk can be used. Autoflex could be used for the screen workflow analyzing several Petri dishes. There are literature sources introducing MALDI-MS optimization and choosing MALDI matrix (Barwick et al., 2006; Calvano et al., 2018; Leopold et al., 2018). For verification of compound identification, mass spectrum for the colony sample by tandem mass spectrometry (MS/MS) should be compared with the mass spectrum from chemical standard. For strains with inducible production of the compound of interest (i.e., enzyme expression by arabinose induction), compound verification can be helped if the peak appears only in the presence of the inducing agent (i.e., arabinose or IPTG).
If there is a distance between the laser ablation point and the center target mark of the video feed, perform fiducial training using the actual laser ablation point. For example, if the laser ablation point is slightly below the target mark in the video feed, perform fiducial training using the laser target point not the target mark of the video feed (Choe et al., 2021). Additionally, the finalized version of the macroMS software does not use numeric values to label the fiducial points like the case at 1:56 in the YouTube video.
Occasionally, check the MALDI-MS source cleaning is performed well. Without cleaning, the MALDI-MS sensitivity can drop significantly. Click the Status tab in the flexControl software, click Details button, open Processor subsystem tab, and check source cleaning maintenance interval. Check if the value is below 12%–15%. If the value is above this range, check with the instrument administrator if the value is acceptable for MALDI-MS sensitivity.
The following describes setting up an AutoXecute file. On the AutoXecute tab of the main flexControl window, select the AutoXecute method file to open in the dropdown menu and click Edit. In the General tab of the AutoXecute Method editor, select the method file to use. In the Laser tab of the AutoXecute file, turn off the Fuzzy Control. In the Evaluation tab, select none for Use background list. In the Accumulation tab, turn off the Dynamic termination. Click on for the Fuzzy control, type in the desired number of laser shots in the input box for Sum up and Satisfactory shots and click Off for the Fuzzy control. In the Movement tab, check the checkbox for Random walk and put in 50 for Shots at raster spot. In Processing tab, select none for all fields. Click Save button. When the AutoXecute file is saved after the above steps, only the number of laser shots need to be set before the next screen.
An alternative MALDI test for the regrown colonies is macroMS-guided MALDI-MS analysis of imprinted colonies of the isolates prepared by the same spraying procedure and MALDI-MS method that were used for the original library screen. In this case, the colonies of experiment group and the control group should be imprinted using the same filter paper. Another alternative is: 1) pull all colonies from each of the sample into vials of 50 μL of water; 2) thoroughly mix the liquid samples; 3) in triplicates, spot 1.5 μL of the samples at the precise center of a well of MTP 384 target plate; 4) dry the sample by leaving in the room air; 5) apply MALDI matrix by the same spray procedure as the original colony screen; and 6) perform MALDI-MS at > 5,000 laser shots, 800 µm random walk at the center of dried droplet, and 40%~60% laser power. After these methods, perform the same data analysis steps mentioned in the protocol.
For DHB and CHCA matrix that can show reduced MALDI-MS sensitivity of the colonies sample, the spot and transfer extraction method can be used. Using a P2 pipette, spot 1 μL of the matrix solution onto the colony in the MALDI target well, pull the liquid in 1 s after making the contact, and spot onto the second well. MALDI-MS measurement is done on the second well.
For qualitative screen searching for the presence of specific compounds from colonies, MALDI matrix can be sprayed using an airbrush instead of using the spraying station. However, this can result in a reduced throughput in sample preparation and increased variability in sensitivity. Performing step D1b to obtain the connector between the pipe and gas tube is still necessary.
A common size for the female port is 3/8″. In this case, an example piece that can be used is 1/4″ tube OD × 3/8″ ID-18 NPTF male connector.
The capillary should be changed every 1–2 months. Without replacement, the capillary shows reduced mechanical strength resulting in breaking.
The manual for building the CNC machine can be found at the following link: http://www.bachinmaker.com/?p=46&a=view&r=209
It is important not to bend the terminal of the USB cable that is plugged into the USB port of the CNC device; the USB port can be easily destroyed. Stabilize the USB cable by taping the cable onto the bottom surface of the chemical safety hood. Minimize moving the CNC machine.
The vertical and horizontal dimensions of the spray area defined by the files are the following: ALL_AREA-4 Sweeps.svg: 273 mm × 190 mm, PACII-4 Sweeps.svg: 126 mm × 150 mm, TLC-4 Sweeps.svg: 62 mm × 150 mm. The files perform four rounds of spraying.
If chemical derivatization is performed, make 1.5 mL of derivatization solution in a glass vial (i.e., 5 mg/mL fmoc-chloride and 1.5 mg/mL pyridine dissolved in toluene), put into a 1 mL glass syringe, mount the syringe onto the syringe pump, connect the capillary, and perform one sweep of spraying at 10 mL/h. These steps should be done quickly because derivatization agents can start losing reactivity (i.e., 20 min for fmoc-chloride). Immediately after spraying, purge the liquid circuit with 1 mL of methanol (step D6y) and take at least 30 min in step D6x for better disappearance of the aerosols. Then, perform spraying MALDI matrix solution onto samples. For gas-based derivatization such as ozonolysis, uniform mixing of gas in the reaction chamber by applying the gas via a pump may be necessary.
If the CNC machine does not respond even after powering on the machine and connecting to the PC, go to device manager in the control panel, go to Universal Serial Bus controllers, right-click the USB device connected to the CNC machine, and try disabling and enabling the USB device. Trial and error may be needed to find the correct USB device that is linked to the CNC machine. Alternatively, turn off and on both the device and the PC.
If the air turbulence inside the hood is strong enough to visibly disturb the flow of the spray, install a mini plastic cup around the spray nozzle as a protection against the air turbulence. Also, where the spraying is intermittent at approximately 2 s cycle, perform capillary height adjustment and liquid purging (step D2giii, step D6i, and step D6m). Lastly, an alternative method for starting the gas spray would be to 1) start the pump without turning on the nitrogen gas, 2) wait for the appearance of droplets, then 3) start the nitrogen gas.
An alternative method for creating a m/z ranges list with peak alignment is aligning to a single spectrum that shows large peaks for all compounds of interest and mass calibrant. The method is justified when there is a contaminating peak at the shoulder of a target compound; this makes setting the m/z ranges using the standard method extremely difficult. Using flexAnalysis, find a mass spectrum that shows large peaks for all target compounds and calibrants. Next, maximally zoom into the tip of the peak for the mass calibrant and get the X-value where the maximum Y-value appears. Write the X-value into the text file as the mass value for the calibrant. Repeat for the second mass calibrant. For m/z ranges of the compounds of interest, use the m/z ranges (i.e., 521.1 521.3) of the peaks of the spectrum as the input values for the compounds.
Acknowledgments
This work was funded by the Department of Energy Center for Advanced Bioenergy and Bioproducts Innovation (United States Department of Energy, Office of Science, Office of Biological and Environmental Research) under Award No. DE-SC0018420. The instrumentation used was partially supported by the National Institute on Drug Abuse under Award No. P30 DA018310. Any opinions, findings, and conclusions or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the views of the funding agencies. The protocol is adapted from the validation publication (Jindra et al., 2023). The idea of imprint-transferring agar colonies by a filter paper is adapted from Si et al. (2017).
Competing interests
The authors declare no competing financial interest.
References
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Supplementary information
The following supporting information can be downloaded here:
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Analysis of Plasmodium falciparum Mitochondrial Electron Transport Chain Activity Using Seahorse XFe96 Extracellular Flux Assays
SR SaiShyam Ramesh
DC Daniela Cihalova
ER Esther Rajendran
GD Giel G. van Dooren
AM Alexander G. Maier
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4863 Views: 503
Reviewed by: Alka MehraHangjun Ke Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in PLOS Pathogens Jul 2023
Abstract
The mitochondrial electron transport chain (ETC) is a multi-component pathway that mediates the transfer of electrons from metabolic reactions that occur in the mitochondrion to molecular oxygen (O2). The ETC contributes to numerous cellular processes, including the generation of cellular ATP through oxidative phosphorylation, serving as an electron sink for metabolic pathways such as de novo pyrimidine biosynthesis and for maintaining mitochondrial membrane potential. Proper functioning of the mitochondrial ETC is necessary for the growth and survival of apicomplexan parasites including Plasmodium falciparum, a causative agent of malaria. The mitochondrial ETC of P. falciparum is an attractive target for antimalarial drugs, due to its essentiality and its differences from the mammalian ETC. To identify novel P. falciparum ETC inhibitors, we have established a real-time assay to assess ETC function, which we describe here. This approach measures the O2 consumption rate (OCR) of permeabilized P. falciparum parasites using a Seahorse XFe96 flux analyzer and can be used to screen compound libraries for the identification of ETC inhibitors and, in part, to determine the targets of those inhibitors.
Key features
• With this protocol, the effects of candidate inhibitors on mitochondrial O2 consumption in permeabilized asexual P. falciparum parasites can be tested in real time.
• Through the sequential injection of inhibitors and substrates into the assay, the molecular targets of candidate inhibitors in the ETC can, in part, be determined.
• The assay is applicable for both drug discovery approaches and enquiries into a fundamental aspect of parasite mitochondrial biology.
Graphical overview
Seahorse assay experimental workflow. Prior to the assay, coat the cell culture microplate with Cell-Tak to help adhere the parasites to the wells; hydrate the cartridge wells to ensure proper sensor functionality and design the assay template using the Agilent Seahorse Wave Desktop software (Analyze Seahorse data files, Seahorse Wave desktop software|Agilent). On the day of the assay, prepare the inhibitors/substrates that are to be injected into the ports. Then, separate 3 × 108 trophozoite-stage parasites from the uninfected red blood cells (RBCs) and ring-stage parasites using a MACS® magnetic column. Check the purity of the parasites with Giemsa-stained smears. Determine the concentration of infected RBCs in the sample using a hemocytometer and dilute to approximately 5 × 107 parasites per milliliter. Treat infected RBCs with saponin to permeabilize the host cell membrane and seed approximately 5 × 106 parasites (100 μL) per well in mitochondria assay solution (MAS) buffer. Supplement MAS buffer with digitonin to permeabilize the parasite plasma membrane. Load the ports with the prepared inhibitors/substrates and run the assay using a Seahorse XFe96 analyzer. Once the assay is completed, analyze the data using the Wave desktop software. Further data processing can be done using statistical analysis software.
Keywords: Plasmodium falciparum Malaria Seahorse XFe96 Mitochondria Electron transport chain Antimalarial drugs Drug discovery
Background
Plasmodium species cause malaria in humans and were responsible for 247 million malaria cases and 619,000 deaths in 2021 (WHO report, 2022). The emergence of parasites resistant to frontline antimalarial drugs has contributed to the increase in malaria cases and deaths in recent years (Conrad and Rosenthal, 2019; Ippolito et al., 2021). Hence, it is important to look beyond the currently available antimalarial drugs for new treatments. The mitochondrion of Plasmodium is a validated drug target, and some common antimalarial compounds interfere with its electron transport chain (ETC) (Hikosaka et al., 2015; Ke and Mather, 2017).
The ETC is located in the inner mitochondrial membrane of eukaryotes and facilitates the transfer of electrons through a series of redox centers to O2. Electrons are sourced from the oxidation of substrates such as malate, succinate, NADH, glycerol 3-phosphate, and dihydroorotate in the mitochondrion (Hayward and van Dooren, 2019). These electrons are donated to coenzyme Q (CoQ). Reduced CoQ docks at the cytochrome bc1 complex (also known as Complex III). Complex III facilitates the transfer of electrons to an electron carrying protein, cytochrome c (CytC). From CytC, electrons are donated to the cytochrome c oxidase complex (Complex IV), where the reduction of O2, the final electron acceptor of the ETC, occurs. The transfer of electrons through Complexes III and IV is coupled to the net movement of protons from the mitochondrial matrix to the mitochondrial intermembrane space (Figure 1). The proton motive force that is established across the inner mitochondrial membrane by the ETC is critical for the synthesis of ATP by the ATP synthase complex (Figure 1).
Figure 1. Mitochondrial electron transport chain. Electrons from the oxidation of a range of different substrates in the mitochondrion are donated to coenzyme Q (CoQ). CoQ donates these electrons to Complex III, from where they are donated via cytochrome c (CytC) to Complex IV, where electrons ultimately reduce O2 to form H2O. Electron transport events are indicated by arrows. The passage of electrons through Complexes III and IV is coupled to the net transport of protons (H+) from the mitochondrial matrix to the intermembrane space, generating an H+ gradient that can be harnessed by ATP synthase to generate ATP. DHODH, dihydroorotate dehydrogenase; G3PDH, glycerol 3-phosphate dehydrogenase; MQO, malate: quinone oxidoreductase; NDH2, type II-NADH dehydrogenase; FADH2, Dihydroflavine-adenine dinucleotide; FAD, Flavin adenine dinucleotide. The inset indicates the position of the enlarged portion within the mitochondrion. Created with BioRender.com.
The P. falciparum ETC varies considerably from the ETC of their mammalian hosts (Hayward and van Dooren, 2019). For example, the P. falciparum ETC has a single subunit “type II” NADH dehydrogenase instead of the multi-subunit Complex I found in mammals and a malate:quinone oxidoreductase enzyme that feeds electrons from malate oxidation directly into the ETC (Figure 1; Rajaram et al., 2022). Complex III is a validated drug target in apicomplexans, and the compounds that inhibit Complex III often target either (or in rare cases both) of the CoQ binding sites in the complex. Structural differences in the CoQ binding sites of parasite Complex III compared with human Complex III likely account for the high degree of selectivity of inhibitors like atovaquone against the parasite complex (Srivastava et al., 1999; Fisher et al., 2020). Mutations in the CoQ binding sites of parasite Complex III can confer clinically relevant levels of resistance against compounds like atovaquone (Staines et al., 2018). It is therefore of interest to identify novel inhibitors of Complex III or the other components of the P. falciparum ETC, ideally ones that remain potent against parasites already resistant to existing ETC inhibitors.
Measuring cellular O2 consumption is a powerful means of assessing ETC function. We have recently developed a suite of Seahorse XFe96 flux analyzer assays to measure mitochondrial O2 consumption rates in the apicomplexan parasite Toxoplasma gondii, a species from the same phylum as Plasmodium (Hayward et al., 2022). We have now established similar assays to enable the measurement of mitochondrial O2 consumption rates in permeabilized asexual-stage Plasmodium falciparum parasites and describe them in this protocol (Graphical overview; General note 1). The assay can be used to screen compound libraries to identify ETC inhibitors in P. falciparum. The assay can aid in determining the molecular target of identified inhibitors in the ETC and the effectiveness of novel inhibitors against drug-resistant parasite strains (Hayward et al., 2023; Nguyen et al., 2023). The assay can also be used to obtain molecular insights into the functionality of the P. falciparum ETC.
Materials and reagents
175 cm2 tissue culture flasks (Sarstedt, catalog number: 83.3912)
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Sigma-Aldrich, catalog number: H3375-250G)
5% (w/v) digitonin (Thermo Fisher Scientific, InvitrogenTM, catalog number: BN 20061)
Albumax II (Gibco, catalog number: 11021-045-1kg)
Atovaquone (Sigma-Aldrich, catalog number: A7986-10MG)
Cell-Tak (Corning, catalog number: 354241)
D-glucose (Sigma-Aldrich, catalog number: G7021)
D-mannitol (Sigma-Aldrich, catalog number: M9546-250G)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D8418-250G)
Ethylene glycol-bis(2-amino-ethylether)-N,N,N,N’-tetra acetic acid (EGTA) (Sigma-Aldrich, catalog number: E3889-25G)
Fatty acid-free bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A3803-100G)
General plastic consumables [sterile Eppendorf tubes, sterile falcon tubes (15, 50 mL), sterile pipette tips (20, 200, and 1,000 μL), sterile serological pipettes (10, 25 mL); Sarstedt or Corning]
Gentamicin (Invitrogen, catalog number: 15710-072)
Hypoxanthine (Sigma-Aldrich, catalog number: H-9636)
L-ascorbic acid (Sigma-Aldrich, catalog number: A5960-25G)
L-malic acid (Sigma-Aldrich, catalog number: M1000-100G)
Magnesium chloride hexahydrate (MgCl2·6H2O) (Sigma-Aldrich, catalog number: M2670-100g)
Milli-Q water (fresh from any available purification system)
N,N,N,N’-tetramethyl-p-phenylenediamine (TMPD) (Sigma-Aldrich, catalog number: T7394-5G)
Potassium chloride (Sigma-Aldrich, catalog number: 7447-40-7, 500G)
Potassium phosphate monobasic (KH2PO4) (ChemSupply, catalog number: PA009-500G)
RPMI-HEPES with Glutamax (Thermo Fisher ScientificTM, catalog number: 72400120)
Sodium azide (AJAX-Finechem Univar, catalog number: AJA1222-100G)
Sodium chloride (Sigma-Aldrich, catalog number: S5886-5KG)
Sodium dihydrogen phosphate monohydrate (AnalaR, catalog number: 102454R)
Sorbitol (Sigma-Aldrich, catalog number: 50-70-4)
Sucrose (Sigma-Aldrich, catalog number: S9378-5KG)
Ascorbate stock (200 mM) (see Recipes)
Albumax II (5% w/v) (see Recipes)
Atovaquone stock (25 mM) (see Recipes)
Complete culture medium (see Recipes)
Malate stock (500 mM) (see Recipes)
MAS buffer (3× stock) (see Recipes)
MAS buffer (1× working solution) (see Recipes)
Phosphate buffered saline (10×) (see Recipes)
Saponin (0.15% w/v) (see Recipes)
Sodium azide stock (100 mM) (see Recipes)
Sodium bicarbonate (0.1 M) (see Recipes)
Sorbitol (5% w/v) (see Recipes)
TMPD stock (500 mM) (see Recipes)
Biological material
Plasmodium falciparum cells (wild type 3D7)
Recipes
Ascorbate stock (200 mM)
Dissolve 352 mg of L-ascorbic acid in 5 mL of 1× MAS buffer. Adjust pH to 7.4 with potassium hydroxide (KOH) and adjust final volume with 1× MAS buffer to 10 mL. Aliquot ascorbate stock (500 μL/aliquot) and store at -20 °C for long-term storage.
Albumax II (5% w/v)
Dissolve 25 g of Albumax II in 500 mL of RPMI medium. Filter sterilize using the Millipore Stericup Vacuum Driven Disposable Filtration System. Aliquot Albumax II stock (50 mL/aliquot) and store at -20 °C for long-term storage.
Atovaquone stock (25 mM)
Dissolve 10 mg of atovaquone in 1.09 mL of DMSO. Aliquot atovaquone stock (50 μL/aliquot) and store at -20 °C for long-term storage.
Complete culture medium
Prepare complete culture medium from RPMI 1640-HEPES with Glutamax supplemented with the cell culture media components shown below. Store complete culture medium at 4 °C and pre-warm to 37 °C before use.
Components Final concentration Volume
RPMI medium - 500 mL
Additional glucose (2.5 M) 10 mM 2 mL
Hypoxanthine (200 mM) 480 μM 1.2 mL
Gentamicin (10 mg/mL) 20 μg/mL 1 mL
5% Albumax solution 0.375% w/v 37.5 mL
Heat-inactivated human serum 2.5% v/v 12.5 mL
Malate stock (500 mM)
Dissolve 670.45 mg of L-malic acid in 5 mL of MAS buffer. Adjust pH to 7.4 with KOH and adjust the final volume with MAS buffer to 10 mL. Aliquot malate stock (1.5 mL/aliquot) and store at -20 °C for long-term storage.
MAS buffer (3× stock)
Weigh out the components (listed below) and dissolve in 50–75 mL of Milli-Q water by placing the solution into a water bath and/or use a magnetic stirrer. Adjust pH of the solution to 7.4 using KOH. Bring the final volume to 125 mL. Filter sterilize for long-term storage.
Components Final concentration Weight
Mannitol 660 mM 15.03 g
Sucrose 210 mM 8.99 g
KH2PO4 30 mM 0.51 g
MgCl2 15 mM 0.381 g
HEPES 6 mM 0.1785 g
EGTA 3 mM 0.1425 g
MAS buffer (1× working solution)
On the day of the experiment, mix 15 mL of 3× MAS buffer with 30 mL of Milli-Q water, to get 45 mL of 1× MAS buffer. Add 90 mg of fatty acid-free BSA and 2.25 mL of 0.5 M malate.
Phosphate buffered saline (10×)
Components Weight
NaCl 80 g
KCl 2 g
Na2HPO4 14.4 g
KH2PO4 1.9 g
Dissolve the compounds in 800 mL of Milli-Q water. Adjust pH of the solution to 7.4 using HCl. Bring the final volume to 1 L. Sterilize the solution by autoclaving for 20 min at 15 psi. Store at room temperature. When needed, mix 100 mL of 10× PBS with 900 mL of Milli-Q water to get 1× PBS.
Saponin (0.15% w/v)
Dissolve 0.15 g of saponin in 100 mL of PBS. Filter-sterilize the solution using a 0.2 μm filter. Store at 4 °C for long-term storage. When needed, mix 1 mL of 0.15% saponin with 2 mL of PBS to obtain a 0.05% (w/v) saponin working solution.
Sodium azide stock (100 mM)
Dissolve 65.1 mg of sodium azide in 10 mL of MAS buffer. Aliquot sodium azide stock (500 μL/aliquot) and store at -20 °C for long-term storage.
Sodium bicarbonate (0.1 M)
Dissolve 84 mg of sodium bicarbonate in approximately 8 mL of Milli-Q water. Adjust pH to 8.0 and make up the volume to 10 mL. Filter-sterilize using a 0.2 μm filter. Store at room temperature.
Sorbitol (5% w/v)
Dissolve 5 g of sorbitol in 100 mL of Milli-Q water. Filter-sterilize the solution using a 0.2 μm filter. Store at 4 °C for long-term storage.
TMPD stock (500 mM)
Dissolve 82.2 mg of TPMD in 1 mL of ethanol. Aliquot TMPD solution (50 μL/aliquot) and store at -20 °C for long-term storage.
Equipment
4 °C fridge and -20 °C freezer
Tabletop centrifuge for 15 and 50 mL tubes (Beckman Coulter, model: Allegra X-15R)
Tabletop centrifuge for microcentrifuge tubes (Thermo Scientific, model: Heraeus Pico 17)
Improved Neubauer hemocytometer (Hirschmann, model: 8100104)
Humidified 37 °C non-CO2 incubator
MACS® magnetic column (CS columns, Miltenyi Biotec, catalog number: 130-041-305) with a sterile 3-way stopclock
SuperMACS II separator (Miltenyi Biotec, catalog number: 130-044-104)
Multichannel pipettes (30–300 μL) (Gilson)
pH meter (Cyberscan pH510)
Pipettes (0.2–2, 2–20, 20–200, and 100–1,000 μL) (Gilson)
Reagent reservoirs
Seahorse XFe96 Analyzer (Agilent)
Seahorse XFe96 FluxPak (XFe96 cell culture microplate, XFe96 sensor cartridge and utility plate, XF Calibrant Solution) (Agilent, catalog number: 102416-100)
Standard inverted light microscope (Olympus CKX41), fitted with 20× objective lense
Biological safety cabinet (Safemate 1.2 vision, Edwards Group)
Water bath, 37 °C (WB7, Ratek)
Zip-lock bags
Software
Wave Desktop Software, version 2.6.3 (for running the Seahorse XFe96 assay)
GraphPad Prism 9 (for statistical analyses of the results)
Procedure
Parasite culturing
Maintain P. falciparum parasites in complete culture medium at 4% hematocrit and approximately 3% parasitaemia in cell culture plates and flasks (Maier and Rug, 2013).
Every second day, check the parasitaemia using Giemsa staining technique; replace the culture with fresh complete culture medium pre-warmed to 37 °C and dilute the parasite cultures to 0.2% parasitaemia. Gas the culture flask with malaria gas mixture of 1% O2, 5% CO2, and 94% N2 for 30 s; close the cap of the flask and incubate at 37 °C.
Maintain a synchronized P. falciparum culture by treating the parasites with sorbitol according to Lambros and Vanderberg (1979).
Expand the cell culture to approximately 3 × 108 trophozoite-infected RBCs and proceed with the Seahorse XFe96 assay (General note 2).
XF96 cell culture microplate pre-treatment (at least 1 day prior to the experiment)
Figure 2 depicts various parts of the Seahorse XFe96 sensor cartridge used in the protocol. The XFe96 analyzer uses only 96-well plates. A 24-well plate format is available for use with a Seahorse XFe24 analyzer, with a previously published protocol describing the analysis of O2 consumption rates in P. falciparum using this approach (Sakata-Kato and Wirth, 2016).
Figure 2. Parts of the Seahorse XFe96 sensor cartridge. A) Side view shows the bottom of the sensor probes attached to the green sensor cartridge. B) Top view shows an example of the four injection ports above each of the 96 wells. The inset shows ports A–D and the top of the sensor probe. Created with BioRender.com.
In order to immobilize Plasmodium falciparum cells on the Agilent Seahorse XFe96 plate during the experiment, coat each well with Cell-Tak, according to the manufacturer’s protocol, 1–5 days prior to the experiment. Briefly, prepare Cell-Tak in 0.1 M sodium bicarbonate at a final concentration of 22.12 μg/mL (General note 3); 2.5 mL is enough for coating one 96-well plate. Dilute Cell-Tak right before use, mix thoroughly, and dispense immediately.
Dispense 25 μL of Cell-Tak in each well of the XFe96 cell culture microplate; 25 μL is usually sufficient to cover the entire bottom of the well, but care must be taken to spread Cell-Tak throughout the well.
Incubate the plate for 20 min at room temperature. Aspirate the solution. Invert and pat the plate to remove any residual Cell-Tak solution.
Wash each well with 200 μL of Milli-Q water, twice, to reduce any trace sodium bicarbonate.
Discard the water and air-dry the wells under sterile conditions. The plate can be left in the cell culture hood for drying.
Store the XFe96 cell culture microplate in a zip-lock bag in the fridge (4 °C) for up to one week.
On the day of the experiment, warm the plate to room temperature in a biological safety cabinet before seeding cells.
Hydrate the sensor cartridge (1 day prior to the experiment)
The sensor cartridge consists of solid-state sensor materials, which detect changes in pH and O2 concentration during the experiment. Prepare the sensors by hydrating the sensor cartridge according to Agilent’s user guide as follows. Open the XFe96 FluxPak (cartridge + utility plate package) and place the cartridge upside down (sensor end up) next to the utility plate. Do not touch the sensors. Fill all the wells of the utility plate with 200 μL of Milli-Q water.
Place the cartridge back on the utility plate and make sure that the sensors are submerged completely. Incubate the sensor cartridge and utility plate in a 37 °C humidified incubator overnight.
On the day of the assay, remove the cartridge and place it upside down (sensor end up). Discard the water from the utility plate and fill all the wells with 200 μL of XF Calibrant pre-warmed to 37 °C.
Place the cartridge back in the utility plate and move the cartridge up and down, twice, to remove any trapped air bubbles from the sensor.
Incubate the utility plate with the sensor cartridge and calibrant solution in a 37 °C humidified incubator for at least 1 h prior to the start of the experiment.
Wave program template preparation
Set up Wave desktop software.
Under the group definitions tab, define the injection strategies (Figure 3A). An example is shown below.
Port A, inhibitors/atovaquone (5 μM).
Port B, TMPD (0.2 mM) and ascorbic acid (2 mM).
Port C, sodium azide (NaN3; 10 mM).
Configure the plate map by assigning individual groups to different conditions of the experiment (Figure 3B). Make sure to include background wells (measurement without cells). A–D represents different candidate P. falciparum ETC inhibitors. Atovaquone is used as the positive control. Control wells have parasites without drug treatment (i.e., solvent control injected from Port A). Background wells do not have parasites. Four technical replicates are included for each condition.
Under the protocol tab, click on injection and specify the timing and repeats of the mixing and measurement cycles for each injection (Figure 3C). e.g., eight cycles of measurement after inhibitor injection (five cycles of measurements otherwise). Each cycle of measurement has:
20 s of mixing,
1 min of waiting,
2.5 min of measuring.
Figure 3. Designing a Seahorse XFe96 experimental template in the Wave software. A) Under Group Definitions tab, fill in the injection strategies as follows. Port A: Inhibitors/atovaquone; Port B: TMPD; Port C: Sodium azide. B) Under the Plate Map tab, customize the plate map according to your group definition. Ensure background wells (wells without cells) are included. C) Under the Protocol tab, specify the timing and substrates to be injected from a specific port. Specify the number of mixing and measurement cycles as follows. After inhibitor injection 8 cycles (5 cycles for all other substrate injection); 20 s mixing; 1 min waiting; 2.5 min measuring.
Save this template. On the day of the experiment, open the template and click on the run assay tab to start the assay.
Inhibitor/substrate preparation
Prepare 45 mL of 1× MAS buffer (see Recipes). Pre-warm 1× MAS buffer, PBS, and saponin in a water bath (37 °C) until use.
Prepare the substrates/inhibitors/controls that will be injected in the ports of the sensor cartridge before parasite preparations (General note 4). For the model experiment, we used:
8× atovaquone/inhibitors solution. Prepare a 40 μM solution to obtain a final concentration of 5 μM following injection into the wells during the assay. To make up 40 μM stocks, add 1.4 μL of 10 mM inhibitor to 348.6 μL of MAS buffer, or add 1.1 μL of 25 mM atovaquone to 698.9 μL of MAS buffer.
9× TMPD/ascorbate solution. Prepare a solution containing 1.8 mM TMPD and 18 mM ascorbate to obtain a final concentration following injection of 0.2 mM TMPD/2 mM ascorbate. Add 6.5 μL of 500 mM TMPD and 162 μL of 200 mM ascorbate to 1,631.5 μL of MAS buffer.
10× sodium azide solution. Prepare a solution of 100 mM NaN3 to obtain a final concentration following injection of 10 mM NaN3. Add 300 μL of 1 M NaN3 to 2,700 μL of MAS buffer (General note 5).
Parasite preparation
Assemble the MACS® magnetic column with three-way stopcock inside the biological safety cabinet. Pre-wash the column by passing 30 mL of 100% ethanol, 30 mL of 80% ethanol, 30 mL of PBS (pre-warmed to 37 °C), and 30 mL of RPMI-HEPES with Glutamax, pre-warmed to 37 °C (General note 6).
Enrich the culture for trophozoite-stage parasites by passing 3 × 108 trophozoite-infected RBCs drop-by-drop through a MACS® magnetic column placed in the magnetic field of a SuperMACS II separator (Ridgway et al., 2021). The volume of the culture is adjusted to yield a hematocrit of approximately 15% before passing through the column. Uninfected and ring-stage infected RBCs will pass through the column, while RBCs infected with mature-stage parasites (trophozoites and schizonts) will be retained.
Remove any residual uninfected red blood cells or ring-stage parasites retained in the column by passing 30–50 mL of complete culture medium pre-warmed to 37 °C through the column.
Remove the column from the magnet and elute RBCs infected with mature-stage parasites in 20–30 mL of complete culture medium.
Determine the parasite-infected RBC yield using a hemocytometer. Check the purity of the parasites in the eluate using Giemsa-stained blood smears.
Free 2 × 108–3 × 108 trophozoite-stage parasites from erythrocytes by treating infected RBCs with 1 mL of pre-warmed 0.05% (w/v) saponin in PBS at 37 °C for 3 min. Wash the cells with pre-warmed PBS three times or until the supernatant is no longer red in color (i.e., until the hemoglobin is completely removed), pelleting the parasites by centrifugation at 2,000× g for 2 min at room temperature after each wash.
The parasite yield should be approximately 2 × 108–3 × 108 cells. Adjust the parasite concentration to 5 × 107 parasites/mL in mitochondrial assay solution (MAS) buffer containing 0.002% (w/v) digitonin. Digitonin permeabilizes the parasite plasma membrane. Pre-heat digitonin solution to 62 °C (to dissolve it) before adding. Parasites start losing viability after detergent treatment and it is advisable to start the O2 consumption rate (OCR) measurements within one hour of the detergent treatment.
Seed 100 μL of the permeabilized parasites per well in the Cell-Tak-coated XFe96 cell culture plate. This results in a density of 5 × 106 cells per well. Centrifuge at 800× g for 10 min at room temperature with the centrifuge brake set to low. This step ensures the parasites adhere to the bottom of the wells. Check the plates under the microscope to ensure that a monolayer of cells has formed (see Figure 4).
Figure 4. Outline of P. falciparum in a cell monolayer. Cell-Tak-coated P. falciparum cell monolayer in XFe96 cell culture plate, after centrifuging the plate with low brake.
Carefully add 75 μL of MAS buffer to the side of the wells at an angle, without agitating and detaching the adhered parasites. Check the plates under the microscope again to confirm that the monolayer of cells has not been disturbed.
Transfer the plate to an incubator (37 °C) at atmospheric CO2 to degas the plate of excess CO2 levels, and store until use. This step is crucial to effectively degas the plastic material of any CO2 (Thangaraj et al., 2018). Degassing adjusts the CO2 in the assay plate to (low) atmospheric CO2 levels, ensuring pH values of the assay solutions are not influenced by changes in CO2 levels during the assay.
OCR measurements
At least one hour prior to the experiment, turn on the XFe96 flux analyzer and the temperature controller to calibrate the instrument.
Open the prepared experiment template in Wave software and export it as a design file (.asyt) to run the assay.
Carefully load 25 μL of each inhibitor/substrate into the injection ports after the cartridge has been placed in the utility plate. For the assay we describe in this manuscript: Port A—Compounds (8× the final concentration), port B—TMPD (9× the final concentration), port C—NaN3 (10× the final concentration) port D—empty in this experiment. To load the solution into the port, insert the pipette tip to near the bottom of the port. Lift the pipette tip slightly and angle the tip alongside the wall. Slowly dispense the inhibitor/substrate into the port (General note 7).
Ensure all ports are filled, including ports of blank wells and ports of unused wells without cells. In the sample experiment, assay ports A, B, and C of all wells must be filled to enable compound/substrate dispensing. Port D, which is not used, can remain empty.
Click Run assay in the Wave software and, when prompted, load the cartridge (filled with the substrates/inhibitors to be injected) with the utility plate onto the plate holder. Make sure to remove the lid from the utility plate and place the plate in the right orientation onto the plate holder.
The sensor calibration takes approximately 25–30 min.
Once the calibration is finished, the utility plate is automatically ejected. The sensor cartridge is retained inside the XFe96 analyzer. Replace the utility plate with the cell plate containing the adhered parasites (again without the lid) and resume the run.
Once the experiment is complete, the cell culture plate is ejected. Remove the plate and the sensor cartridge and examine the injection ports to verify that all compounds were injected (General note 8).
The results file (.asyr) is automatically saved on the computer. Save the result file onto a USB drive to transfer to another computer with the analysis software.
Discard the cell culture plate with the cartridge containing the parasites and the substrates safely and appropriately (General note 9).
Data analysis
Open the results file, using Wave software 2.6.3 or another version.
Click on add view and select overview to view the results and plate map.
Make sure the background correction box is ticked. Background wells lack parasites and, once background correction box is ticked, the OCR values from these background wells will be subtracted from the experimental wells.
Occasionally, substrates from one or more ports may not dispense properly (e.g., fail to inject or inject prematurely). This will lead to errors in the data. In addition to checking the injection ports following the experiment (General note 8), analyze the data from each group on the plate map and look for instances where premature injection of substrates from some ports may have occurred. If you have a strong reason to believe that improper injection has occurred into a well, you can consider excluding the data from that particular well from the analysis by unselecting it (General note 10).
The result indicates the effect of different substrates or inhibitors on the OCR of P. falciparum over time (Figure 5A). The results can be plotted onto a graph as time (minutes) vs. OCR (pmol/min) (Figure 5B), typically as the average OCR value of the technical replicates at each time point in the plate. From these data, it is possible to calculate the extent of OCR inhibition caused by the candidate inhibitor, the extent of TMPD-dependent OCR recovery, and NaN3-dependent OCR inhibition. To ensure confidence in the data, independent biological repeats should be performed on different days.
Figure 5. Model data based on the injection strategies performed. A) Schematic depicting the effects of the inhibitors and substrates on the mitochondrial electron transport chain. B) Initially, malate-dependent O2 consumption rate (OCR) is measured to establish the basal OCR. The compound of interest, either a putative inhibitor or atovaquone (positive control), is then injected into the wells to measure their effect on the OCR. The OCR will be diminished if the inhibitor affects the mitochondrial electron transport chain (ETC). Following several measurements, TMPD is injected into the wells. TMPD donates electrons to cytochrome c and thereby bypasses Complex III. If TMPD injection restores the OCR, it implies that the test compound is inhibiting OCR upstream of cytochrome c. After a few measurements, the Complex IV inhibitor sodium azide (NaN3) is injected into the wells. If NaN3 injection inhibits the OCR, it confirms that the TMPD-dependent OCR recovery is dependent on Complex IV activity. Created with BioRender.com.
Click on export to export the results to statistical analysis software like GraphPad Prism or Microsoft Excel to further process the data.
Further uses of the Seahorse XFe96 assay
In the experiment described in this manuscript, we test candidate ETC inhibitors for their ability to inhibit mitochondrial OCR in P. falciparum, and whether these inhibitors target up- or downstream of cytochrome c in the ETC. The Seahorse XFe96 assay is a versatile one and can be used in a range of other experiments. For example, testing the extent of mitochondrial OCR inhibition at a range of compound concentrations enables determination of the potency of those inhibitors on the ETC [e.g., enables calculation of the half maximal effective concentration (EC50) of the compound on OCR (Hayward et al., 2023)], and the extent to which this changes in drug-resistant parasites. The Seahorse XFe96 assay can also be used for screening large libraries for putative ETC inhibitors, as we have done previously for similar assays in the related parasite Toxoplasma gondii (Hayward et al., 2023). The presence of four injection ports on the 96-well plate enables several hundred compounds to be screened using a single Seahorse XFe96 assay. The Seahorse XFe96 assay we describe here also has uses beyond compound screening strategies. For example, it can be used to measure ETC impairment in mutant parasite strains defective in aspects of mitochondrial biology, or to gain insights into fitness costs associated with mutations conferring resistance to ETC inhibitors.
Validation of protocol
Jenni, A. Hayward, F. Victor Makota, Daniela Cihalova, Rachel A. Leonard, Esther Rajendran, Soraya M. Zwahlen, Laura Shuttleworth, Ursula Wiedemann, Christina Spry, Kevin J. Saliba, Alexander G. Maier, Giel G. van Dooren: A screen of drug-like molecules identifies chemically diverse electron transport chain inhibitors in apicomplexan parasites, PLOS Pathogens 19(7): e10111517.
Figure 6. Most of the candidate ETC inhibitors target the ETC upstream of cytochrome c in P. falciparum parasites.
Table 3. Inhibitory activities of MMV Pathogen Box compounds against O2 consumption rate in T. gondii and P. falciparum.
William Nguyen, Madeline G. Dans, Iain Currie, Jon Kyle Awalt, Brodie L. Bailey, Chris Lumb, Anna Ngo, Paola Favuzza, SaiShyam Ramesh, Jocelyn Pennington, Kate E. Jarman, Alexander G. Maier, Giel G. van Dooren, Tony Papenfuss, Sergio Wittlin, Jake Baum, Delphine Baud, Stephen Brand, Paul F. Jackson, Alan F. Cowman, and Brad E. Sleebs: 7-N-Substituted-3-Oxadiazole Quinolones with Potent Antimalarial Activity Target the Cytochrome bc1 Complex; ACS Infectious Diseases. doi: https://doi.org/10.1021/acsinfecdis.2c00607
Figure 5. P. falciparum oxygen consumption rate (OCR) assay.
General notes and troubleshooting
General notes
The current protocol to measure mitochondrial O2 consumption rates in P. falciparum has key differences from the ones we established in T. gondii. P. falciparum preparation and permeabilization requires numerous steps that are detailed in section F. P. falciparum parasites are particularly sensitive to the depletion of carbon sources, which means that, unlike in T. gondii, we need to maintain Plasmodium in a carbon source at all times. In contrast to a previous study using a Seahorse XFe24 analyzer approach to measure O2 consumption in P. falciparum (Sakata-Kato and Wirth, 2016), we find that permeabilization of the parasite plasma membrane with low concentrations of digitonin is critical for enabling the robust measurement of mitochondrial O2 consumption rates in the parasite.
Make sure the parasites are synchronized and the majority is at the late-trophozoite stage on the day of the experiment.
Cell-Tak concentration varies from batch to batch and the final dilution may need to be adjusted depending on the batch. For the model experiment, we used Cell-Tak Lot # 0055009 (1.58 mg/mL) and diluted 35 μL of Cell-Tak in 2.5 mL of 0.1 M sodium bicarbonate solution (final concentration 22.1 μg/mL).
The time from initial parasite preparation to compound injection from the ports is substantial. To best maintain parasite viability, prepare substrate and inhibitor compounds (section E) before commencing parasite preparation (section F).
The substrates from port A are injected first, followed by substrates in port B, C, and D, according to the constant concentration protocol described in the manufacturer’s instructions. Once injected, the added MAS buffer volume in the well dilutes the substrate. To account for the dilution, the concentration of the substrates being injected increases proportionally with each injection. E.g., 8× the desired final concentration for compounds injected from port A, 9× the concentration for port B, 10× the concentration for port C, and 11× for port D.
Sterilizing the column with 25 mL of 100% and 80% ethanol is necessary if the magnetic columns are reused. If a new column is used, the ethanol washes are not required.
Make sure to dispense the correct substrate or inhibitor into the appropriate port, as it is very difficult to completely remove the sample if a wrong compound is dispensed into a port. It will also make the data unreliable if traces of different compounds are mixed. If a wrong compound is dispensed into a port, exclude those wells from the analysis altogether.
One way to verify whether all the ports were injected during the course of the assay is to take a photo of the cartridge before and after the experiment and then checking if all the ports have been discharged during the assay.
The Seahorse assay plates contain reagents including TMPD, sodium azide, and potentially some of the candidate inhibitors that are toxic to humans and/or the environment, and also contain potentially infectious P. falciparum parasites. Hence, following the experiment, the parasites and the substrates should be discarded in accordance with local policies for disposing hazardous materials.
Improper injections are not common. To minimize improper injections, care should be taken not to damage the port with the pipette tip when dispensing the substrate into the port. Also, the plate should be handled gently after dispensing the substrates into the ports. Finally, including four technical replicates per experimental condition can aid in ensuring sufficient confidence in the data in instances where improper compound injection occurs, and particular wells are excluded from the analysis.
Acknowledgments
We are thankful to the Australian Red Cross for providing human RBCs and serum. This work was supported by the Research School of Biology (RSB) International Ph.D. Scholarship to S.R., an Australian National Health and Medical Research Council Ideas grant (GNT1182369) to G.v.D and A.G.M, and a Research School of Biology Innovation grant to G.v.D., A.G.M., E.R. and D.C.
Graphical overview created with BioRender.com.
Competing interests
The authors declare that they have no competing interests.
Ethical considerations
The use of human erythrocytes was approved as part of the Human Ethics committee protocol HEC 2017/351 of the Australian National University.
References
Conrad, M. D. and Rosenthal, P. J. (2019). Antimalarial drug resistance in Africa: the calm before the storm? Lancet Infect. Dis. 19(10): e338–e351.
Fisher, N., Meunier, B. and Biagini, G.A., (2020). The cytochrome bc1 complex as an antipathogenic target. FEBS Letters 594(18): 2935–2952.
Hayward, J. A. and van Dooren, G. G. (2019). Same same, but different: Uncovering unique features of the mitochondrial respiratory chain of apicomplexans. Mol. Biochem. Parasitol. 232: 111204.
Hayward, J. A., Makota, F. V., Cihalova, D., Leonard, R.A., Rajendran, E., Zwahlen, S.M., Shuttleworth, L., Wiedemann, U., Spry, C., Saliba, K. J., Maier, A. G., et al. (2023). A screen of drug-like molecules identifies chemically diverse electron transport chain inhibitors in apicomplexan parasites. PLOS Pathog. 19(7): e1011517.
Hayward, J. A., Rajendran, E., Makota, F. V., Bassett, B. J., Devoy, M., Neeman, T. and van Dooren, G.G. (2022). Real-time analysis of mitochondrial electron transport chain function in Toxoplasma gondii parasites using a Seahorse XFe96 extracellular flux analyzer. Bio Protoc. 12(1): e4288.
Hikosaka, K., Komatsuya, K., Suzuki, S. and Kita, K. (2015). Mitochondria of malaria parasites as a drug target. An Overview of Tropical Diseases.
Ippolito, M. M., Moser, K. A., Kabuya, J. B. B., Cunningham, C. and Juliano, J. J. (2021). Antimalarial drug resistance and implications for the WHO global technical strategy. Curr. Epidemiol. Rep. 8(2): 46–62.
Ke, H. and Mather, M. W. (2017). +Targeting Mitochondrial Functions as Antimalarial Regime, What Is Next? Curr. Clin. Microbiol. Rep. 4(4): 175–191.
Lambros, C. and Vanderberg, J. P. (1979). Synchronization of Plasmodium falciparum Erythrocytic Stages in Culture. J. Parasitol. 65(3): 418–420.
Maier, A. G. and Rug, M. (2013). In vitro Culturing Plasmodium falciparum Erythrocytic Stages. Methods Mol. Biol. 923: 3–15.
Nguyen, W., Dans, M. G., Currie, I., Awalt, J. K., Bailey, B. L., Lumb, C., Ngo, A., Favuzza, P., Palandri, J., Ramesh, S., et al. (2023). 7-N-Substituted-3-oxadiazole Quinolones with Potent Antimalarial Activity Target the Cytochrome bc1 Complex. ACS Infect. Dis. 9(3): 668–691.
Rajaram, K., Tewari, S. G., Wallqvist, A. and Prigge, S. T. (2022). Metabolic changes accompanying the loss of fumarate hydratase and malate–quinone oxidoreductase in the asexual blood stage of Plasmodium falciparum. J. Biol. Chem. 298(5): 101897.
Ridgway, M., Cihalova, D. and Maier, A. (2021). Sex-specific Separation of Plasmodium falciparum Gametocyte Populations. Bio Protoc 11(11): e4045.
Sakata-Kato, T. and Wirth, D. F. (2016). A Novel Methodology for Bioenergetic Analysis of Plasmodium falciparum Reveals a Glucose-Regulated Metabolic Shift and Enables Mode of Action Analyses of Mitochondrial Inhibitors. ACS Infect. Dis. 2(12): 903–916.
Srivastava, I. K., Morrisey, J. M., Darrouzet, E., Daldal, F. and Vaidya, A. B. (1999). Resistance mutations reveal the atovaquone-binding domain of cytochrome b in malaria parasites. Mol. Microbiol. 33(4): 704–711.
Staines, H. M., Burrow, R., Teo, B. Y., Chis Ster, I., Kremsner, P. G. and Krishna, S. (2018). Clinical implications of Plasmodium resistance to atovaquone/proguanil: a systematic review and meta-analysis. J. Antimicrob. Chemother. 73(3): 581–595.
Thangaraj, A., Periyasamy, P., Liao, K., Bendi, V. S., Callen, S., Pendyala, G. and Buch, S. (2018). HIV-1 TAT-mediated microglial activation: role of mitochondrial dysfunction and defective mitophagy. Autophagy 14(9): 1596–1619.
World Health Organization (2022). World malaria report 2022. World Health Organization (8 Dec 2022, ISBN: 978 92 4 0066489 8).
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Cell Biology > Cell metabolism > Respirometry
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Retraction Notice: Paper Lateral Flow Biosensor for Nodavirus Reverse Transcribed RNA Detection doi: 10.21769/BioProtoc.3711
DT Dimitra K. Toubanaki
Evdokia Karagouni
Published: Sep 20, 2023
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This article (10.21769/BioProtoc.3711) has been retracted at the request of the authors of the paper due to unforeseen conflicts concerning intellectual property rights. At the time of publication, the authors were unaware of relevant patents, International Patent EP1436420A2 and National Patent GR1003966B, that encompass certain methodologies used in this study. The intellectual property holder, not a co-author, has raised concerns regarding proprietary information use. Both authors agree with the retraction and affirm the scientific integrity and quality of the protocol. The retraction is for legal reasons and does not reflect on the scientific validity of the work.
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Medullary Thymic Epithelial Cell Antigen-presentation Assays
AB Alexia Borelli §
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MI Magali Irla
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Published: Vol 13, Iss 21, Nov 5, 2023
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The authors used this protocol in eLIFE Feb 2022
Abstract
Medullary thymic epithelial cells (mTEC) are bona fide antigen-presenting cells that play a crucial role in the induction of T-cell tolerance. By their unique ability to express a broad range of tissue-restricted self-antigens, mTEC control the clonal deletion (also known as negative selection) of potentially hazardous autoreactive T cells and the generation of Foxp3+ regulatory T cells. Here, we describe a protocol to assess major histocompatibility complex (MHC) class II antigen-presentation capacity of mTEC to CD4+ T cells. We detail the different steps of thymus enzymatic digestion, immunostaining, cell sorting of mTEC and CD4+ T cells, peptide-loading of mTEC, and the co-culture between these two cell types. Finally, we describe the flow cytometry protocol and the subsequent analysis to assess the activation of CD4+ T cells. This rapid co-culture assay enables the evaluation of the ability of mTEC to present antigens to CD4+ T cells in an antigen-specific context.
Key features
• This protocol builds upon the method used by Lopes et al. (2018 and 2022) and Charaix et al. (2022).
• This protocol requires transgenic mice, such as OTIIxRag2-/- mice and the cognate peptide OVA323–339, to assess mTEC antigen presentation to CD4+ T cells.
• This requires specific equipment such as a Miltenyi Biotec AutoMACS® Pro Separator, a BD FACSAriaTM III cell sorter, and a BD® LSR II flow cytometer.
Graphical overview
Keywords: Medullary thymic epithelial cells CD4+ T cells Antigen-presentation assay Cell purification Co-culture
Background
The thymus is a primary lymphoid organ that ensures the production of functional and self-tolerant naïve T cells. The development of those cells depends on stromal niches composed of thymic epithelial cells (TEC) that provide essential cues for the survival, proliferation, differentiation, and migration of developing T cells, called thymocytes (Irla, 2022). According to their localization in the thymus, TEC are subdivided into two main subsets: cortical (cTEC) and medullary TEC (mTEC). cTEC support early stages of thymopoiesis, including T-cell progenitor homing, T-cell lineage commitment, proliferation and survival of immature thymocytes, and positive selection of CD4+ and CD8+ single positive thymocytes. In contrast, mTEC control late stages of T-cell development, i.e., the clonal deletion of highly autoreactive thymocytes and the generation of mature Foxp3+ regulatory T cells (Treg). mTEC are commonly subdivided into two main subsets based on the expression of MHC class II (MHC-II) and CD80 molecules: mTEClo (MHC-IIloCD80lo) and mTEChi (MHC-IIhiCD80hi), with mTEClo containing progenitors capable of differentiating into mTEChi (Gray et al., 2006; Gäbler et al., 2007; Miragaia et al., 2018). mTEC constitute a crucial antigen reservoir due to their unique capacity to express a wide range of tissue-restricted self-antigens. The expression of the majority of these tissue-restricted self-antigens is mediated by the transcription factors Aire (Autoimmune regulator) and Fezf2 (Fez family zinc finger 2), which regulate the expression of 3,000–4,000 and 600–700 genes, respectively (Sansom et al., 2014; Takaba et al., 2015). mTEC are bona fide antigen-presenting cells, as they can efficiently induce the clonal deletion of highly autoreactive CD4+ and CD8+ T cells, as well as the differentiation of CD4+ T cells towards the Foxp3+ Treg cell lineage (Aschenbrenner et al., 2007; Hinterberger et al., 2010; Aichinger et al., 2013). Interestingly, our lab recently showed that Aire+ mTEC also have the ability to restimulate recirculating Foxp3+ Treg upon their re-entry in the thymus in an antigen-specific manner (Charaix et al., 2022). It is important to notice that mTEC–CD4+ T cell interactions act as bidirectional signals, as not only mTEC are crucial for CD4+ T cell selection but also CD4+ T cells control mTEC differentiation (Irla et al., 2008; Lopes et al., 2015 and 2022).
The antigen-presentation capacity of mTEC to CD4+ T cells can be assessed by co-culture experiments using variable amounts of mTEC or peptide concentration (Aschenbrenner et al., 2007; Wirnsberger et al., 2009; Hinterberger et al., 2010; Lopes et al., 2018 and 2022; Charaix et al., 2022). This in vitro co-culture protocol describes the steps to analyze the antigen-presentation capacity of mTEC through the activation of CD4+ T cells in an antigen-specific manner. It describes the different steps of thymus collection, enzymatic digestion, immunostaining, cell sorting of mTEC and CD4+ T cells, peptide-loading of mTEC, and the co-culture between these two cell types. Although this approach may be limited by the fact that it is an in vitro assay, it has the advantage of specifically and rapidly evaluating, within two days, the antigen-presentation capacity of mTEC. This protocol can also be used to assess the antigen-presentation capacity of human mTEC or dendritic cell subsets, the latter also implicated in the induction of T-cell tolerance (Lopes et al., 2015; Irla, 2022).
Materials and reagents
Biological species
C57BL/6 WT mice at the age of 4–6 weeks (Charles River)
OTIIxRag2-/- mice on a C57BL/6 background at the age of 4–6 weeks (Shinkai, 1992; Barnden et al., 1998)
Note: Mice are age- and sex-matched.
Reagents
Fetal bovine serum (Pan Biotech, catalog number: P30-3306)
Bovine serum albumin (Sequens IVD, catalog number: 1000-70)
LiberaseTM (Roche, catalog number: 5401127001)
DNase I (Roche, catalog number: 10104159001)
HBSS, without calcium and magnesium, no phenol red (Thermo Fisher Scientific, GibcoTM, catalog number: 14175-053)
PBS 1×, without calcium and magnesium (Thermo Fisher Scientific, GibcoTM, catalog number: 10010-023)
EDTA, 0.5 M pH 8.0 (Thermo Fisher Scientific, Invitrogen, catalog number: AM9261)
L-glutamine (Thermo Fisher Scientific, GibcoTM, catalog number: 25030024)
Penicillin and streptomycin (Thermo Fisher Scientific, GibcoTM, catalog number: 15140-122)
2-mercaptoethanol (Thermo Fisher Scientific, GibcoTM, catalog number: 31350-010)
Sodium pyruvate (Thermo Fisher Scientific, GibcoTM, catalog number: 11360-039)
D-MEM (Thermo Fisher Scientific, GibcoTM, catalog number: 41965-039)
Red blood cell (RBC) lysis buffer (Thermo Fisher Scientific, catalog number: 00-4333-57)
Antibodies:
Purified Rat Anti-Mouse CD16/CD32 (Mouse BD Fc BlockTM) (BD Biosciences, clone 2.4G2, catalog number: 553142)
Biotin anti-mouse CD45 antibody (BioLegend, clone 30-F11, catalog number: 103104)
Biotin anti-mouse CD8 antibody (BioLegend, clone 53-6.7, catalog number: 100704)
Biotin anti-mouse CD11c antibody (BioLegend, clone N418, catalog number: 117304)
PE/Cyanine7 conjugated anti-mouse EpCAM (CD326) antibody (BioLegend, clone G8.8, catalog number: 118216)
PE conjugated anti-mouse Ly51 (CD249) antibody (BD Biosciences, clone BP-1, catalog number: 553735)
Fluorescein conjugated UEA1 (Vector Laboratories, catalog number: FL-1061-5)
Brilliant Violent 421TM conjugated anti-mouse CD80 antibody (BioLegend, clone 16-10A1, catalog number: 104726)
Alexa Fluor® 700 conjugated anti-mouse MHC-II (I-A/I-E) antibody (BioLegend, clone M5/114.15.2, catalog number: 107622)
Brilliant Violent 421TM conjugated anti-mouse CD4 antibody (BD Biosciences, clone RM4.5, catalog number: 740007)
PerCP-CyTM 5.5 conjugated anti-mouse CD8 antibody (BD Biosciences, clone 53-6.7, catalog number: 551162)
PE conjugated anti-mouse CCR6 antibody (BioLegend, clone 29-2L17, catalog number: 129804)
PE/Cyanine7 conjugated anti-mouse CD25 antibody (BioLegend, clone PC61, catalog number: 102016)
APC conjugated anti-mouse CD69 antibody (BioLegend, clone H1.2F3, catalog number: 104514)
Anti-Biotin MicroBeads (Miltenyi Biotec, catalog number: 130-090-485)
Ovalbumine323–339 (OVA323–339) peptide (Polypeptide group SC1303 then Anaspec Inc., catalog number: AS-27024)
Solutions
Digestion buffer (see Recipes)
FACS buffer (see Recipes)
AutoMACS buffer (see Recipes)
Complete culture medium (see Recipes)
Note: All buffers and culture medium are filtered through a 0.22 μm filter.
Recipes
Digestion buffer
Reagent Final concentration Quantity
HBSS 1× n/a 9.9 mL
LiberaseTM 50 μg/mL 100 μL
DNase I 100 μg/mL 10 μL
Total n/a 10 mL
Note: Enzymes are sensitive to temperature; therefore, prepare before use.
FACS buffer
Reagent Final concentration Quantity
PBS 1× n/a 1,980 mL
EDTA (0.5 M, pH 8.0) 5 mM 20 mL
Bovine serum albumin 0.5% 10 g
Total n/a 2,000 mL
Note: Store up to one month at 4 °C and keep on ice for the duration of the protocol.
AutoMACS buffer
Reagent Final concentration Quantity
PBS 1× n/a 1,982 mL
EDTA (0.5 M, pH 8.0) 2 mM 8 mL
Fetal bovine serum 0.5% 10 mL
Total n/a 2,000 mL
Note: Store up to one month at 4 °C and keep on ice for the duration of the protocol.
Complete culture medium
Reagent Final concentration Quantity
D-MEM n/a 435 mL
Fetal bovine serum 10% 50 mL
L-Glutamine
Sodium pyruvate
2-mercaptoethanol
Penicillin and streptomycin
2 mM
1 mM
2 × 10-5 M
100 IU/mL
5 mL
5 mL
500 μL
5 mL
Total n/a 500 mL
Note: Store up to one month at 4 °C. One hour before use, place the medium in a water bath at 37 °C.
Laboratory supplies
Curved forceps (Fine Science Tools, catalog number: 11271-30)
Straight forceps (Fine Science Tools, catalog number: 11064-07)
Dissection scissors (BochemTM, catalog number: 4070)
Micropipette tips [Sarstedt, catalog number: 70.3050.205 (1,000 μL); Starlab, catalog numbers: S1111-1810 (200 μL) and S1110-3700 (0.1–10 μL)]
5 mL Falcon round-bottom polystyrene tube (Fisher Scientific, Corning, catalog number: 352008)
15 mL tube (Sarstedt, catalog number: 62.554.502)
50 mL tube (Sarstedt, catalog number: 62.547.254)
1.5 mL tubes (Eppendorf, Dutscher, catalog number: 033305)
96-well U-bottom cell culture plate (Cellstar®, Greiner Bio-One, catalog number: 650180)
96-well V-bottom cell culture plate (Cellstar®, Greiner Bio-One, catalog number: 651180)
70 μm pore cell strainers (Sarstedt, catalog number: 83.3945.070)
30 μm pre-separation cell strainers (Miltenyi Biotec, catalog number: 130-041-407)
1 mL syringe (Terumo; catalog number: SS+01H1)
Steritop E-GP sterile vacuum 0.22 μm filtration system (Millipore Merck, catalog number: SEGPT0045)
Equipment
Micropipettes 0.1–2.5 μL, 2–20 μL, 20–200 μL, and 100–1,000 μL (Eppendorf, Thermo Fisher Scientific, catalog number: 05-403-152)
Centrifuge (Eppendorf, Thermo Fisher Scientific, model: 5810R and 5415R)
37 °C 5% CO2 incubator Forma Scientific 3548 (Labexchange, Forma Scientific, catalog number: B00032422)
4 °C refrigerator (Candy)
-20 °C freezer (Liebherr)
Water bath (Grant Instruments)
AutoMACS® Pro Separator (Miltenyi Biotec, catalog number: 130-092-545)
BD FACSAriaTM III cell sorter (BD Biosciences)
BD® LSR II flow cytometer (BD Biosciences)
Software and datasets
FlowJo (BD Biosciences, Version 10.8.1)
BD FACSDivaTM software (BD Biosciences, Version 9.0)
Procedure
Thymus withdrawal
Mouse euthanasia was performed through 100% CO2 inhalation at a flow rate of 20 L/min in accordance with National and European laws for laboratory animal welfare (EEC Council Directive 2010/63/UE) and the Marseille Ethical Committee for Animal experimentation no. 14. Verify that the mouse is unresponsive to a toe pinch.
Place the mouse on its back and pin each limp down to the dissection board. Spray with 70% ethanol to sterilize the mouse body.
Open the thoracic cavity with scissors and carefully expose the heart and thymus above without cutting major blood vessels.
Gently remove the thymus with a pair of forceps.
Note: Make sure to remove fat and connective tissue from the thymus.
Transfer the thymus into a 15 mL collection tube pre-filled with 2 mL of enzymatic digestion buffer for WT mice or 2 mL of cold FACS buffer for OTIIxRag2-/- mice.
Thymic epithelial cell isolation
Digest the thymus from WT mice in a water bath at 37 °C for 15 min in 2 mL of digestion buffer; then, dissociate the tissue by approximately 20 recurrent aspirations through a 1,000 μL tip.
Note: Agitate the fractions gently during the digestion process and avoid bubbles.
Filter thymic cells through a 70 μm pore cell strainer into a 50 mL tube with 3 mL of cold FACS buffer.
Collect remnant undigested tissue from the cell strainer with forceps and return it to the 15 mL tube containing digestion buffer.
Digest remnant tissue in a water bath at 37 °C for 15 min. Then, dissociate the tissue by approximately 20 recurrent aspirations through a 1,000 μL tip.
Filter thymic cells through a 70 μm pore cell strainer into the 50 mL tube with 3 mL of cold FACS buffer.
Repeat steps B3–B5 until complete tissue digestion.
Centrifuge at 450× g for 5 min at 4 °C.
Remove the supernatant and resuspend the cell pellet in 1 mL of RBC lysis buffer for 3 min.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Prepare the biotin antibody mix No. 1: dilute the biotin anti-mouse CD45 antibody in cold FACS buffer at the final concentration of 2 μg/mL.
After the centrifugation, remove the supernatant and resuspend the cell pellet with 500 μL of the biotin antibody mix No. 1.
Incubate the cells for 15 min at 4 °C.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Prepare the anti-biotin microbead mix: dilute 50 μL of anti-biotin microbeads in 450 μL of cold FACS buffer.
After the centrifugation, remove the supernatant and resuspend the cell pellet with 500 μL of the anti-biotin microbead mix.
Incubate the cells for 15 min at 4 °C.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Remove the supernatant and resuspend the cell pellet in 4 mL of cold AutoMACS buffer.
Pass the cells through a pre-separation filter into a 15 mL tube to remove cell clumps that may clog the AutoMACS® Pro Separator columns.
Proceed to magnetic separation with the AutoMACS® Pro Separator using the DepleteS program.
Keep the CD45- fraction and discard the CD45+ fraction.
Centrifuge the CD45- fraction at 450× g for 5 min at 4 °C.
Prepare the antibody mix in cold FACS buffer for mTEC staining with the following concentrations:
Purified Rat Anti-Mouse CD16/CD32: final concentration 1 μg/million cells.
PE/Cyanine7 conjugated anti-mouse EpCAM antibody: final concentration 60 ng/mL.
PE conjugated anti-mouse Ly51 antibody: final concentration 60 ng/mL.
Fluorescein conjugated UEA1: final concentration 6.25 μg/mL.
Note: Based on the desired mTEC subset for the co-culture assay, include anti-mouse CD80 and anti-mouse I-A/I-E antibodies to discriminate mTEClo and mTEChi with:
Brilliant Violent 421TM conjugated anti-mouse CD80 antibody: final concentration 0.5 μg/mL.
Alexa Fluor® 700 conjugated anti-mouse I-A/I-E antibody: final concentration 0.9 μg/mL.
Remove the supernatant and resuspend the cell pellet with 500 μL of antibody mix for mTEC staining.
Incubate the cells for 15 min at 4 °C.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Remove the supernatant and resuspend the stained cell pellet with 500 μL of cold FACS buffer.
Keep on ice until cell sorting.
Sort total mTEC (or alternatively mTEClo and mTEChi) with a BD FACSAriaTM III cell sorter (Figure 1). Cells are collected in 1.5 mL Eppendorf tubes containing 100 μL of complete culture medium.
Figure 1. Gating strategy used to purify medullary thymic epithelial cells (mTEC) from WT mice by cell sorter after AutoMACS pre-enrichment of CD45- cells. EpCAM+ TEC were divided into mTEC (UEA-1+Ly51lo cells) and subsequently into mTEClo (UEA-1+Ly51loCD80-I-A/I-E- cells) and mTEChi (UEA-1+Ly51loCD80+ I-A/I-E+ cells) based on the level of the CD80 and MHC-II molecules.
Isolation of OTII CD4+ T cells
Mechanically dissociate the thymus from OTIIxRag2-/- mice by scratching it in cold FACS buffer on a 70 μm pore cell strainer fixed on a 50 mL tube with the plunger of a 1 mL syringe.
Centrifuge at 450× g for 5 min at 4 °C.
Remove the supernatant and resuspend the cell pellet in 1 mL of RBC lysis buffer for 3 min.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Prepare the biotin antibody mix No. 2: Dilute the biotin anti-mouse CD8 and CD11c antibodies in cold FACS buffer at the final concentration of 2.5 μg/mL and 1.5 μg/mL, respectively.
After the centrifugation, remove the supernatant and resuspend the cell pellet with 500 μL of the biotin antibody mix No. 2.
Incubate the cells for 15 min at 4 °C.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Prepare the anti-biotin microbead mix: dilute 50 μL of anti-biotin microbeads in 450 μL of cold FACS buffer.
After the centrifugation, remove the supernatant and resuspend the cell pellet with 500 μL of the anti-biotin microbead mix.
Incubate the cells for 15 min at 4 °C.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Remove the supernatant and resuspend the cell pellet in 4 mL of cold AutoMACS buffer.
Pass the cells through a pre-separation filter into a 15 mL tube to remove cell clumps that may clog the AutoMACS® Pro Separator columns.
Proceed to magnetic separation with the AutoMACS® Pro Separator, using the Deplete program.
Keep the CD8- and CD11c- fraction and discard the other fraction.
Centrifuge the CD8- and CD11c- fraction at 450× g for 5 min at 4 °C.
Prepare the antibody mix in cold FACS buffer for CD4+ T cells staining with the following concentrations:
Brilliant Violent 421TM conjugated anti-mouse CD4 antibody: final concentration 1 μg/mL.
PerCP-CyTM 5.5 conjugated anti-mouse CD8 antibody: final concentration 1 μg/mL.
PE conjugated anti-mouse CCR6 antibody: final concentration 0.7 μg/mL.
PE/Cyanine7 conjugated anti-mouse CD25 antibody: final concentration 0.5 μg/mL.
APC conjugated anti-mouse CD69 antibody: final concentration 1 μg/mL.
Remove the supernatant and resuspend the cell pellet with 500 μL of antibody mix for CD4+ T-cells staining.
Incubate the cells for 15 min at 4 °C.
Wash the cells by adding 10 mL of cold FACS buffer.
Centrifuge at 450× g for 5 min at 4 °C.
Remove the supernatant and resuspend the stained cell pellet with 500 μL of cold FACS buffer.
Keep on ice until cell sorting.
Sort mature OTII CD4+ T cells with a BD FACSAriaTM III cell sorter (Figure 2). Cells are collected in 1.5 mL Eppendorf tubes containing 100 μL of complete culture medium.
Figure 2. Gating strategy used to purify CD4+ T cells from OTIIxRag2-/- mice by cell sorter after AutoMACS pre-enrichment of CD8- and CD11c- cells. Recirculating cells from the periphery into the thymus were excluded based on CCR6 expression. Mature OTII CD4+ T cells were gated as CD4+CD8-CCR6-CD25-CD69-.
In vitro co-culture assays
After the purification of mTEC, centrifuge the Eppendorf collection tubes at 450× g for 5 min at 4 °C.
Remove the supernatant and carefully resuspend the cell pellet in culture medium supplemented with 5 μg/mL of OVA323–339 peptide.
Note: OVA323–339concentration can be adjusted as required.
Distribute mTEC in a plate with U bottom (100 μL/well) according to the cell concentration needed for the experimental plan.
Incubate the cells in a 37 °C, 5% CO2 air humidified incubator for 1 h.
After the purification of CD4+ T cells, centrifuge the Eppendorf collection tubes at 450× g for 5 min at 4 °C.
Remove the supernatant and gently resuspend the cell pellet in complete culture medium.
Add 100 μL of complete culture medium containing 105 CD4+ T cells in each well containing OVA323–339-loaded mTEC.
Incubate the cells in a 37 °C, 5% CO2 air humidified incubator for 18 h.
Fluorescence-activated cell sorting (FACS) analysis of OTII CD4+ T-cell activation after co-culture
After 18 h of co-culture, transfer the cells to a 96-well V-bottom plate for staining.
Centrifuge at 450× g for 5 min at 4 °C.
Prepare the antibody mix in cold FACS buffer for the analysis of OTII CD4+ T cell activation with the following concentrations:
Brilliant Violent 421TM conjugated anti-mouse CD4 antibody: final concentration 1 μg/mL.
APC conjugated anti-mouse CD69 antibody: final concentration 1 μg/mL.
Remove the supernatant and resuspend the cell pellet with 50 μL of antibody mix.
Incubate the cells for 15 min at 4 °C.
Centrifuge at 450× g for 5 min at 4 °C. Then, remove the supernatant and add 150 μL of cold FACS buffer.
Repeat step E6.
Resuspend the cells in 150 μL of cold FACS buffer.
Transfer the cells to a 5 mL tube.
Acquire the samples on the BD® LSR II flow cytometer.
Data analysis
FACS data were acquired on a BD® LSR II flow cytometer using violet laser 405 nm, blue laser 488 nm, green laser 561 nm, and red laser 633 nm. FACS data were then analyzed using FlowJo software (version 10.8.1). The gating strategy used to identify the proportion of activated CD69+ OTII T cells after co-culture with OVA323–339-loaded mTEC is shown in Figure 3A. This protocol can be used with different ratios of mTEC to OTII CD4+ T cells to assess the antigen-presentation capacity of mTEC. As depicted in Figure 3B, the activation of OTII CD4+ T cells, reflected by the upregulation of CD69, gradually increases with the numbers of OVA323–339-loaded mTEC. The details regarding the analysis can be found in the original article, e.g., Figure 2A (Lopes et al., 2022).
Figure 3. Representative results of OTII CD4+ T-cell activation after co-culture with OVA323–339-loaded medullary thymic epithelial cells (mTEC). (A) Gating strategy used to analyze the frequency of activated CD69+ OTII CD4+ T cells by flow cytometry. (B) Percentage of CD69+ OTII CD4+ T cells cultured with variable numbers of OVA323–339-loaded WT mTEC for 18 h.
Validation of protocol
This protocol has been adapted from the previously published paper from our team (Lopes et al., 2022).
General notes and troubleshooting
General notes
An enzymatic digestion to isolate OTII CD4+ T cells should be avoided since it may affect their viability and/or activation.
This protocol can be used in a polyclonal context.
This protocol can be applied to assess MHC-I antigen-presentation capacity to CD8+ T cells with transgenic mouse models, such as OTI mice and the cognate peptide OVA257–264 (SIINFEKL).
A viability marker can be added in steps B26, C21, and E3.
This protocol can be applied for a range of downstream applications, such as FACS analysis (Charaix et al., 2022; Lopes et al., 2022) or gene expression profiling (Lopes et al., 2018).
Each step described in the section “CD4+ T cell isolation” can be applied for the isolation of splenic CD4+ T cells and Foxp3+ Treg. This procedure was used in the publication of Charaix et al. (2022).
This protocol can be used to test the capacity of mTEC to generate or activate Foxp3+ Treg (Charaix et al., 2022).
Troubleshooting
The time of digestion must be optimized to avoid over-digestion, which can lead to cell death and epitope removal.
To ensure an optimal pre-enrichment with the AutoMACS® Pro Separator, carefully follow the manufacturer’s recommendations concerning cell concentration.
Clogging of the AutoMACS® Pro Separator: depending on the cell density, cell suspension should be diluted and separated into two different tubes.
During FACS analysis: make sure to use a cytometer compatible with the fluorochromes used in the experiment, use fluorochrome combinations to avoid excessive spectral overlap, and verify compensation parameters.
To avoid cell death, make sure to work quickly through the protocol.
Acknowledgments
We thank the CIML animal housing and flow cytometry core facilities. This work was supported by the Marie Skłodowska-Curie Actions (Career Integration Grants, CIG_SIGnEPI4Tol_618541), a CoPoC-proof of concept (MAT-PI-17326-A-01 to M.I.), a pre-maturation grant from A*MIDEX, a French “Investissements d'avenir” program (LTalpha-Treg to M.I.) and Agence Nationale de la Recherche (grant SelfExpress ANR-22-CE15-0045 and grant Reality ANR-22-CE18-0045-01 to M.I.). This work was supported by institutional grants from INSERM, CNRS and Aix-Marseille Université. AB and CZ were supported by a PhD fellowship from the Ministère de l’Enseignement Supérieur et de la Recherche et de l’Innovation (MESRI) and by the Fondation ARC pour la Recherche sur le Cancer (to AB, ARCDOC42022010004424). This protocol has been adapted from the previously published paper in Elife by our team (Lopes et al., 2022). The graphical abstract was generated using biorender.com.
Competing interests
Authors declare no competing interests.
Ethical considerations
All experiments were done in accordance with national and European laws for laboratory animal welfare (EEC Council Directive 2010/63/UE) and were approved by the Marseille Ethical Committee for Animal Experimentation (Comité National de Réflexion Ethique sur l’Expérimentation Animale no. 14).
References
Aichinger, M., Wu, C., Nedjic, J. and Klein, L. (2013). Macroautophagy substrates are loaded onto MHC class II of medullary thymic epithelial cells for central tolerance. J. Exp. Med. 210(2): 287–300.
Aschenbrenner, K., D’Cruz, L. M., Vollmann, E. H., Hinterberger, M., Emmerich, J., Swee, L. K., Rolink, A. and Klein, L. (2007). Selection of Foxp3+ regulatory T cells specific for self antigen expressed and presented by Aire+ medullary thymic epithelial cells. Nat. Immunol. 8(4): 351–358.
Barnden, M. J., Allison, J., Heath, W. R. and Carbone, F. R. (1998). Defective TCR expression in transgenic mice constructed using cDNA-based α- and β-chain genes under the control of heterologous regulatory elements. Immunol. Cell Biol. 76(1): 34–40.
Charaix, J., Borelli, A., Santamaria, J. C., Chasson, L., Giraud, M., Sergé, A. and Irla, M. (2022). Recirculating Foxp3+ regulatory T cells are restimulated in the thymus under Aire control. Cell. Mol. Life Sci. 79(7): e1007/s00018-022-04328-9.
Gäbler, J., Arnold, J. and Kyewski, B. (2007). Promiscuous gene expression and the developmental dynamics of medullary thymic epithelial cells. Eur. J. Immunol. 37(12): 3363–3372.
Gray, D. H. D., Seach, N., Ueno, T., Milton, M. K., Liston, A., Lew, A. M., Goodnow, C. C. and Boyd, R. L. (2006). Developmental kinetics, turnover, and stimulatory capacity of thymic epithelial cells. Blood 108(12): 3777–3785.
Hinterberger, M., Aichinger, M., Prazeres da Costa, O., Voehringer, D., Hoffmann, R. and Klein, L. (2010). Autonomous role of medullary thymic epithelial cells in central CD4+ T cell tolerance. Nat. Immunol. 11(6): 512–519.
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Irla, M., Hugues, S., Gill, J., Nitta, T., Hikosaka, Y., Williams, I. R., Hubert, F. X., Scott, H. S., Takahama, Y., Holländer, G. A., et al. (2008). Autoantigen-Specific Interactions with CD4+ Thymocytes Control Mature Medullary Thymic Epithelial Cell Cellularity. Immunity 29(3): 451–463.
Lopes, N., Boucherit, N., Santamaria, J. C., Provin, N., Charaix, J., Ferrier, P., Giraud, M. and Irla, M. (2022). Thymocytes trigger self-antigen-controlling pathways in immature medullary thymic epithelial stages. eLife 11: e69982.
Lopes, N., Charaix, J., Cédile, O., Sergé, A. and Irla, M. (2018). Lymphotoxin α fine-tunes T cell clonal deletion by regulating thymic entry of antigen-presenting cells. Nat. Commun. 9(1): e1038/s41467-018-03619-9.
Lopes, N., Sergé, A., Ferrier, P. and Irla, M. (2015). Thymic Crosstalk Coordinates Medulla Organization and T-Cell Tolerance Induction. Front. Immunol. 6: e00365.
Miragaia, R. J., Zhang, X., Gomes, T., Svensson, V., Ilicic, T., Henriksson, J., Kar, G. and Lönnberg, T. (2018). Single-cell RNA-sequencing resolves self-antigen expression during mTEC development. Sci. Rep. 8(1): e1038/s41598-017-19100-4.
Sansom, S. N., Shikama-Dorn, N., Zhanybekova, S., Nusspaumer, G., Macaulay, I. C., Deadman, M. E., Heger, A., Ponting, C. P. and Holländer, G. A. (2014). Population and single-cell genomics reveal the Aire dependency, relief from Polycomb silencing, and distribution of self-antigen expression in thymic epithelia. Genome Res. 24(12): 1918–1931.
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A Guideline for Assessment and Characterization of Bacterial Biofilm Formation in the Presence of Inhibitory Compounds
BE Bassam A. Elgamoudi
VK Victoria Korolik
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4866 Views: 878
Reviewed by: Emilia KrypotouElizabeth LibbyEsteban Paredes-Osses
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Original Research Article:
The authors used this protocol in Antibiotics Nov 2020
Abstract
Campylobacter jejuni, a zoonotic foodborne pathogen, is the worldwide leading cause of acute human bacterial gastroenteritis. Biofilms are a significant reservoir for survival and transmission of this pathogen, contributing to its overall antimicrobial resistance. Natural compounds such as essential oils, phytochemicals, polyphenolic extracts, and D-amino acids have been shown to have the potential to control biofilms formed by bacteria, including Campylobacter spp. This work presents a proposed guideline for assessing and characterizing bacterial biofilm formation in the presence of naturally occurring inhibitory molecules using C. jejuni as a model. The following protocols describe: i) biofilm formation inhibition assay, designed to assess the ability of naturally occurring molecules to inhibit the formation of biofilms; ii) biofilm dispersal assay, to assess the ability of naturally occurring inhibitory molecules to eradicate established biofilms; iii) confocal laser scanning microscopy (CLSM), to evaluate bacterial viability in biofilms after treatment with naturally occurring inhibitory molecules and to study the structured appearance (or architecture) of biofilm before and after treatment.
Keywords: Biofilm assay Biofilm method Microtiter plate assay Antibiofilm compounds Natural compounds D-amino acids Biofilm of Campylobacter
Background
Biofilm formation is considered to be important for the survival and transmission of bacterial pathogens to humans where they are able to cause disease [1]. Inhibition of bacterial biofilms by naturally occurring compounds such as polyphenols, essential oils, and D-amino acids [i.e., serine (D-Ser)] has been previously investigated [2–6] using several methods, such as spectrophotometric or fluorescence-based methods, to quantify total formed biofilms [7–9]. The most common system used to assess biofilm formation is a spectrophotometer, as it does not require overly specialized or expensive equipment, a great quantity of analytical reagents, or a high level of user expertise. This protocol presents a proposed guideline for consistent and reproducible analysis of biofilm formation in the presence of naturally occurring inhibitory compounds using Campylobacter jejuni as a model organism.
C. jejuni presents itself as a suitable model for this method as it is an opportunistic pathogen widely believed to be responsible for most cases of bacterial gastroenteritis worldwide. C. jejuni is a common commensal bacterium of poultry, especially chickens [10, 11], and has been reported to be able to form mono-species biofilms and to integrate into composite biofilms with other bacterial species, such asPseudomonas aeruginosa[8, 12–18]. The ability of Campylobacter to form biofilms plays a critical role in its survival in the environment as well as in the dissemination of infection and the emergence of antibiotic resistance [19–23].
Materials and reagents
Materials
Media: Mueller-Hinton agar/broth (MHA/MHB) (Thermo Fisher Scientific, catalog number: CM0337) and Luria-Bertani broth (LB) (Oxoid, catalog number: CM0996B)
Antibiotic stock: Trimethoprim (2.5 μg/mL) (Sigma-Aldrich, catalog number: T7883-5G) and Vancomycin (10 μg/mL (Sigma-Aldrich, catalog number: 1709007)
Strains: Campylobacter jejuni NCTC 11168-O [24] and Pseudomonas aeruginosa PA0-1
Culture conditions: cultures ofC. jejuniare grown microaerobically (85% N2, 10% CO2, and 5% O2) at 42 °C for 36 h while Pseudomonas aeruginosa is grown aerobically at 37 °C for 24 h
Chemicals: natural compounds that have high anti-biofilm properties such as D-serine (D-Ser) (Sigma-Aldrich, catalog number: S4250)
Phosphate-buffered saline (PBS, pH 7.4) (Sigma-Aldrich, catalog number: P4417)
Multi-channel pipette (e.g., 100–200 μL, 300–1,000 μL) (Thermo Scientific, catalog number: 4661180N)
15 mL conical tubes or glass test tubes for growing liquid cultures (Thermo Scientific, catalog number: 339650)
24- or 96-well clear flat-bottom plates (Geiner Bio-One, catalog number: 655101)
Tray or box slightly larger than a 96-well plate (e.g., tray dimension ~200 mm × 100 mm)
0.1% Crystal violet solution (0.1 g of Crystal violet in 100 mL demineralized water) (Sigma-Aldrich, catalog number: 548-62-9)
Modified biofilm dissolving solution (MBDS) [sodium dodecyl sulfate (SDS) (Sigma-Aldrich, catalog number: 436143) dissolved to a final concentration of 10% with 80% Ethanol in H2O) [25]
Paper towels
Large beaker
For confocal laser scanning microscopy (CLSM):
6-well polystyrene clear flat-bottom plates (Geiner Bio-One, catalog number: 655101)
Glass coverslips (e.g., 22 mm2 coverslips)
Glass slides
5% formaldehyde solution (in distilled H2O)
4′,6-diamidino-2-phenylindole (DAPI) (Sigma-Aldrich, catalog number: D8417)
Mounting medium (Ibidi GmbH, catalog number: 50001)
Nail varnish
CLSM (Nikon Microscopy, model: Nikon A1R+)
Equipment
Plate reader (Tecan, model: Infinite M200 Pro) for measuring optical density (OD) range at 570–600 nm
Incubator (SHEL LAB, model: SM12)
Laminar flow cabinet (ESCO, model: AHL-4A2)
Software
ImageJ analysis software (National Institutes of Health, Bethesda, MD, USA)
Procedure
Protocol 1: Biofilm formation inhibition assay (Figure 1)
Figure 1. Protocol 1 for screening biofilm formation inhibition. (1–2) Prepare the bacterial suspension and dispense it into a 24-well microplate. (3) Dispense different concentrations of the tested compound (D-Ser) in desired increments into the microplate. (4) Incubate the microplate to generate the biofilm inside the wells. (5) After incubation, the wells are washed, allowed to dry, and then stained with crystal violet solution (6). The stained biofilms are washed to remove excess dye and a volume of MBDS solution is added to the wells to dissolve the crystal violet stained biofilm. (7) The absorbance (OD) of the de-stained MBDS solution is read on a spectrophotometer.
Single-species assay
Note: To investigate the effect of a compound to be tested on biofilm-formed C. jejuni.
Recover C. jejuni NCTC 11168-O from -80 °C storage, plate the stock on MHA media supplemented with appropriate antibiotics [Trimethoprim (2.5 μg/mL) and Vancomycin (10 μg/mL)], and incubate microaerobically (85% N2, 10% CO2, and 5% O2) at 42 °C overnight.
Harvest bacterial cells from the agar plates into 1 mL of MHB using a glass or a disposable plastic rod to lift the cells off the agar plates into a test tube, and transfer ~100–200 μL of the resulting bacterial suspension into 15 mL of MHB (supplemented with appropriate antibiotics). Incubate microaerobically at 42 °C and 125 rpm in a shaker incubator (18–20 h overnight).
Dilute overnight culture in fresh MHB (supplemented with appropriate antibiotics) to achieve cell density of OD600 of 0.05 at the start of the logarithmic growth phase (~107 CFU/mL).
Dispense 2 mL of diluted bacterial suspension into each well of a 24-well plate (minimum of three rows of four wells to each sample) or 180 μL per well for 96-well plates. Uninoculated MHB is used as a negative control.
To test the inhibitory effect of the tested compound (e.g., D-Ser), add the chosen concentrations (i.e., 1–50 mM) of the compounds to be tested directly to the culture in the wells. For example, 1 M of D-Ser (in PBS) was used as stock solution and 100 μL of stock solution was added to each well, except for negative control.
Cover the 24-well plates and incubate them at 42 °C under microaerophilic conditions without shaking (static culture) for 24 h (see Notes 1 and 2). Each plate must include medium-only control in four wells.
To quantify biofilm formation, go to section “Assessment of biofilm formation” from Protocol 2.
Dual/multiple species assay
Note: To investigate the effect of a compound to be tested on biofilms formed by a mixed culture (in this example: C. jejuni and P. aeruginosa).
Grow P. aeruginosa cells aerobically in MHB overnight at 37 °C and then adjust the cell density with fresh MHB to OD600 of 0.1, at the start of the logarithmic growth phase (107 CFU/mL) [26].
Grow C. jejuni cells as described above (steps A1–A3).
Use a ratio of 1:1 of the cell suspensions for both/multiple bacteria for the assay.
Repeat steps A4–A6 as described above (see Notes 1 and 2).
To quantify biofilm formation, go to section “Assessment of biofilm formation” from Protocol 2.
Protocol 2: Biofilm dispersal assay (Figure 2)
Figure 2. Protocol 2 for screening biofilm formation dispersal. (1–2) Prepare the bacterial suspension and dispense it into a 24-well microplate. (3) Incubate the microplate to generate the biofilm inside the wells. (4) The bacterial culture is then removed, and the wells are washed to remove planktonic cells before dispensing different concentrations of the tested compound (D-Ser) in desired increments into the microtiter plate. (5) After incubation, the wells are washed, allowed to dry, and then stained with crystal violet solution. (6) The stained biofilms are washed to remove excess dye and MBDS solution is added to the wells to dissolve the crystal violet stained biofilm. (7) The absorbance (OD) of the de-stained MBDS solution is read on a spectrophotometer.
Repeat steps A1–A6, except step 5.
Remove the media from the 24-well plate and add 500 μL of PBS with an appropriate concentration (i.e., 10–50 mM) of the compound to be tested in each well (PBS-only is used as negative control).
Incubate the plates at 42 °C under microaerobic conditions without shaking for 24 h (see Notes 1 and 2).
Take the supernatants and measure the OD600 for each well.
To quantify biofilm formation, go to section “Assessment of biofilm formation.”
Assessment of biofilm formation
Remove the media from plates (by inverting over an absorbent paper towel in a tray) and rinse gently with distilled water twice to remove planktonic cells.
Dry plates by gently tapping on a paper towel until no liquid remains in the wells.
Additionally, air-dry the plates for 15 min in a laminar flow cabinet.
Stain the attached biofilm material by adding 300 μL of 0.1% crystal violet solution (125 μL for a 96-well plate) to each well and let stand for 10 min at room temperature.
Remove the crystal violet solution by pipetting out and rinse out unbound crystal violet with distilled water until all wells are free of liquid crystal violet (see Note 6).
Tap the plates over the paper towel and leave them face up on the bench overnight at room temperature to dry or air-dry for 15 min in laminar flow cabinet.
Add 500 μL (or 200 μL for a 96-well plate) of MBDS to each well to solubilize the crystal violet and incubate for 10 min at room temperature.
Mix the MBDS and crystal violet in the wells by pipetting up and down.
Transfer 125–200 μL of the MBDS/crystal violet solution from each well into a corresponding well of a flat-bottomed 96-well plate.
Quantify OD (570–600 nm) of each well in a plate reader in the flat-bottomed plate.
Subtract the measurement for blank wells (MBDS only) from the OD of each well that contained a sample/concentration and calculate the average.
Normalize the average to the percentage of biofilm inhibition and dispersion (%) as described in [6, 27, 28]:
% = (Con - Exp)/Con) × 100
Where Con = Control (Untreated) and Exp = Experimental (Treated).
CLSM microscopy (Figure 3)
Figure 3. Confocal laser scanning microscopy (CSLM) images of dual-species biofilms with Campylobacter jejuni and Pseudomonas aeruginosa. C. jejuni/P. aeruginosa biofilm (48 h) imaged using dual fluorescence labeling, DAPI (blue), and Thioflavin T (green) (scale bar = 20 μm).
Prepare the cells as described in steps A1–A3.
Place a coverslip into each well of the 6-well plate.
Dispense 2 mL of diluted bacterial suspension into 2 wells of the 6-well plate to enable the formation of biofilm on the coverslip.
Incubate plates at 42 °C under microaerobic condition without shaking for at least 24 h and up to 72 h (see Notes 1 and 2).
Remove the media from plates over a tray and rinse gently with PBS twice to remove planktonic cells.
Fix the cells on the coverslips using a 5% formaldehyde solution for 1 h at room temperature, rinse gently with 2 mL of PBS, and stain with fluorescent dyes (e.g., DAPI) that label DNA [6].
DAPI is usually used to visualize bacterial cell distribution in the biofilm [29] and is often included in the anti-fade mounting medium.
Remove the coverslips using forceps, carefully hold the coverslip edge to avoid disrupting the biofilm, and invert onto a glass slide containing a drop of the anti-fade mounting medium with DAPI (10 μg/mL) (see Note 7).
Seal coverslips mount on glass slides using transparent nail varnish. Two coverslips per sample/concentration from at least two separate experiments should be examined microscopically.
All images are processed using ImageJ [30]. ImageJ was used to combine/overlay two images [e.g., DAPI and Thioflavin T (green)] into a single image to generate a representative image as shown inFigure 3. Briefly, open the images in ImageJ and adjust the contrast (go to Image > Adjust > Brightness). To combine two/three images into a single-color image, go to Image > Color > Merge channels; each image represents one of the three channels (red, green, and blue) and the merge channels box will appear. Then, select the channels that you want and click OK to create the composite image; save it as .tiff file.
Notes
To create a humid environment for the biofilm assay when using 24-well plates, place a small container of water and a stack of wet paper towels around the plate during incubation.
If using a 96-well plate, add 200 μL of sterile water to each outer well to prevent drying.
All solutions should be prepared in ultrapure water and all reagents should be stored at room temperature (unless otherwise stated).
It is important to note that there are factors that can affect the results of this assay, such as the ability of the organism to adhere to the surface. For example, C. jejuni 81116 forms more abundant biofilms than C. jejuni 11168-O. In addition, C. jejuni strains display flagellin phase variation, and it is important to make sure that the culture is motile (e.g., check the cell motility by using “wet drop” light microscopy) prior to seeding of the plates, as non-motile variants do not attach nor form biofilms. Briefly, the wet mount is made by placing a drop of water on a microscopic slide and suspending a single colony of the selected strain into the water drop. The suspension is then covered with a cover slide and examined via light microscopy.
Each test plate should contain both a strong positive control (i.e., a wild-type strain, untreated sample) and a negative control (e.g., inoculated medium). Higher variability can appear following the incubation due to dryness or overwashing (detaching of cells).
Make sure that the only remaining crystal violet is bound to the biofilm at the bottom of the well. A crystal purple ring around the well is not indicative of biofilm formation and should be rinsed again as overstaining can affect assay results.
Using a sterile needle and forceps, carefully remove the coverslip from the 6-well plate and fix it with 5% formaldehyde solution for 1 h at room temperature.
Data analysis
Expected results
Here, we introduce two assays—inhibition and dispersion assays—to evaluate the effect of natural compounds (e.g., DAs). Treatment of C. jejuni culture with the DA D-Ser showed a significant inhibitory effect (p < 0.001) on biofilm formation (Figure 4) in a dose-dependent manner. Also, D-Ser had a significant disruptive effect on the existing biofilm (p < 0.001), up to 71% at 50 mM (Figure 4). Both assays can be used to screen the inhibitory and dispersal effect of selected compounds on biofilms and determine the right concentration for any further investigations. We have tested the C. jejuni 81–176 (81–176) andC. jejuni 81116 (81116) [6] strains with this protocol.
Figure 4. Effect of D-serine (D-Ser) on Campylobacter jejuni 11168-O biofilm. A) Inhibition of biofilm formation in the presence of D-Ser at different concentrations. B) Dispersion of the existing biofilm induced by different concentrations of D-Ser. The asterisk (*) indicates a statistically significant difference using the unpaired Student’st-test, p < 0.05. At least three biological replicates were included for each experiment and each biological replicate included at least three technical repeats.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Elgamoudi et al. (2020). Inhibition of Campylobacter jejuni Biofilm Formation by D-Amino Acids. Antibiotics (Figures 2 and 3). Data are representative of three independent experiments (n = 3), and values are presented as mean ± standard errors. Statistical significance of data generated in this study was determined using two tailed Student’s t-test. p ≥ 0.05 was considered statistically significant.
Acknowledgments
This protocol was derived from and validated in the original research papers (Elgamoudi et al., 2020 [6]; Elgamoudi et al., 2022 [31]). The work has been partially supported by Griffith University.
Author contributions: Bassam Elgamoudi: Conceptualization, Methodology, Validation, Formal analysis, Data curation, Writing: original draft, Writing: review & editing. Victoria Korolik: Conceptualization, Writing: original draft, Writing: review & editing.
Competing interests
The authors declare no conflicts of interest.
References
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4,867 | https://bio-protocol.org/en/bpdetail?id=4867&type=0 | # Bio-Protocol Content
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Peer-reviewed
Studying Cellular Focal Adhesion Parameters with Imaging and MATLAB Analysis
LY Ling-Yea Yu *
TT Ting-Jeng Tseng
HL Hsuan-Chao Lin
CH Chi-Lin Hsu *
TL Ting-Xuan Lu
YL Yu-Chiao Lin
MT Miranda Tseng
FT Feng-Chiao Tsai
(*contributed equally to this work)
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4867 Views: 525
Reviewed by: Alka Mehra Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances Jul 2021
Abstract
Cell signaling is highly integrated for the process of various cell activities. Although previous studies have shown how individual genes contribute to cell migration, it remains unclear how the integration of these signaling pathways is involved in the modulation of cell migration. In our two-hit migration screen, we revealed that serine-threonine kinase 40 (STK40) and mitogen-activated protein kinase (MAPK) worked synergistically, and the suppression of both genes could further lead to suppression in cell migration. Furthermore, based on our analysis of cellular focal adhesion (FA) parameters using MATLAB analysis, we are able to find out the synergistic reduction of STK40 and MAPK that further abolished the increased FA by shSTK40. While FA identification in previous studies includes image analysis using manual selection, our protocol provides a semi-automatic manual selection of FAs using MATLAB. Here, we provide a method that can shorten the amount of time required for manual identification of FAs and increase the precision for discerning individual FAs for various analyses, such as FA numbers, area, and mean signals.
Keywords: Focal adhesion (FA) Serine-threonine kinase 40 (STK40) Cell signaling Mitogen-activated protein kinase (MAPK) Mean signal Thresholding Segmentation FA selection
Background
Our protocol can be used to identify focal adhesion (FA) molecules such as paxillin, talin, or vinculin. Images of FA molecules are usually taken by a confocal fluorescent microscope (Doss et al., 2020) or through super-resolution fluorescence microscopy (Kanchanawong et al., 2010) to show clear images. Our protocol allows fast imaging using fluorescent microscopy, acquiring a large number of images for FA identification and quantification. Selecting FAs using prior methods such as ImageJ requires researchers to manually select all FAs by the naked eye. This creates great deviations between different operators and even different times of selection by the same operator. The semi-automatic manual selection provides a time-saving and precise method, running MATLAB scripts to select all FAs in images. Researchers may then select specific FAs for further analysis. By using MATLAB software instead of the traditional analysis by ImageJ plugins (Horzum et al., 2014), we are able to analyze different FA parameters, such as number, area, and signal intensity in a semi-automatic manual method. Furthermore, we chose paxillin for the representation of FA due to its clearer immunofluorescent staining under fluorescent microscope compared to other FA proteins, such as talin and vinculin. By understanding the general properties of FA followed by different treatments, such as STK40 knockdown, we can further explore subsequent cellular mechanisms.
Materials and reagents
Cell culture
Cells
SAS and HSC-3 cells [gifts from J. S. Chia’s Lab (National Taiwan University, Taipei, Taiwan)]
HepG2 and HEK-293T cells [American Type Culture Collection (Manassas, VA, USA)]
HUVEC cells (Lonza, Basel Stücki, Switzerland)
HaCaT cells [American Type Culture Collection (Manassas, VA, USA)]
Culture medium
Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, Thermo Fisher Scientific, catalog number: 11965092 and HyClone, catalog number: 16750-074) for SAS, HSC-3, HepG2, and HEK-293T cells
Endothelial cell growth medium-2 (EGM2) (Lonza, catalog number: CC-3162) for HUVEC cells
1% of penicillin/streptomycin (P/S) (Gibco, catalog number: 15140122)
10% fetal bovine serum (FBS) (HyClone, catalog number: 12389802)
0.5% trypsin-EDTA (Gibco, catalog number: sc-363354)
Phosphate-buffered saline (PBS) (Corning, catalog number: 21-040-CM)
Coating and infection
96-well plate (Thermo Fisher Scientific, NuncTM, catalog number: 165305)
Coated chamber slides [Thermo Fisher Scientific, catalog number: 155411 (4-well) or Nunc Lab-TEK, catalog number: 155383]
6-well plate (Jetbiofil, catalog number: TCP011006)
Poly-D-lysine (Sigma-Aldrich, catalog number: A3890401)
Collagen (100 μg/mL; collagen I, bovine) (Gibco, catalog number: A1048301)
Lentiviruses of control or STK40 [shRNA plasmids bought from National RNAi Core (NRC; Academia Sinica, Taipei, Taiwan); plasmids of shSTK40 were transfected into HEK-293T cells to produce lentiviruses while control lentiviruses were directly bought from NRC]
Puromycin (2 μg/mL) (Sigma-Aldrich, catalog number: A1113803)
Blasticidin (10 μg/mL) (Sigma-Aldrich, catalog number: 15205)
Reagents used prior to immunofluorescent staining
20 mM HEPES (Gibco, catalog number: 15630080)
0.1% BSA (BioShop, catalog number: ALB001)
25 ng/mL FGF1 (Invitrogen, catalog number: PHG0244)
10 U heparin (Sigma, catalog number: H3393)
Drugs
Trametinib 100 nM (LC laboratories, catalog number: T-8123)
Y27632 5 μM (Sigma-Aldrich, catalog number: 688000)
Blebbistatin 5 μM (Sigma-Aldrich, catalog number: 203390)
Verteporfin 2.5 μM (MedChemExpress, catalog number: HY-B0146)
Leptomycin B 100 nM (Cayman Chemical Company, catalog number: 10004976)
Immunofluorescent staining
4% paraformaldehyde (Sigma-Aldrich, catalog number: 158127)
0.25% Triton X-100 (J.T. Baker, catalog number: 10421871)
Primary antibody: purified mouse anti-PXN (BD Biosciences, catalog numbers: 610051 and 610052) in 1% BSA. Paxillin was chosen as the optimal representation of FA due to its clearer visibility compared to other antibodies such as talin and vinculin
5% BSA for blocking
4’,6-diamidino-2-phenylindole (DAPI); 10 μg/mL (Invitrogen, catalog number: D1306)
Secondary antibodies: goat anti-mouse immunoglobulin G (IgG) (H + L) cross-adsorbed secondary antibody and Alexa Fluor 488 (Invitrogen, catalog number: A11001) at 1:500; goat anti-mouse IgG (H + L) cross-adsorbed secondary antibody and Alexa Fluor 594 (Invitrogen, catalog number: A11005) at 1:500 in 1% BSA
Equipment
Microscope (Nikon Eclipse Ti)
Camera (Nikon DS-Qi2)
Excitation light source: X-cite 120 (Excelitas Technologies Corp.)
Filter cubes (Chroma Technology Corporation)
DAPI filter set (excitation wavelength: 360 nm; dichroic mirror wavelength: 400 nm; emission wavelength: 460 nm)
Fluorescein isothiocyanate (FITC) filter set (excitation wavelength: 480 nm; dichroic mirror wavelength: 505 nm; emission wavelength: 535 nm)
Texas Red (Tx-Red) filter set (excitation wavelength: 560 nm; dichroic mirror wavelength: 595 nm; emission wavelength: 630 nm)
Yellow fluorescent protein (YFP) filter set (for over-expression) (excitation wavelength: 500 nm; dichroic mirror wavelength: 520 nm; emission wavelength: 542 nm)
Autofluorescent plastic slides (Chroma Technology Corporation, catalog number: 92001)
Software
MATLAB (MathWorks, Natick, MA, USA)
Install MATLAB Image Processing Toolbox to run script.
Microsoft Excel (Redmond, WA, USA)
Procedure
Preparation for immunofluorescence fixation
Seed SAS, HaCaT, HUVEC, or HepG2 cells on culture medium in a 6-well plate. For SAS and HaCaT cells, seed 3 × 105 cells/well. Seed cells at 60% confluency. For SAS, HaCaT, and HepG2 cells, use DMEM with 1% penicillin/streptomycin, 10% FBS, and 0.5% trypsin-EDTA as culture medium. For HUVEC cells, use EGM2 as culture medium.
On the next day, transduce lentiviruses that knockdown or over-express STK40 into seeded SAS, HaCaT, HUVEC, or HepG2 cells. The lentiviruses are generated by transfecting vectors of (1) shRNA plasmid targeting STK40 bought from Academia Sinica or (2) plasmid of over-expression construct targeting STK40 into HEK-293T cells. For our over-expression construct of STK40, YFP is tagged at the cytoplasmic domain of STK40 to ensure fluorescent visibility under the microscope.
Select cells in a 6-well plate with antibiotics such as puromycin or blasticidin over 24–48 h.
On the same day of antibiotics selection, perform collagen coating in a new 96-well plate or 4- or 8-well chamber slide. Coat poly-D-lysine 100 μg/mL in PBS for 5 min at 37 °C. Wash SAS, HaCaT, HUVEC, or HepG2 cells with PBS and then coat cells with bovine collagen 100 μg/mL in PBS. Collagen coating ensures better cell attachment for cell migration and subsequent observation of focal adhesion. Incubate chamber slides or 96-well plates at 37 °C overnight.
On the next day, re-seed cells from the 6-well plate onto the coated 96-well plate or 4- or 8-well chamber slide. To observe nascent and prominent FAs, the transduced cells are re-seeded differently:
To observe nascent FAs: seed cells at a higher density at 70%–80% confluency to perform wound scratch. The FAs on the first row of the wound will be analyzed later. For SAS cells, seed 2–3 × 104 cells per 96-well plate and 1.5 × 105 cells per 4-well chamber slide. For HaCaT cells, seed 1 × 104 cells per 96-well plate and 5 × 104 per 4-well chamber slide.
To observe prominent FAs: seed cells at a lower density at 40% confluency. Cells will perform random migration and all of their prominent FAs will be analyzed later. For SAS cells, seed 6,000 cells per 96-well plate and 3 × 104 cells per 4-well chamber slide. For HaCaT cells, seed 5,000 cells per 96-well plate and 2.5 × 104 cells per 4-well chamber slide. The prominent FAs of HaCaT cells are shown as demonstration in this protocol.
Incubate cells at 37 °C in supplemented medium with or without drugs according to their groups. The supplemented medium contains 20 mM HEPES, 0.1% BSA, 25 ng/mL FGF1, and 10 U of heparin. The experimental groups include untreated and drug-treated groups.
Untreated group.
Drug-treated groups:
i. Blebbistatin (5 μM): inhibits myosin II.
ii. Y27632 (5 μM): inhibits ROCK (Rho-kinase). Blebbistain or Y27632 are added to inhibit force-mediated FA. The FA results from the co-inhibition of STK40, and force-generating FA tell us whether the effect of STK40 on FA is due to force-mediated FA strengthening.
iii. Trametinib (100 nM): inhibits MAPK kinase. The FA results from the co-inhibition of STK40, and MAPK kinase tell us whether MAPK is involved in the effect of STK40 on FA.
iv. Leptomycin (100 nM): inhibits exportin to block nuclear exportation. The blockage of nuclear export by leptomycin renders the accumulation of our other target protein, YAP (Yes-associated protein), in the nucleus. The FA results of the co-inhibition of STK40 and exportin provide partial clues for us to explore whether the spatial distribution of YAP is involved in the effect of STK40 on FA.
v. Verteporfin (2.5 μM): inhibits YAP activity. The FA results from the co-inhibition of STK40, and YAP tell us whether YAP is involved in the effect of STK40 on FA. Furthermore, the FA results of the inhibition of STK40, YAP, and MAPK support the STK40-YAP-MAPK system on FA in our published paper (Yu et al., 2021).
To observe nascent and prominent FAs, the duration of drug treatment is different.
To observe nascent FAs, treat cells with drugs as follows:
i. 5 μM blebbistatin, 5 μM Y27632, or 100 nM trametinib: overnight
ii. 100 nM leptomycin B: 1 h
iii. 2.5 μM verteporfin: 6 h.
Go to steps 7–10 in this section to create wound scratch.
To observe prominent FAs, treat cells with drugs as follows:
i. 5 μM blebbistatin, 5 μM Y27632, or 100 nM trametinib: overnight
ii. 100 nM leptomycin B: 3 h
iii. 2.5 μM verteporfin: 8 h.
Go directly to section B for fixation.
After incubation, keep the media by transferring it to a new 96-well plate or chamber slide. Wash cells with PBS.
Create wounds with a tip for chamber slides and a scratcher for 96-well plates in cells immersed in PBS. A horizontal wound is created in the middle of the well for 96-well plates. To create wounding for chamber slides, draw a cross to create two wounds in each well.
After wounding, put back media onto the cells.
Incubate cells at 37 °C for another 2 h. During this period, cells migrate towards the wound, as shown in Video 1. The video demonstrates SAS cells wound-healing migration for 10 h. The purpose of the 2 h migration is to ensure that the cells migrate towards the wound to form nascent FAs (Olson and Nechiporuk, 2021). The lamellipodia of migrating cells towards the wound are stretched out. Nascent FAs are visible under this condition.
Video 1. Wound-healing migration of SAS cells for 10 h
Immunofluorescence fixation and staining
Fix cells with 4% paraformaldehyde in PBS at room temperature for 15 min.
Permeate cells with 0.25% Triton X-100 at room temperature for 10 min.
Block cells with 5% BSA at room temperature for 1 h.
Incubate cells with primary antibodies in 1% BSA in PBS overnight at 4 °C. Anti-PXN is used as primary antibody to stain FA.
Stain cells with secondary antibodies and 10 μg/mL of DAPI (10 μg/mL) in 1% BSA in PBS.
Use a Nikon Eclipse Ti for FA imaging. Focus on the FAs of the fixed cells and take z-stack images. Take images of FA with a step size of 1 μm. We took seven z-stack images with a total range of 5 μm for our analysis (Figure 1A).
Figure 1. Z-stack images of prominent focal adhesion (FA) of HaCaT cells. We took seven Z-stack images of FA with the step size of 1 μm. (A) The original fluorescence images. The images are listed sequentially from left to right. Their file names are: adj_xy04c1z02.tif, adj_xy04c1z03.tif, adj_xy04c1z04.tif, adj_xy04c1z05.tif, adj_xy04c1z06.tif, adj_xy04c1z07.tif and adj_xy04c1z08.tif. (B) The images of FA with their background subtracted. Their file names are: adj_bs_xy04c1z02.tif, adj_bs_xy04c1z03.tif, adj_bs_xy04c1z04.tif, adj_bs_xy04c1z05.tif, adj_bs_xy04c1z06.tif, adj_bs_xy04c1z07.tif and adj_bs_xy04c1z08.tif. Scale bars: 50 μm.
Data analysis
Immunofluorescent images of FA are analyzed using MATLAB software written scripts: FA1_save_fluoref_cor_img.m, FA2_FA_background_20200313.m, and FA3_FA_identification_FAclearer_20200316chiao.m. The overall pipeline of data analysis is shown in Figure 2. We utilized the cell line HaCaT as our demonstration of FA analysis. We adjusted cells and FA contrast equally for better visualization. All seven Z-stack images (adj_xy04c1z02.tif, adj_xy04c1z03.tif, adj_xy04c1z04.tif, adj_xy04c1z05.tif, adj_xy04c1z06.tif, adj_xy04c1z07.tif, and adj_xy04c1z08.tif) are included in our data analysis. These images are in the zip folder “Adjusted_xy04.zip.” The seven images are shown in Figure 1A.
Figure 2. Pipeline of data analysis. Process of the overall analysis of focal adhesion (FA). Script 1 refers to “FA1_save_fluoref_cor_img.m,” script 2 refers to “FA2_FA_background_20200313.m,” and script 3 refers to “FA3_FA_identification_FAclearer_20200316chiao.m.”
Calibration of fluorescence
Calibrated fluorescent images of FA (FITC for FA in our demonstration) are obtained by correcting the original fluorescent images with reference images. The reference images are taken with fluorescent light on and off under the autofluorescent plastic slide (Figure 3 and Figure 4). The calibration of fluorescence is required due to the change of illumination by LED excitation light under confocal microscopes. Inconsistency and unevenness occur, reducing the fluorescence on the edges of sample images. This calibration step ensures that the fluorescence in our analyses is closest to the actual FA fluorescence of the sample.
To calibrate the fluorescence of images, load the original FA images and reference images. Then, run the first script (“FA1_save_fluoref_cor_img.m”) to crop the edges of our FA images for analysis.
Figure 3. Representative images of autofluorescent plastic slides. (A) All the slides in the slide storage box. (B) (left to right, up to down): YFP, FITC, TX-Red, and DAPI slides. The transparent slide is used for creating compartments in the slide storage box.
Figure 4. Reference images utilized for calibration of fluorescence. (A) Off: reference image taken with lights off. (B) DAPI: reference image taken under DAPI autofluorescent plastic plate and DAPI light on. (C) FITC: reference image taken under FITC autofluorescent plastic plate and FITC light on. For tif files of these images, see zip file “reference_images.zip.” The names of the files are: off.tif, dapi.tif, and fitc.tif.
FA background subtraction
The background signal of FA is subtracted using the second script (“FA2_FA_background_20200313.m”). The purpose of carrying out background subtraction is to eliminate the background noises of the images, contrasting FAs of the cells with the background to ensure better discerning. To carry out background subtraction, we first determine the background signal of each individual pixel. Each signal of each pixel is separately determined by the median value of its surrounding pixels within the radius of 1.46 μm. Then, we subtract the background signal by its original signal by running the script, completing background subtraction. Images adj_bs_xy04c1z02.tif, adj_bs_xy04c1z03.tif, adj_bs_xy04c1z04.tif, adj_bs_xy04c1z05.tif, adj_bs_xy04c1z06.tif, adj_bs_xy04c1z07.tif, and adj_bs_xy04c1z08.tif are generated after running the script. These images are also in the zip folder “Adjusted_xy04.zip.” The contrast of the images of FAs was adjusted equally, subtracting their background signal.
Thresholding: The purpose of thresholding for the mask of FA is to allow clear identification of Fas for quantification and qualification. The threshold is determined as two standard deviations above the mode of background value.
Segmentation: To distinguish each FA from one another for analysis, segmentation is required. In other words, the process of segmentation prevents repeated analysis of the same FA. To be specific, double analysis of a single FA generates results for two separate Fas with identical data in all parameters. To carry out FA segmentation, use the function of watershed segmentation in MATLAB.
FA analysis
FA analysis is processed using the third script: “FA3_FA_identification_FAclearer_20200316chiao.m.” Steps to run the script are as follows:
Cells are selected differently to analyze nascent or prominent FAs:
i. Nascent FA: The FAs of the first row of the wound are selected for analysis. Select only the cells with adjacent cells at their two sides. These are the cells that perform directed cell migration towards the wound. Nascent FAs forming in the lamellipodia of these cells at the leading edges of the wound are analyzed.
ii. Prominent FA: All cells with clear FAs are selected for analysis. We analyze the prominent FAs of HaCaT as demonstration.
For both analyses of nascent and prominent FAs, all of the FAs within 7 μm (15 pixel) of the lamellipodia edge are analyzed per run. The detailed steps are listed below.
Circle target cell, then run script:
Circle the target cell manually by pinpointing the surroundings of the cell in the panel of Figure 5A. Run script, which loads in the original fluorescent image (Figure 5B). The script will show again the background-subtracted image of the selected target cell (Figure 5C). Afterwards, the script automatically identifies all the FAs (Figure 5D). This will be the background-subtracted image that the script will analyze later on.
Figure 5. HaCaT cells are selected utilizing MATLAB scripts to analyze prominent focal adhesion (FA). Individual HaCaT cells are selected manually by pinpointing the surrounding of the cell. (A–B) Images of cells after performing background subtraction (A) and the original FA image (B). Blue crosses of (A) indicate selected cell. (C–D) Image of selected cell after background subtraction shows up again in (C). MATLAB script then automatically identifies FAs in (D). Scale bars: 50 μm.
Determine lamellipodia border:
We pinpoint the border of the cell showing clear lamellipodial protrusion toward the wound to determine lamellipodia border of target cell.
Determine nuclear center:
Nucleus of the target cell is marked so that the clear lamellipodium for FA analysis is completely determined. The completion of determining lamellipodia border and nuclear center is shown in Figure 6A–6C. The red dash line demonstrates the lamellipodia border and the nucleus center from which the lamellipodia border is determined.
Figure 6. Focal adhesions (FA) of chosen cells are then selected for further analysis. (A–C) The nucleus center and lamellipodia border are determined. FA under background subtraction (A); red dash lines indicate the chosen cell with determined nucleus (B) and lamellipodia border (C). (D–G) Thresholding of FAs after background subtraction and FA numbering of chosen cell. Green dash lines indicate the chosen cell with determined nucleus and lamellipodia border (D). Yellow X indicates manually selected FAs (E) with its corresponding red dots (F) of actual FAs in the selection. FAs are numbered according to sequence of selection (G). Scale bars: 10 μm.
FA selection:
Under determined lamellipodium, all FAs are selected manually for analysis (Figure 5D to 5G). FAs within 7 μm of the lamellipodia edge are analyzed. After analysis, FA signal intensities are generated as shown in the heatmap of Figure 7. In this process, the script corrects the FA signal to get rid of the effects of photobleaching. The generated heatmap is shown on Figure 7; the left panel demonstrates original FA signal (Ori Cell x87y101), while the right panel depicts corrected FA signal (Cor Cell x87y101). The data of corrected FA signal with its photobleaching corrected will then be utilized for final FA analysis.
Figure 7. Heatmap of focal adhesion (FA) signal intensity. Left panel: OriCellx87y101 indicates original FA signal intensity before photobleaching correction. X87 and y101 indicate x and y axis of selected target cell. CorCellx87y101 indicates corrected FA signal intensity, depicting the signals processed with photobleaching correction. FA# depicts the sequential numbering of the 11 FA selected manually in Figure 6. Frame depicts the seven z-stack images taken for analysis.
Generate data of different FA parameters:
Complete script running. Different FA parameters for quantification and qualification are generated: (a) FA location, (b) FA eccentricity, (c) FA area, (d) mean FA signal, and (e) FA numbers. The integrated FA is utilized for FA analysis. The formula for integrated FA is as follows:
Integrated FA = FA area × mean FA signal
*The integrated FA is generated by multiplying (c) by (d).
*(a) FA location: coordinates x and y are generated.
*(b) FA eccentricity determines whether the FA is closer to circular or linear shape. Numeric value ranging from 0 to 1 is generated after running the script. Larger numbers symbolize greater eccentricity.
Organize FA data:
Data of FAs and cells from multiple sites analyzed are pulled up and organized using Microsoft Excel. See Excel file “xy04.” The Excel file is all the FA data of all cells of the well xy04. The highlighted portion demonstrates the FA data of our demo cell, x87y101.
Acknowledgments
This work was supported by grants from The Ministry of Science and Technology in Taiwan (MOST 107-2320-B-002-038-MY3 and MOST 108-2926-I-002-002-MY4), National Taiwan University Hospital (NTUH 106-T02, NTUH 107-T13, NTUH 108-T13, and VN 109-14), and The Liver Disease Prevention and Treatment Research Foundation in Taiwan. This protocol was adapted from Yu et al. (2021).
Competing interests
The authors declare that they have no competing interests.
References
Doss, B. L., Pan, M., Gupta, M., Grenci, G., Mège, R.-M., Lim, C. T., Sheetz, M. P., Voituriez, R. and Ladoux, B. (2020). Cell response to substrate rigidity is regulated by active and passive cytoskeletal stress. Proc. Natl. Acad. Sci. U. S. A. 117(23): 12817–12825.
Kanchanawong, P., Shtengel, G., Pasapera, A. M., Ramko, E. B., Davidson, M. W., Hess, H. F. and Waterman, C. M. (2010). Nanoscale architecture of integrin-based cell adhesions. Nature 468(7323): 580–584.
Horzum, U., Ozdil, B. and Pesen-Okvur, D. (2014). Step-by-step quantitative analysis of focal adhesions. MethodsX 1: 56–59.
Yu, L. Y., Tseng, T. J., Lin, H. C., Hsu, C. L., Lu, T. X., Tsai, C. J., Lin, Y. C., Chu, I., Peng, C. T., Chen, H. J., et al. (2021). Synthetic dysmobility screen unveils an integrated STK40-YAP-MAPK system driving cell migration. Sci. Adv. 7(31): eabg2106.
Olson, H. M. and Nechiporuk, A. V. (2021). Lamellipodia-like protrusions and focal adhesions contribute to collective cell migration in zebrafish. Dev. Biol. 469: 125–134.
Supplementary information
The following supporting information can be downloaded here:
Adjusted_xy04 files
MATLAB files
reference images
xy04.xlsx
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Preparation of Whole-mount Mouse Islets on Vascular Extracellular Matrix for Live Islet Cell Microscopy
KH Kung-Hsien Ho
GG Guoqiang Gu
IK Irina Kaverina
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4868 Views: 395
Reviewed by: Pilar Villacampa AlcubierreAlba Casellas Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Nov 2021
Abstract
Pancreatic islet β cells preferentially secrete insulin toward the plasma membrane, making contact with the capillary extracellular matrix (ECM). Isolated islets separated from the exocrine acinar cells are the best system for cell biology studies of primary β cells, whereas isolated islets lose their capillary network during ex vivo culture. Providing the appropriate extracellular signaling by attaching islets to vascular ECM-coated surfaces can restore the polarized insulin secretion toward the ECM. The guided secretion toward ECM-coated glass coverslips provides a good model for recording insulin secretion in real time to study its regulation. Additionally, β cells attached to the ECM-coated coverslips are suitable for confocal live imaging of subcellular components including adhesion molecules, cytoskeleton, and ion channels. This procedure is also compatible for total internal reflection fluorescence (TIRF) microscopy, which provides optimal signal-to-noise ratio and high spatial precision of structures close to the plasma membrane. In this article, we describe the optimized protocol for vascular ECM-coating of glass coverslips and the process of attachment of isolated mouse islets on the coverslip. This preparation is compatible with any high-resolution microscopy of live primary β cells.
Key features
• Optimized coating procedure to attach isolated islets, compatible for both confocal and TIRF microscopy.
• The ECM-coated glass coverslip functions as the artificial capillary surface to guide secretion toward the coated surface for optimal imaging of secretion events.
• Shows the process of islets attachment to the ECM-coated surface in a 6-day ex vivo culture.
Graphical overview
Keywords: Pancreatic β cells Islet attachment Islet ex vivo culture Islet flattening Human ECM Live islet cells visualization
Background
Pancreatic β cells regulate blood glucose homeostasis by secreting insulin to the blood stream in response to elevated glucose levels. The secretion of insulin is tightly regulated to prevent over-secretion, which causes hypoglycemia, or under-secretion, which in the long term leads to diabetes. Islet β cells preferentially secrete insulin toward the plasma membrane, making contact with the capillary extracellular matrix (ECM) or basement membrane (Low et al., 2014; Gan et al., 2018). Isolated islets lose their capillaries and the polarity of β cells during ex vivo culture. When studying β-cells secretion, recording secretion events on the plasma membrane making contact with the vascular ECM can better recapitulate the polarized insulin secretion of β cells in vivo. Coating coverslips with human ECM, derived from placenta and containing the main components of vascular ECM, is an ideal surface for islet attachment to study insulin secretion (Patterson et al., 2000; Arous and Wehrle-Haller, 2017; Barillaro et al., 2022). The coated coverslip functions as the artificial capillary surface to guide the secretion of attached islet β cells toward it and is suitable for high-resolution imaging to record secretion events.
Total internal reflection fluorescence (TIRF) microscopy is a powerful tool to study subcellular structures in attached β cells, such as adhesion molecules, cortical actin cytoskeleton, ion channels, and lipids in the plasma membrane. Another powerful application of TIRF microscopy is to record insulin secretion from individual β cells in real time. Multiple studies have reported different approaches to labeling insulin vesicles (Ma et al., 2004; Ivanova et al., 2013; Schifferer et al., 2017). Integration of an engineered insulin fusion probe to the genome of β cells in isolated rodent or human islets is usually technically challenging. Label-free approaches bypass this challenge and the possibility that a fluorescence probe may interfere with the maturation or secretion of insulin (Pouli et al., 1998; Schifferer et al., 2017). Successful label-free recording of insulin exocytosis has been reported using either two-photon microscopy or correlative scanning ion conductance microscopy-fluorescence confocal microscopy (Takahashi et al., 2002; Bednarska et al., 2021). Our lab has previously established a novel label-free approach to record insulin secretion using TIRF microscopy and a cell-impermeable zinc-sensitive fluorescence dye, FluoZinTM-3 (Gee et al., 2002; Zhu et al., 2015). The fusion of individual insulin vesicles to the plasma membrane transiently elevates local zinc concentration, which activates FluoZinTM-3 to emit fluorescence upon excitation. This strategy allows visualization of insulin secretion at the cell–ECM interface with high spatial precision in real time. The conventional coating procedure for islet attachment and confocal microscopy includes a layer of matrigel between the coverslip and the ECM-coating. The matrigel layer provides a three-dimensional-like cushion to facilitate islet attachment but is not compatible with TIRF microscopy due to its thickness. In this article, we describe the details of an optimized ECM-coating and attachment of isolated islets on glass coverslips, which is compatible with TIRF microscopy and any high-resolution live-cell microscopy.
Materials and reagents
Mice [GFP-Lifeact; Ins2Apple, 12–20 weeks (Riedl et al., 2008; Stancill et al., 2019)]
Human extracellular matrix (ECM) (Corning, catalog number: 354237)
Fetal bovine serum (FBS) (Atlanta Biologicals, catalog number: S11550)
RPMI 1640 (Gibco, catalog number: 11875)
Penicillin-Streptomycin solution (Pen-Strep) (Gibco, catalog number: 15140)
NalgeneTM Rapid-FlowTM Sterile Disposable Filter Units, 0.2 μm PES membrane (Thermo Fisher Scientific, catalog number: 569-0020)
Hanks’ Balanced Salt Solution with calcium & magnesium (HBSS) (Gibco, catalog number: 21-020-CV)
Collagenase from Clostridium histolyticum (Millipore Sigma, catalog number: C5138, lot number: 0000164940)
Islet media (see Recipes)
Collagenase solution (see Recipes)
ECM coating solution (see Recipes)
Recipes
Islet media
RPMI-1640 (the media contains 11 mM glucose)
10% heat-inactivated FBS
100 U/mL penicillin-100 mg/mL streptomycin
Filter the media through a 0.2 μm PES membrane, store at 4 °C, and used within one month of preparation.
Collagenase solution
0.5 mg/mL collagenase in HBSS
Store aliquots (1 mL) of the prepared solution at -20 °C for up to one year.
ECM coating solution
9 μg/mL ECM in islet media
Store aliquots (100 μL) of the prepared solution at -20 °C for up to two years.
Equipment
Plasma cleaner (Harrick Plasma, model: PDC-001)
Nikon A1R laser scanning confocal microscope
CFI Apochromat TIRF 100×/1.45 oil objective (Nikon)
Olympus SZH10 dissection scope
MatTek glass bottom dishes, 35 mm dish, 10 mm microwell (MatTek, catalog number: P35G-1.5-10-C)
150 mm dish (Corning, catalog number: 430599)
Tissue wipers (VWR, catalog number: 82003-820)
5 mL syringe (BD, catalog number: 309646)
Needle [BD, catalog numbers: 305109 (27 G 1/2) and 305106 (30 G 1/2)]
50 mL centrifuge tubes (VWR, catalog number: 525-1075)
Precision water bath (Precision Scientific, Model 182)
60-mm polystyrene Petri dish (Thermo Fisher Scientific, catalog number: AS4052)
Water-jacked CO2 incubator (NuAire, model: NU-8700 series 5)
Procedure
Day 0
The collagenase digestion of mouse pancreas and islet isolation follows the published procedure (Li et al., 2009; Ho et al., 2023). Briefly, mouse pancreata are injected with 2 mL of collagenase solution (see Recipes) using a 5 mL syringe and a 27 G 1/2 needle (or a 30 G 1/2 needle for smaller mice) through the common bile duct. The pancreata are digested in a 50 mL centrifuge tube in a water bath at 37 °C for 16 min with gentle shaking every 4 min. The digested homogenate is washed using 10 mL (per animal’s pancreata) of chilled islet media (see Recipes) with vigorous shaking and centrifuged at 700× g for 2 min at 4 °C. Islets are hand-picked using a P20 pipette with 200 μL tips to a 60 mm Petri dish with islet media under a dissection scope. Repeat the picking three times to separate islets from pancreatic acinar cells (Note 1). The isolated islets are incubated in a water-jacked incubator with 5% CO2 (Note 2).
Remove the lid and place the 35 mm MatTek dish with a 10 mm microwell in the plasma cleaner. Seal the chamber and establish vacuum. Set the RF (radio frequency) level to medium and initiate the cleaning for 1 min (Note 3). Place the lid back immediately after retrieving the dish from the cleaner. The plasma-cleaned MatTek dish can be stored in its packaging sleeve for six months at room temperature, but a coated dish is for immediate use.
Coat the coverslip of a plasma-cleaned MatTek dish by applying a small volume (~5 μL) of ECM coating solution (see Recipes) to the center of the coverslip (Figure 1A). When coating MatTek dishes, instead of measuring the exact volume of coating solution for each dish, we use a P100 pipette holding enough coating solution, push out a small amount of solution from the tip, smear it on the area to be coated, and suck excess solution back to the tip. This ensures the coating is not too thick to interfere with the downstream TIRF microscopy. We do not coat the entire coverslip to reduce the chance of an islet attaching to the edge of the microwell.
Figure 1. Illustration of coating and setup for islet attachment. (A) Schematic of coating of extracellular matrix (ECM) to the center of coverslip. (B) Schematic of MatTek dishes arrangement in a humidified 150 mm dish. (C) Schematic of media added and contained inside the microwell. (D) Side view of the MatTek dish showing the media contained inside the microwell. (E) Schematic showing two islets of different size plated per MatTek dish.
Cover the MatTek dish with its own lid and place it on the lid of a 150 mm dish. Create a humidified chamber by placing a sheet of tissue wiper wetted with distilled water on the bottom of the 150 mm dish and place the MatTek dish inside such that the tissue is at the top of the chamber (Figure 1B). Incubate at 37 °C in a water-jacked incubator for 10 min to allow ECM coating on the coverslip.
Wash the coated coverslip once by pipetting 100 μL of islet media to the microwell and aspirate it from its edge that is not coated with ECM. Refill the microwell with 100 μL of islet media (Figure 1C). During washing and refilling, the media should be contained inside the microwell. This helps to retain the surface tension of islet media and constrain islet attachment inside the microwell (Figure 1D). If the media covers the plastic dish bottom, the planted islets may float out of the microwell and attach to the plastic.
Under a dissection scope, transfer 1–2 islets to the center of the microwell using a P20 pipette (Figure 1E) (Note 4).
Place the MatTek dish in the 150 mm dish (the humidified chamber) and carefully move it to the incubator (Figure 1B). Choose islets smaller than 120 μm in diameter for better dye penetration to the space between the islet attachment surface and the coverslip in downstream imaging. Without a wet tissue wiper inside the 150 mm dish, the small volume of 100 μL of islet media will dry out overnight even in a water-jacketed incubator (Note 5).
Incubate it for two days. There is no need to replace media or take the MatTek dish out to check the status of islets before day 2 (see below). To follow the attachment process on the ECM-coated coverslip, islets isolated from the GFP-Lifeact; Ins2Apple mouse (Riedl et al., 2008; Stancill et al., 2019) are shown here. GFP-Lifeact labels cortical actin to delineate cell borders and Ins2Apple (Ins2 promoter driven H2B-mApple) marks β cells (Figure 2A). Approximately 70% of islets loosely attach to the ECM overnight (Figure 2B–2C). Taking the MatTek dish incubated overnight out for observation can easily stir up those loosely attached islets. Most islets (~90%) are firmly attached to the ECM after two nights.
Figure 2. Attachment and flattening process of isolated mouse islets. (A) Single slice of confocal microscopy image of an islet isolated from the GFP-Lifeact; Ins2Apple mouse. GFP-Lifeact (green) delineates cell borders and β-cells nuclei are labeled by Ins2Apple (magenta). (B–C) Projected images of isolated islets incubated on extracellular matrix (ECM)-coated coverslip for one day. Dashed line delineates the attachment site of an islet. Arrow indicates the side in contact with the coverslip. (D–N) Confocal images of isolated islets incubated on ECM-coated coverslip for 2–6 days. D, G, and J show projected images. E, H, and K show the side view of the projected images. F, I, and L show single slice of confocal images at the z-plane of attachment. M and N show the enlargement of image at dashed square 1 and 2 respectively. Dashed lines delineate the attachment sites of islets (D, G, and J) or represent the location of coverslips (E, H, and K). (O) Projected image of an isolated islet attached to ECM-coated coverslip and flattened to single-cell-layer thickness at the periphery of the islet. Arrows indicate the periphery of the flattened islet.
Day 2
Take the dish out from the incubator and gently add 150 μL of islet media from the edge of the microwell under a dissection scope. Because the majority of islets have now attached to the ECM on the coverslip, they are resistant to minor vibrations while moving the dish and it is okay to let the media exceed the microwell (Figure 2D–2F and 2M). Do not remove old media before adding the 150 μL of fresh media. If the media is removed and then replaced, attached islets will be lifted up above the surface of media. For this reason, there should be no media change during the entire ex vivo culture of attached islets. Instead, we add more fresh media to the dish every other day.
Incubate the attached islets for two more days.
Day 4
Gently add 500 μL of islet media from the edge of the microwell. The media will overflow from the microwell to the plastic dish bottom at this point. Islets are firmly attached to the ECM and will not float out of the microwell (Figure 2G–2I and 2N). The MatTek dish can hold 2 mL of media. The attached islets can be subjected to microscopy from day 4 to day 6.
Day 6
Islets attached firmly and further expanded their surfaces attached to the ECM. The area of attachment reaches 85%–90% of the hemispherical cross section area of an islet (Figure 2J–2L). It is noteworthy that the so-called “flattening” of isolated mouse islets refers only to the surface attached to the ECM-coated coverslip. The rest of the islet retains its spherical appearance even after 6 days of ex vivo culture. This is likely due to residual extracellular matrix (the peripheral capsule) surrounding an isolated mouse islet that maintains its spherical shape. Approximately 5% of attached islets can be fully flattened on a coated coverslip, likely due to incomplete peripheral capsule, and the islet periphery can reach single-cell-layer thickness (Figure 2O). The majority of attached islets with a hemispherical shape is suitable for live imaging using confocal or TRIF microscopy (Zhu et al., 2015; Trogden et al., 2021).
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Trogden, K.P., Lee, J., Bracey, K.M., Ho, K.H., McKinney, H., Zhu, X., Arpag, G., Folland, T.G., Osipovich, A.B., Magnuson, M.A., Zanic, M., Gu, G., Holmes, W.R., Kaverina, I. (2021). Microtubules regulate pancreatic β-cell heterogeneity via spatiotemporal control of insulin secretion hot spots. eLife (Figure 2).
Zhu, X., Hu, R., Brissova, M., Stein, R.W., Powers, A.C., Gu, G., Kaverina, I. (2015). Microtubules Negatively Regulate Insulin Secretion in Pancreatic β Cells. Dev. Cell. (Figure 2, panel C–F).
Notes
A 10 μL tip may injure large islets during picking and transferring. We usually can obtain 200–300 islets from one mouse using this procedure. Do not use cell culture dish to incubate isolated islets to prevent their attachment to the dish.
The isolated islets are incubated in 3 mL of islet media in a 60 mm Petri dish overnight for recovery. The islet number should not exceed 100 per dish to minimize the formation of hypoxia core (necrosis due to insufficient O2) in larger islets.
The 10 mm microwell reduces the search time for an attached islet on a TIRF microscope and provides enough room for islets to attach to the glass coverslip instead of the edge of the microwell.
By choosing two islets with clear size difference, e.g., a mid-sized islet with a diameter of 100–120 μm and a small islet with a diameter of 60–80 μm, these two islets can be distinguished under the microscope without diamond pen marking on the coverslip.
Ideally, each 150 mm dish accommodates no more than six MatTek dishes to reduce the chance of knocking nearby MatTek dishes over while retrieving one.
Acknowledgments
This work is supported by National Institutes of Health, grants R01-DK106228 (to IK and GG), R35-GM127098 (to IK), and P30-DK020593-44S1 (to KHH). We thank Margret Fye for constructive feedback on the manuscript.
This protocol was derived from the original work of Trogden et al. (2021).
Competing interests
The authors declare no competing interests.
References
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4,869 | https://bio-protocol.org/en/bpdetail?id=4869&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Purification of Long Non-coding RNAs on Replication Forks Using iROND (Isolate RNAs on Nascent DNA)
WZ Weidao Zhang
MT Min Tang
LW Lin Wang
PZ Ping Zheng
BZ Bo Zhao
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4869 Views: 406
Reviewed by: Emilia KrypotouThirupugal GovindarajanZheng Zachory WeiDr. Amit K. Tripathi
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Original Research Article:
The authors used this protocol in Science Advances Jan 2023
Abstract
Fork stability is key to genome DNA duplication and genetic integrity. Long non-coding RNAs (LncRNAs) may play vital roles in fork stabilization and chromatin remodeling. Existing techniques such as NCC-RNA sequencing are useful to identify LncRNAs on nascent chromatin DNA. However, there is still a lack of methods for LncRNAs purification directly from replicative forks, hindering a deep understanding of the functions of LncRNAs in fork regulation. Here, we provide a step-by-step protocol named iROND (isolate RNAs on nascent DNA). iROND was developed and modified from iPOND, a well-known method for purifying fork-associated proteins. iROND relies on click chemistry reaction of 5'-ethynyl-2'-deoxyuridine (EdU)-labeled forks and biotin. After streptavidin pull down, fork-associated LncRNAs and proteins are purified simultaneously. iROND is compatible with downstream RNA sequencing, qPCR confirmation, and immunoblotting. Integrated with functional methods such as RNA fluorescent in situ hybridization (RNA FISH) and DNA fiber assay, it is feasible to screen fork-binding LncRNAs in defined cell lines and explore their functions. In summary, we provide a purification pipeline of fork-associated LncRNAs. iROND is also useful for studying other types of fork-associated non-coding RNAs.
Key features
• Purify long non-coding RNAs (LncRNAs) directly from replication forks.
• Connects to RNA sequencing for screening easily.
• Allows testing various genotoxic stress responses.
• Provides LncRNA candidate list for downstream functional research.
Graphical overview
Schematic overview of isolate RNAs on nascent DNA (iROND) protocol. Cells were pulse-labeled with 5'-ethynyl-2'-deoxyuridine (EdU) for 10 min before paraformaldehyde fixation. EdU-positive forks were ligated with biotin through Click-IT chemistry reaction. Genomic DNA was ultrasonically cracked and crosslinked with streptavidin for pulling down. Both RNA and protein components were purified. RNA components were used for downstream RNA sequencing and qPCR validation. Protein components were used for immunoblotting to evaluate binding dynamics of fork-associated proteins such as helicase, topoisomerase, and DNA polymerases.
Keywords: LncRNA Purification Replication fork Genotoxic stress Genomic stability
Background
DNA replication and DNA repair are central topics in genome stability research. Currently, a vast majority of studies focuses on protein components intensively. Through the investigation of core factors such as ATM, ATR, CHK1, CHK2, DNA-PK, and P53, as well as their associated signaling pathways, the fundamental regulatory mechanisms in DNA replication and DNA repair have been revealed (Jackson and Bartek, 2009). For example, using isolate proteins on nascent DNA (iPOND) technology, numerous replication fork-associated protein components have been identified, including DNA polymerases, helicases, and epigenetic modifiers (Sirbu et al., 2012). The initiation, elongation, and termination of steady-state replication forks are strictly controlled to ensure the integrity of the genome replication process (Dungrawala and Cortez, 2015). In order to cope with genotoxic drug-induced fork collapse or stalling, endogenous DNA replication stress response mechanisms can ensure DNA replication progression and reduce the accumulation of DNA breaks (Berti et al., 2020). In terms of DNA repair, numerous specific techniques have been developed. For instance, GFP-based reporter systems were used to evaluate DNA repairing efficiency (Pierce et al., 1999). Single-cell comet assay is used broadly to examine general DNA damage levels (Collins, 2004). Overall, the involvement of protein components in genome stability maintenance is presently well understood.
Besides protein components, a few recent studies raised the potential importance of RNA components in DNA repair and replication, facilitated by indirect identification techniques. For example, the NCC-RNA-seq technique, based on the principle of biotin affinity purification, has been used to identify multiple long and short RNA molecules associated with nascent chromosomes (Gylling et al., 2020). The functions of these RNAs in chromosome reshaping and maturation warrant further investigation. Additionally, using RNA affinity purification coupled with mass spectrometry, the long non-coding RNA (LncRNA) NORAD plays a critical role in genome stability through interaction with PUMILIO protein (Elguindy and Mendell, 2021). Comparative transcriptomics is an effective approach for discovering functional non-coding RNAs. In our previous studies, we identified a novel LncRNA, DISCN, by comparing the expression profiles of LncRNAs in mouse embryonic stem cells (mESCs) with differentiated cells. DISCN exhibits specific high expression in embryonic and neural stem cells. DISCN forms a functional complex with the nucleolar protein nucleolin and the DNA single-strand binding protein RPA, through which it keeps an RPA protein pool to ensure the efficiency of DNA replication stress response and DNA repair (Wang et al., 2021). These studies support the notion that non-coding RNAs form a new regulatory layer in genomic stability system. However, limited by direct purification techniques, the roles of non-coding RNAs in DNA replication or DNA repair are still not understood.
We have recently developed a novel method called iROND (isolate RNAs on nascent DNA). Using iROND, we identified a specific fork-binding LncRNA Lnc956, in the embryonic stem cell system. Lnc956 forms a stable nucleic acid–protein complex with TRIM28 and HSP90B1, facilitating the integration of the CMG helicase at replication fork sites and ensuring replication fork stability in mESCs (Zhang et al., 2023). Here, we provide a step-by-step protocol for iROND. iROND is applicable in different types of rapidly proliferating cell lines and compatible with downstream functional validation systems such as RNA fluorescence in situ hybridization (RNA-FISH) and DNA fiber assays.
Materials and reagents
Biological materials
Mouse embryonic stem cells (derived from C57Bl/6J blastocysts)
Mouse NIH3T3 cells (kindly provided by Dr. Bingyu Mao at Kunming Institute of Zoology, Chinese Academy of Sciences)
Materials
35 mm cell culture dish (Corning, catalog number: 430165)
60 mm cell culture dish (NEST, catalog number: 705001)
100 mm culture dish (NEST, catalog number: 704001)
150 mm culture dish (Corning, catalog number: 430599)
200 μL PCR tube (Axygen, catalog number: AXYPCR02LC)
1.5 mL centrifugal tube (Axygen, catalog number: AXYMCT150C)
15 mL centrifugal tube (NEST, catalog number: 601052)
50 mL centrifugal tube (NEST, catalog number: 602002)
0.22 μm filter (Millipore, catalog number: SLGP033RB)
Reagents
5-ethynyl-2′-deoxyuridine (EdU) (Life Technology, catalog number: A10044)
Thymidine (Sigma-Aldrich, catalog number: T1895)
37% (w/v) formaldehyde solution (Sigma-Aldrich, catalog number: F1635); caution: toxic
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P3911)
Potassium phosphate monobasic (KH2PO4) (Sigma-Aldrich, catalog number: P5655)
Sodium phosphate dibasic (Na2HPO4) (Sigma-Aldrich, catalog number: S5136)
Glycine (Sigma-Aldrich, catalog number: G7126)
Triton X-100 (Sigma-Aldrich, catalog number: X100)
BSA (Sigma-Aldrich, catalog number: V900933)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D8418)
Copper (II) sulfate pentahydrate (CuSO4·5H2O) (Sigma-Aldrich, catalog number: 209198)
(+) Sodium L-ascorbate (Sigma-Aldrich, catalog number: A4034)
Biotin azide (Invitrogen, catalog number: B10184)
SDS (Sangon Biotech, catalog number: A600485)
Tris (Sangon Biotech, catalog number: A610195)
Glycerol (Sangon Biotech, catalog number: A600232)
Bromophenol blue (Sangon Biotech, catalog number: A602230)
EDTA (Sigma-Aldrich, catalog number: E9884)
Agarose (Merck, catalog number: A4718)
Dithiothreitol (DTT) (Merck, catalog number: D8255)
Aprotinin (Merck, catalog number: A6279)
Leupeptin (Merck, catalog number: L2884)
Streptavidin agarose (Novagen, catalog number: 69203-3)
RNase inhibitor (Thermo Fisher, catalog number: EO0381)
Diethyl pyrocarbonate (DEPC) (Sigma-Aldrich, catalog number: D5758)
Protease K (Thermo Fisher, catalog number: 4333793)
Chloroform (Sigma-Aldrich, catalog number: 366911)
Isopropyl alcohol (Sigma-Aldrich, catalog number: W292912)
Ammonium acetate (Sigma-Aldrich, catalog number: A7262)
Alcohol (Sigma-Aldrich, catalog number: AX0442)
Acetic acid (Sigma-Aldrich, catalog number: 695092)
TRIzol (Thermo Fisher, catalog number: 15596026)
Hydroxyurea (HU) (Sigma-Aldrich, catalog number: H8627)
Solutions
EdU solution (see Recipes)
DEPC-H2O (see Recipes)
Thymidine solution (see Recipes)
Chase medium (see Recipes)
1× PBS solution (see Recipes)
1% Formaldehyde solution (see Recipes)
1.25 M Glycine solution (see Recipes)
Permeabilization buffer (see Recipes)
Wash buffer (see Recipes)
Biotin-azide solution (see Recipes)
100 mM CuSO4 solution (see Recipes)
Sodium L-ascorbate solution (see Recipes)
Lysis buffer (pH 8.0) (see Recipes)
Salt wash buffer (see Recipes)
2× SDS Laemmli sample buffer (2× SB) (see Recipes)
50× Tris acetate-EDTA (TAE) buffer (see Recipes)
Recipes
EdU solution
Reagent (use for storing) Final concentration Quantity
EdU (50 mg) 10 mM 25.2 mg
DMSO n/a 10 mL
Total n/a 10 mL
Reagent (use for cell labeling) Final concentration Quantity
EdU (10 mM) 10 μM 1 μL
mESCs medium (see Step A1) n/a 999 μL
Total n/a 1 mL
DEPC-H2O
Reagent Final concentration Quantity
DEPC n/a 1 mL
H2O n/a 999 mL
Total n/a 1,000 mL
Thymidine solution
Reagent (use for storing) Final concentration Quantity
Thymidine (1 g) 10 mM 24.2 mg
DEPC-H2O n/a 10 mL
Total n/a 10 mL
Chase medium
Reagent (use for cell labeling) Final concentration Quantity
Thymidine (10 mM) 10 μM 1 μL
mESCs Medium n/a 999 μL
Total n/a 1 mL
1× PBS solution
Reagent Final concentration Quantity
NaCl 137 mM 8 g
KCl 3 mM 0.2 g
Na2HPO4 8 mM 1.15 g
KH2PO4 2 mM 0.24 g
DEPC-H2O n/a 1 L
Total n/a 1 L
1% Formaldehyde solution
Reagent Final concentration Quantity
Formaldehyde (37%) 1% 0.27 mL
DEPC-PBS n/a 9.73 mL
Total n/a 10 mL
1.25 M Glycine solution
Reagen Final concentration Quantity
Glycine 1.25 M 46.92 g
DEPC-H2O n/a 500 mL
Total n/a 500 mL
Permeabilization buffer
Reagent Final concentration Quantity
Triton X-100 0.25% 2.5 mL
DEPC-PBS n/a 997.5 mL
Total n/a 1 L
Wash buffer
Reagent Final concentration Quantity
BSA 0.5% 0.5 g
DEPC-PBS n/a 100 mL
Total n/a 100 mL
Biotin-azide solution
Reagent Final concentration Quantity
Biotin-azide 1 mM 1 mg
DMSO n/a 1.624 mL
Total n/a 1.624 mL
100 mM CuSO4 solution
Reagent Final concentration Quantity
CuSO4·5H2O 100 mM 1.248 g
DEPC-H2O n/a 50 mL
Total n/a 50 mL
Sodium L-ascorbate solution
Reagent Final concentration Quantity
(+) sodium L-ascorbate 100 mM 20 mg
DEPC-H2O n/a 1 mL
Total n/a 1 mL
Lysis buffer (pH 8.0)
Reagen Final concentration Quantity
Tris 10 mM 1.21 g
SDS 7 mM 2 g
DEPC-H2O n/a 200 mL
RNase inhibitor 10 U/μL 200 μg
Aprotinin 1 μg/mL 200 μg
Leupeptin 1 μg/mL 200 μg
Total n/a 200 mL
Salt wash buffer
Reagent Final concentration Quantity
NaCl 1 M 14.625 g
DEPC-H2O n/a n/a
Total n/a 250 mL
2× SDS Laemmli sample buffer (2× SB)
Reagent Final concentration Quantity
SDS 123 mM 0.4 g
Bromophenol blue 1 mM 0.01 g
Glycerol 18% 2 mL
1 M Tris (pH 6.8)
1M DTT
1 mM
1 mM
1.25 mL
1.25 mL
H2O n/a 8 mL
Total n/a 12.5 mL
50× Tris acetate-EDTA (TAE) buffer
Reagent Final concentration Quantity
Tris 2 M 242 g
Acetic acid n/a 57.1 mL
EDTA 0.1 M 37.2 g
H2O n/a 942.9 mL
Total n/a 1,000 mL
Equipment
Thermo Forma series II water jacketed CO2 incubator (Thermo Scientific, model: 3111)
Nikon eclipse Ti inverted microscope (Nikon, model: Ti)
Ultrasonic cell disruptor (Diagenode, Bioruptor Plus)
90 µm nylon mesh (Small Parts Inc., catalog number: B000FN0PGQ)
Roll shaker for incubation (Thermo Scientific, model: PHMT)
Real-time fluorescence quantitative PCR instrument (Bio-Rad, model: CFX96Touch)
Vortexer (VWR analog vortex mixer, catalog number: 10153-838)
4 °C refrigerator (Haier, model: HYC-1090)
-20 °C freezer (Haier, model: DWL262)
NanoDrop (Thermo Scientific, ND2000)
-80 °C freezer (Thermo Scientific, model: TSX60086VRAKLB)
Software
Microsoft Excel (Microsoft, https://products.office.com/en-us/excel) (Office Excel 2016, September 22, 2015)
GraphPad Prism (GraphPad, https://www.graphpad.com/scientific-software/prism/) (GraphPad Prism 8.3.0, October 29, 2019)
Procedure
Label cells with EdU
For the preparation of mESC medium and culture and expansion of mESCs and NIH3T3, please refer to our publication (Zhang et al., 2023). Before EdU labeling, incubate cells with 37 °C pre-warmed fresh mESCs medium for 2 h, with 25 mL of medium for each 15 cm diameter dish containing ~1.5 × 107 cells.
To perform EdU labeling, take the dishes to a biological safety cabinet, add 25 μL of EdU stock (10 mM) into 25 mL of pre-existing medium for each dish, and mix well gently to make sure that the final concentration of EdU is 10 μM. Depending on the percentage of mESCs in S phase (more than 60%), we calculated that at least 9 × 109 cells in each dish were labeled by EdU.
Put the dishes back into the incubator quickly and label for 10 min.
Note: Typically, fork-associated RNA components display a low-dose manner. To improve the productivity, using a cell line with high percentage of S-phase cells is critical to secure the RNA output. In our study, we used mESCs, as these always keep more than 60% of cells in S phase. If you need to evaluate replication stress dynamics, add genotoxic drugs such as 4 mM hydroxyurea (HU) into the dish and incubate for 2 h.
To get post-replicated chromatin samples, remove EdU-positive medium, carefully wash cells three times with 5 mL each time of pre-warmed chase medium. Chase medium, containing 10 μM thymidine, is used to compete with EdU to chase DNA replication. After 1 h incubation, the EdU-labeled DNA has been constructed to mature chromatin. So, this group is used as a chromatin control. Then, add 25 mL of pre-warmed chase medium and incubate for 1 h.
Formaldehyde crosslink and harvest cells
Once finished with EdU labeling, thymidine chase, or drugs treatment, remove culture medium quickly and add 10 mL of ice-cold PBS containing 1% formaldehyde to each dish to fix cells for 20 min at room temperature.
Caution: Formaldehyde is toxic to human health.
Critical: Do not wash cells with PBS buffer before fixation since PBS washing may cause RNA degradation.
Add 1 mL of 1.25 M glycine to each dish to quench formaldehyde.
Scrape cells and harvest into 50 mL RNase-free centrifuge tubes. Record the fixation volume.
Critical: Keep tubes on ice all the time.
Spin down at 900× g for 5 min at 4 °C.
Remove supernatant.
Use ice-cold 1× PBS buffer containing 10 U/μL RNase inhibitor to wash sediments three times with the same volume as the fixation volume from step B3.
Pause point: Samples can be quickly immersed into liquid nitrogen and stored at -80 °C for at least four weeks.
Cell permeabilization
Resuspend cells in ice-cold permeabilization buffer containing 10 U/μL RNase inhibitor. Adjust cell density to 1 × 107 cells/mL. Vortex well and incubate at room temperature for 30 min on a shaker (40 rpm).
Centrifuge at 900× g for 5 min at 4 °C.
Remove supernatant.
Wash cells with ice-cold 1× PBS containing 0.5% BSA and 10 U/μL RNase inhibitor; the volume used is equal to the permeabilization buffer in step C1.
Centrifuge at 900× g for 5 min at 4 °C and remove supernatant.
Repeat steps C4 and C5 once. Keep samples on ice for click reaction.
Click reaction
Prepare click reaction cocktail as listed in Table 1.
Table 1. Click reaction cocktail (5 mL for 1 × 108 cells)
Reagent Stock concentration Working concentration Control reaction volume (mL) Experimental reaction volume (mL)
1× PBS 4.225 4.225
Biotin-azide 1 mM 10 μM 0.05
CuSO4 100 mM 2 mM 0.1 0.1
(+) Sodium L-ascorbate 100 mM 10 mM 0.5 0.5
DMSO 0.05
RNase inhibitor 40 U/μL 1 U/μL 0.125 0.125
Total volume 5.0 5.0
Critical: The biotin-azide is sensitive to light. The reaction cocktail should be stored in the dark.
Critical: CuII is unstable in water. The reaction cocktail should be prepared fresh.
Resuspend cells into click reaction cocktail. Adjust reaction volume to reach the density of 1 × 108 cells per 5 mL.
Vortex gently and incubate for 2 h at room temperature on a shaker (40 rpm).
Critical: Keep the reaction tubes in the dark.
Centrifuge at 900× g for 5 min at 4 °C and remove supernatant.
Wash cells with ice-cold 1× PBS buffer containing 0.5% BSA and 10 U/μL RNase inhibitor. The PBS volume used is equal to the permeabilization buffer in step C1.
Centrifuge at 900× g for 5 min at 4 °C and remove supernatant.
Repeat steps D5 and D6 once.
Cell lysis and sonication
Keep lysis buffer on ice. Add leupeptin, aprotinin, and RNase inhibitor before use.
Add 100 μL of lysis buffer per 1.5 × 107 cells and resuspend well. Transfer cell suspension into a 1.5 mL RNase-free centrifuge tube.
Sonicate cells using a microtip sonicator with the following settings: pulse, 20 s; 40 s pause; 15 cycles; power: 13–16 watts.
Critical: Tubes should be immersed into an ice-cold water bath in the process of sonication to prevent overheating.
Note: After sonication, the supernatant should be clear, but not cloudy.
Centrifuge at 12,000× g for 15 min at 4 °C.
Transfer the supernatant through a 90 µm nylon mesh and collect into a new 1.5 mL RNase-free centrifuge tube; then, keep samples on ice.
Note: To examine DNA fragment size at this step, pick up 5 μL of supernatant to extract DNA. Perform electrophoresis using 1% agarose gel at 120 V for 30 min to detect DNA fragments (Figure 1).
Figure 1. DNA fragments after sonication. The size of DNA fragments is mainly concentrated in the 200–1,000 base pair range.
DNA extraction.
Take 5 μL from the supernatant from step E5, add protease K to the final concentration of 50–100 μg/mL, mix well, and incubate at 50 °C for 3 h.
Add an equal volume of chloroform/isopropyl alcohol (volume ratio: 24:1), mix well, centrifuge at 10,000× g for 10 min, and transfer the supernatant to the new centrifuge tube.
Add 10% volume of 10 M ammonium acetate and mix gently.
Add twice the volume of isopropyl alcohol and mix gently.
Centrifuge at 10,000× g for 10 min and discard supernatant.
Add the same volume of 75% ethanol and mix well, centrifuge at 10,000× g for 5 min, and discard the supernatant.
Air-dry pellets for 5 min and dissolve with 10 μL of double-distilled water for electrophoresis.
Dilute the samples with the same volume of ice-cold PBS to improve the efficiency of following Biotin capture. Make sure to add 1 μg/mL leupeptin, 1 μg/mL aprotinin, and 10 U/μL RNase inhibitor.
Pick up 15 μL of each sample, mix well with 15 μL of 2× SB, and boil at 95 °C for 10 min. These samples can be used as input for immunoblotting targeting replication fork proteins such as PCNA.
Critical: This step is crucial since immunoblotting of PCNA can be used to confirm the pull-down efficiency and specificity.
Pick up 15 μL of each sample, mix well with 500 μL of TRIzol, and store at -80 °C for RNA purification.
Critical: This step is crucial; samples will be used to calculate enrichment fold change in the following qPCR confirmation.
Streptavidin capture
Vortex streptavidin agarose beads well.
Calculate agarose beads volume for each sample. Typically, 100 μL of streptavidin agarose beads is used per 1 × 108 cells.
Wash streptavidin agarose beads two times with lysis buffer and one time with PBS before use, using 1 mL for each wash.
Add beads into each sample and incubate for 16 h at 4 °C on a rotator (40 rpm).
Critical: Keep the rotator in the dark.
Centrifuge at 1,800× g for 1 min at 4 °C and remove supernatant.
Wash beads three times with ice-cold lysis buffer and one time with 1 mL of salt wash buffer. For each wash, samples should be rotated for 5 min before centrifugation.
Elution of proteins and RNAs
Separate the beads of each sample into two parts equally to elution protein and RNA components, respectively.
Add 2 μL of 2× SB into one part, mix well, and boil samples at 95 °C for 25 min to elute proteins. These are ready to use for immunoblotting.
For elution of RNAs, add 500 μL of TRIzol into another part of each sample, and mix well.
Add 100 μL of chloroform into each tube, vortex tightly for 30 s, and leave on ice for 5 min.
Centrifuge at 12,000× g for 10 min at 4 °C.
Transfer the supernatant of each sample into a new RNase-free tube and record the volume.
Add an equal volume of isopropanol and leave on ice for 10 min.
Centrifuge at 12,000× g for 10 min at 4 °C. Remove the supernatant.
Wash the pellet with 75% alcohol.
Centrifuge at 12,000× g for 10 min at 4 °C. Remove the supernatant and air dry for 1 min.
Add 40 μL of RNase-free double-distilled water to dissolve the pellet.
Measure RNA concentration using a NanoDrop. Approximately 200–400 ng of RNA can be extracted from 1× 108 cells.
Note: RNA purification is critically important; the optical density (OD) 260/230 value should be between 1.5 and 2.4, and the OD260/280 value must be between 1.8 and 2.4.
Data analysis
Immunoblotting is useful to confirm the pull-down specification of fork DNA fragments. PCNA and Histone H3 can be used as a fork-binding protein marker and mature chromatin marker, respectively. Please refer to Supplementary Figure 1A in our publication (Zhang et al., 2023).
RNA samples can be used either for qPCR analysis or RNA sequencing (Figures 2 and 3). Please refer to Figure 1B, C, and D in our publication (Zhang et al., 2023).
Validation of protocol
Figure 2. Design of isolate RNAs on nascent DNA (iROND)-based LncRNA screening in mouse embryonic stem cells (mESCs) and NIH3T3 cells (Zhang et al., 2023). We set up the following experimental items: 1) no click group: cells were pulse-labeled by EdU but click reaction was omitted, as a non-specific binding control. 2) Click-normal group: cells were cultured in normal conditions and standard iROND was performed, as a steady-state sample. 3) Click-HU group: cells were pulse-labeled by EdU and treated with 4 mM hydroxyurea (HU) to induce replication stress, as a genotoxic sample. 4) Click-Protease group: all procedures were the same as group 3, with extra treatment of 100 μg/mL protease K for 30 min to degrade protein components. 5) Click-chase group: cells were chase-labeled with thymidine to get a mature chromatin control.
Figure 3. RNA sequencing and bioinformatic comparison of isolate RNAs on nascent DNA (iROND) samples from mouse embryonic stem cells (mESCs) and NIH3T3 cells under indicated conditions (Zhang et al., 2023). Two biological repeats were performed for each group. Notably, a set of LncRNAs were significantly enriched in mESC Click-HU group.
General notes and troubleshooting
Compared with published methods for studying LncRNAs in genomic dynamics or stability, iROND is more specific for purifying the fork-associated LncRNA subpopulation (Gylling et al., 2020; Wang et al., 2021). One limitation is that iROND needs billions of cells for each assay. The more S-phase cells, the easier performing the assay is. In our study, we used mESCs and NIH3T3 cells; both proliferate very fast. This is critical for iROND success. We assume that it might be difficult if low proliferating cell lines are used. Alternatively, synchronization into S phase before EdU labeling may be a reasonable choice.
In the steps of EdU labeling and thymidine chase, pre-warming medium is important to minimize disturbance to cells in a very short time.
In order to label replication forks as specifically as possible, EdU labeling must be done in 10 min. It is quite hard for only one person to handle many culture dishes in a very short period of time. So, cooperation of two or three people is recommended.
Acknowledgments
The work was supported by National Natural Science Foundation of China (31930027 to P.Z. and 32000422 to W.Z.), National Key Research and Developmental Program of China (2021YFA1102002), Yunnan Fundamental Research Projects (202001AT070140 and 2019FB049). The protocol is derived from the assay systems described in Zhang et al. (2023).
Competing interests
The authors have no competing interests.
References
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4,870 | https://bio-protocol.org/en/bpdetail?id=4870&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Generation of Human Blood Vessel and Vascularized Cerebral Organoids
XS Xin-Yao Sun §
XJ Xiang-Chun Ju *
HZ Hong-Fang Zhao
ZY Zhi-Wen You
RH Run-Run Han
ZL Zhen-Ge Luo
(*contributed equally to this work, § Technical contact: [email protected])
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4870 Views: 1660
Reviewed by: Philipp WörsdörferAntoine de Morree Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE May 2022
Abstract
Brain organoids have been widely used to study diseases and the development of the nervous system. Many reports have investigated the application of brain organoids, but most of these models lack vascular structures, which play essential roles in brain development and neurological diseases. The brain and blood vessels originate from two different germ layers, making it difficult to induce vascularized brain organoids in vitro. We developed this protocol to generate brain-specific blood vessel and cerebral organoids and then fused them at a specific developmental time point. The fused cerebral organoids exhibited robust vascular network-like structures, which allows simulating the in vivo developmental processes of the brain for further applications in various neurological diseases.
Key Features
• Culturing vascularized brain organoids using human embryonic stem cells (hESCs).
• The new approach generates not only neural cells and vessel-like networks but also brain-resident microglia immune cells in a single organoid.
Graphical overview
Workflow and timeline for vessel organoid and vascularized brain organoid generation. (By Figdraw, ID: RTIURffccf)
Keywords: Blood vessel organoids Cerebral organoids Neural organoids Vascularized brain organoids Human embryonic stem cells
Background
As stem-cell-derived 3D-microtissue, brain organoids have been developed to model developmental programs of the human fetal brain and recapitulate developmental, psychiatric, and degenerative brain diseases. The wide application of brain organoids improves our understanding of the developmental process of the human brain (Lancaster and Knoblich, 2014; Di Lullo and Kriegstein, 2017; Amin and Paşca, 2018; Benito-Kwiecinski, 2021). However, lack of a neurovascular system, which plays essential roles in oxygen supply, neurogenesis regulation, and functional application, limits the applications of brain organoids (Tata et al., 2016).
Blood vessels originate from the mesoderm and develop via sequential vasculogenesis and angiogenesis processes, which invade the neuroepithelial regions and form the brain vascular network (Lee et al., 2009). A self-organizing human blood vessel organoid model has been reported and applied to the study of diabetic vasculopathy (Wimmer et al., 2019). However, because the blood vessels and neurons in the brain are derived from two different germ layers, it has been challenging to generate vascularized brain organoids. Several studies have attempted to induce vascular structures in cerebral brain organoids, including co-cultured brain organoids with endothelial cells (ECs) or their progenitors (Mansour et al., 2018; Pham et al., 2018; Cakir et al., 2019; Shi et al., 2020; Wörsdörfer et al., 2020). These reports provide various ideas for further improvement in this field.
We recently developed an induction approach for brain-specific vascular organoids cultured in the medium containing neurotrophic factors at the maturation stage to obtain vessel organoids with cerebrovascular characteristics (Sun et al., 2022). Although the generation of vascularized neural organoids by assembling neural and mesenchymal aggregates has been reported (Wörsdörfer et al., 2020), we showed more abundant cell types in the vessel organoids, including a substantial number of microglia cells in the fusion vascularized organoids, which enabled better recapitulation of the brain characteristics in vivo. This vessel organoid model showed a variety of cell types similar to those observed in vivo. The mature vessel organoids exhibited robust vasculatures that can invade the brain organoids when wrapped in Matrigel. This approach also induced microglial cells along with other types of vascular cells. The fusion vascularized brain organoids induced by this protocol accurately simulate the structure and function of brain vessels. This protocol provides a platform for studying interactions between neuronal and non-neuronal components during brain development and function.
Materials and reagents
Biological materials
H9 embryonic stem cell (H9-ESC, human) (WiCell, WA09)
H9-EGFP (generated in house)
Reagents
mTeSR1 (STEMCELL Technologies, catalog number: 85850)
ReLeSRTM (STEMCELL Technologies, catalog number: 05872)
Accutase (STEMCELL Technologies, catalog number: 07920)
Y27632 (STEMCELL Technologies, catalog number: 72304)
Human Recombinant bFGF (STEMCELL Technologies, catalog number: 78003)
Human Recombinant VEGF (STEMCELL Technologies, catalog number: 78073)
Human Recombinant BMP4 (R&D, catalog number: 314-BP-050)
DMEM/F12 (Life/Invitrogen, catalog number: 11330032)
Knockout serum replacement (KSR) (Gibco, catalog number: 10828028)
MEM-NEAA (Gibco, catalog number: 11140050)
Glutamax (Gibco, catalog number: 35050061)
β-mercaptoethanol (Sigma-Aldrich, catalog number: M3148)
Dorsomorphine (Tocris, 3093/10)
A83-01 (Tocris, catalog number: 2939/10)
Neurobasal (Life/Invitrogen, catalog number: 21103049)
N2 supplement (Life/Invitrogen, catalog number: 17502048)
B27 supplement (Life/Invitrogen, catalog number: 17504044)
B27 supplement without vitamin A (Life/Invitrogen, catalog number: 12587010)
Insulin (Sigma-Aldrich, catalog number: I9278)
Heparin (Sigma-Aldrich, catalog number: H3393)
Lipidure®-CM5206 (NOF CORPORATION, catalog number: CM5206)
Antibiotic-Antimycotic (Gibco, catalog number: 15240096)
Matrigel hESC-qualified matrix (BD-Biocoat, catalog number: 354277)
Matrigel growth factor reduced (BD-Biocoat, catalog number: 354230)
STEMdiff APEL2 medium (STEMCELL Technologies, catalog number: 05270)
Endothelial cell growth medium MV2 (Promocell, catalog number: C-22022)
CHIR99021 (Selleck, catalog number: S1263)
LDN-193189 2HCl (Selleck, catalog number: S7505)
SB431542 (Selleck, catalog number: S1067)
DPBS (Life/Invitrogen, catalog number: 14190144)
Mouse anti-CD31 (Abcam, catalog number: ab9498, 1:500)
Goat anti-DCX (Santa Cruz, catalog number: sc-8066, 1:200)
Chicken anti-GFP (Aves Lab, SKU number: GFP-1020, 1:1,000)
Rabbit anti-IBA1 (Wako, catalog number: 019-19741, 1:500)
DAPI fluorochrome (Beyotime, catalog number: C1002)
Paraformaldehyde (PFA) (Sigma-Aldrich, catalog number: 158127)
Triton X-100 (Sigma-Aldrich, catalog number: T8787)
Bovine serum albumin (Sigma-Aldrich, catalog number: A9418)
Sucrose (Sinopharm, catalog number: 10021418)
Antigen retrieval buffer (10 mM citric acid, pH 6.0) (Sigma-Aldrich, catalog number: 1159047)
Optimal cutting temperature (OCT) compound (Sakura, catalog number: 4583)
Aggregation medium (see Recipes)
Mesoderm induction medium (see Recipes)
Endothelial induction medium (see Recipes)
Endothelial maturation medium (see Recipes)
Expansion medium (see Recipes)
Maturation medium (see Recipes)
Ectoderm induction medium (see Recipes)
Neuroepithelium induction medium (see Recipes)
Recipes
Note: The basal medium could be stored at -20 °C for six months and thawed immediately before use. However, as presented in the following recipes, all culture media can be stored for up to two weeks at 4 °C. The small molecules and growth factors (Y27632, CHIR99021, VEGF, BMP4, bFGF, dorsomorphine, A83-01, LDN193189, and SB431542) should be added immediately before use.
Aggregation medium
Reagents Final concentration Volume
mTeSR1 50 mL
Y27632 10 μM 50 μL
Mesoderm induction medium
Reagents Final concentration Volume
APEL2 10 mL
CHIR99021 6 μM 12 μL
Endothelial induction medium
Reagents Final concentration Volume
APEL2 20 mL
VEGF 50 ng/mL 20 μL
BMP4 25 ng/mL 10 μL
bFGF 10 ng/mL 4 μL
Endothelial maturation medium
Reagents Final concentration Volume
MV2 50 mL
VEGF 50 ng/mL 50 μL
Expansion medium
Reagents Final concentration Volume
DMEM/F12 500 mL
Neurobasal 500 mL
N2 supplement 5 mL
B27 supplement without vitamin A 10 mL
β-mercaptoethanol 3.5 μL
Insulin 250 μL
Glutamax 10 mL
MEM-NEAA 5 mL
Antibiotic-Antimycotic 10 mL
VEGF 20 ng/mL
Maturation medium
Reagents Final concentration Volume
DMEM/F12 500 mL
Neurobasal 500 mL
N2 supplement 5 mL
B27 supplement 10 mL
β-mercaptoethanol 3.5 μL
Insulin 250 μL
Glutamax 10 mL
MEM-NEAA 5 mL
Antibiotic-Antimycotic 10 mL
VEGF 20 ng/mL
Note: The human recombinant VEGF is added immediately before use in the expansion and maturation mediums, with the final concentration of 20 ng/mL.
Ectoderm induction medium
Reagents Final concentration Volume
DMEM/F12 400 mL
KSR 100 mL
β-mercaptoethanol 3.5 μL
Glutamax 5 mL
MEM-NEAA 5 mL
Dorsomorphine 2.5 μM
A83-01 2 μM
Neuroepithelium induction medium
Reagents Final concentration Volume
DMEM/F12 500 mL
N2 supplement 5 mL
Glutamax 5 mL
MEM-NEAA 5 mL
Heparin 1 μg/mL
SB431542 10 μM
LDN193189 2HCl 200 nM
Laboratory supplies
Cell culture 6-well plates (Corning, catalog number: 3516)
V-bottom 96-well plates (Thermo, catalog number: 277143)
Cell culture 60 mm dishes (Corning, catalog number: 430166)
Cell culture 35 mm dishes (Corning, catalog number: 430165)
Pasteur pipettes, 3 mL (Nest, catalog number: 318314)
Serological pipette, 5 mL (Corning, catalog number: 4487)
Serological pipette, 10 mL (Corning, catalog number: 4492)
Serological pipette, 25 mL (Corning, catalog number: 4251)
Sterile tips (Axygen)
Sterile 15 mL polypropylene centrifuge tubes (Corning, catalog number: 430790)
Sterile 50 mL polypropylene centrifuge tubes (Corning, catalog number: 430828)
Equipment
Olympus FV3000 confocal laser scanning microscope
Olympus CKX53 microscope
Water bath (Jinghong, model: XMTD-8222)
Biosafety cabinet (Thermo, model: 1300 SERIES A2)
CO2 constant-temperature incubator (Thermo, model: HERAcell 150i)
Pipette sets (Eppendorf)
Table shaker (SCILOGEX SLC-O3000-S)
Centrifuge (Eppendorf, model: 5804R)
Cryostat (Thermo, model: NX50)
Freezers (Panasonic, model: Haier)
Software
Fiji/ImageJ (ImageJ 1.52e, https://fiji.sc, access date 07/11/2018)
OLYMPUS FV31S-SW
Figdraw (https://www.figdraw.com/static/index.html#/, access date 02/14/2022)
Procedure
H9-EGFP and H9-ES cell culture
The identity of the H9 human embryonic stem (H9-hES) cell line was confirmed by short tandem repeat (STR) profiling (performed by APPLIED CELL). The cells were tested for mycoplasma, and the result was negative. The H9 cell line stably expressing H9-EGFP was generated by introducing a CAG-EGFP DNA fragment into the genome locus ROSAβgeo26 (ROSA26) using the clustered regularly interspaced short palindromic repeats-associated protein 9 (CRISPR/Cas9) method.
Note: The experiments published in the original research paper and described here were performed using the H9 cell line purchased from APPLIED CELL. Unfortunately, the company no longer provides this cell line. We suggest directly ordering the H9 cell line (WA09) from WiCell Research Institute (Madison, WI, USA) or using an alternative cell line, such as an iPS cell line from WiCell (UCSD093i-1-11), which we have tested yielding similar results. In addition, mycoplasma testing is crucial, because cells infected with mycoplasma can have their growth and differentiation affected; for the testing protocol, refer to this latest study by Siegl et al. (2023).
Coat 6-well plates using hESC-qualified Matrigel
Thaw the frozen Matrigel stock solution in the fridge (4 °C) overnight and place the vial on ice to avoid gel formation.
To prepare the working solution, dilute Matrigel stock solution by adding cold (4 °C) DMEM/F12 into the vial on ice and keep pipetting until the material is evenly dispersed.
Add 1 mL of Matrigel working solution per well and distribute the solution gently to cover the bottom surface. Then, incubate the Matrigel-coated plate for 1 h in the cell incubator (37 °C).
Note: The best dilution ratio of the Matrigel stock solution is based on the batch number (LOT number); please check on the official Corning website. Before coating, Matrigel stock solution dilution must be kept on ice. Otherwise, it will irreversibly solidify quickly when the highly concentrated Matrigel stock solution is placed at room temperature (RT). If you need to pipette the concentrated Matrigel solution, freeze the tips in the freezer ahead of time to avoid the clogging of Matrigel in the tip. The lot numbers we have tested are 2088003/2095003/2109003/2116003/2158003/2160996/2165002/2249003/2250003/2319003 (e.g., visit the website https://www.corning.com/asean/en/products/life-sciences/resource-library.html?productNumber=354277&lotNumber=, input the lot number 2088003, and click on Download certificate. The “dilution factor: 310 μL” implies that 310 μL of Matrigel is diluted by the mTeSR medium up to 25 mL for the final concentration).
Cell thawing
Add 10 μM of Y27632 (Rock inhibitor) to the embryonic stem cells (ESC) culture medium (mTeSR) to prevent cell death and warm the medium at 37 °C for 30 min (the warming time depends on the volume of the medium).
Remove the cryotube from liquid nitrogen and quickly thaw the cells in a 37 °C water bath with only a small ice pellet remaining.
Gently add 1 mL of warm mTeSR medium in a dropwise manner to the cells to avoid a sudden change in the osmolarity of the freezing solution around the cells and improve recovery; then, transfer the cells into a 15 mL conical tube with fresh mTeSR medium (3 mL per cryovial) and centrifuge at 200× g for 2 min.
Note: A sudden change in the osmolarity of the cryoprotectant around the cells may cause a rapid stream of water across the membranes of the cells, which could stress the cells and make them more prone to dying. Avoiding this stressful operation improves cell survival and recovery.
Remove the supernatant medium and resuspend the cells in fresh mTeSR medium containing rho-associated protein kinase (ROCK) inhibitor (2–3 mL per culture well for the 6-well plate) using a 1,000 μL tip by pipetting 2–3 times as gently as possible.
Aspirate the coating Matrigel solution and transfer the cell suspension into the coated well.
Note: When warming the medium, the use of a water bath with circulating water will shorten the time. The experimenter should check the warmed medium by gripping the bottle with a hand and avoid using the cold medium. When resuspending cells, pipette as few times as possible. Do not use sharp tips (e.g., 200 μL tip) to pipette cells, as this will cause severe cell death. Y27632 is added into the medium only on the first day of cell resuscitation.
Cell maintenance
Aspirate the medium from the 6-well plate that contains human ESC (hESC) colonies and add 2 mL of fresh culture medium (mTeSR with 4 ng/mL bFGF) on the following day when the cells are well attached to the well. Replace the culture medium daily.
Note: The culture medium can be changed every other day if only a few dead cells exist. Check the status of stem cells based on the colony and cell morphology (Wakui et al., 2017), e.g., high nucleus/cytoplasm ratio of cells, flat and well-defined edge of colonies (see also step B1).
Cell passage
Prepare a Matrigel-coated plate as described above before starting the cell passage.
Aspirate the mTeSR medium from the well containing the hESC colonies and wash with 1× DPBS.
Replace the DPBS with 1 mL of ReLeSRTM solution for each well of the 6-well plate and incubate for 30 s. Then, aspirate the ReleSRTM quickly and place the cell plate in the incubator for another 3 min.
Take out the culture plate and gently add the warm mTeSR medium along the wall of the wells.
Gently tap the sides of the culture plate to detach the cell colonies from the bottom of the wells into the fresh medium. Then, dissociate detached colonies into small clumps by gentle pipetting using a 1,000 μL tip.
Seed the cell clumps at a split ratio of approximately 1:10 in a new Matrigel-coated well and return the plate to the incubator. Tilt the plate several times in both the horizontal and vertical directions to evenly spread the cell clumps.
Change the culture medium daily until 80%–90% confluence is achieved for organoid generation.
Note: Use ReleSRTM as the dissociation reagent to detach the cell colonies rather than dissociating the colonies into single cells. Generally, cells should be passaged before the clones adhere together. During cell passage and maintenance, basic fibroblast growth factor (bFGF) should be added into the culture medium right just before use, as the bFGF activity may markedly decline in the medium if added too early.
Blood vessel organoid (VOr) generation (schematic view shown in Figure 1)
Figure 1. Schematic procedure of generating vessel organoids (VOrs) from human embryonic stem cells (hESCs)
Blood vessel organoid aggregation, day 0
Culture H9-EGFP cells in a new 6-well plate as described in section A; the EGFP tag marks the vascular part in the subsequent fusion process. When the cell density reaches approximately 80% confluence and shows good growth status (flat and well-defined edges of colonies and high ratio of nucleus to cytoplasm in cells) (Figure 2A), the stem cells are ready for organoid generation.
Figure 2. Processes of generating VOrs and fVBOrs and representative images. (A) An H9-ESC colony showing a good stemness state. Scale bar, 200 μm. (B) Developmental stages of VOrs from D4 (day 4) to D12 in a V-bottom 96-well plate. Top, bright field; bottom, GFP. Scale bars, 200 μm. (C) Developmental stages of VOrs from D12 to D25 in dishes. Top, bright field; bottom, GFP. Scale bars, 200 μm. Note the protrusion of vessels to the Matrigel. (D) The processes of making dimpled Parafilm substrate for Matrigel embedding. (E) Quantification of the VOr diameter from D4 to D25. Data are presented as mean ± SEM (15–20 organoids at each time point). (F) The failed cases of VOrs at D4, D12, and D16. Scale bars, 200 μm. (G) The developmental stages of fVBOrs from D12 to D40. GFP, VOrs; Bright, BOrs. VOrs: vessel organoids; BOrs: brain organoids; fVBOrs: fusion vascularized brain organoids. Scale bar, 200 μm.
Detach H9-EGFP cells from the Matrigel-coated plate using Accutase to dissociate cell colonies into single cells. Wash the cells with 1× DPBS and add 1 mL of Accutase solution to each well of the 6-well plate. Place cells back in the CO2 incubator at 37 °C for 10 min or until the colonies have been dissociated into single cells and then add 2 mL of mTeSR into each well.
Note: After 5–6 min of digestion, observe the cell states to prevent over-digestion.
Collect the medium into a 15 mL conical tube and centrifuge at 200× g for 2 min.
Cell counting.
Remove the supernatant and resuspend the cells in 1 mL of mTeSR (containing 10 μM Y27632). Pipette cell suspension two to three times to avoid the re-aggregation of dissociated single cells and then dilute cells using the fresh aggregation medium at a reasonable ratio for cell counting. Only count the living cells devoid of trypan blue labeling using a hemocytometer or any cell-counting device in the culture room.
Note: Resuspend the cells evenly before counting, which should be as accurate as possible to reduce the batch-to-batch variation of the initial size of embryoid bodies (EBs). Y27632 is added on the first day of single-cell dissociation and culturing.
Making EBs
i. Use a V-bottom 96-well plate to form the EBs (early stage of the organoids) from single-cell suspension. Prepare 0.5% Lipidure® solution by dissolving 0.25 g of Lipidure® powder in 50 mL of absolute ethyl alcohol in a sterile 50 mL tube. Add 20 μL of Lipidure® solution into each well of the V-bottom 96-well plate for 10 min, pipette off the remaining liquid, and then leave the plate upright with the lid slightly off in the biosafety cabinet to dry completely for 5 min.
Note: 20 μL of Lipidure® solution is enough to cover the V-bottom plate. However, more than 20 μL is also acceptable.
ii. Dilute stem cells using fresh and warm aggregation medium (see Recipes) to a concentration of 7,000–9,000 cells per 150 μL of medium.
iii. Plate 150 μL of cells in each well of the Lipidure®-coated V-bottom 96-well plate. Place it back in the CO2 incubator (37 °C) and culture the cells for 48 h to induce EB formation.
Note: Generally, 7,000 cells per well are used for generating brain organoids and 9,000 cells per well for generating vessel organoids. From experience, 7,000–9,000 cells per well are conducive to generating the two organoids.
Organoid induction, days 2–10
Day 2: Replace the aggregation medium with 150 μL of mesoderm induction medium (see Recipes) for each well and continue to culture EBs in the CO2 incubator at 37 °C for 48 h.
Note: Tilt the plate and aspirate the aggregation medium with a pipette without disrupting the EBs. Do not handle many organoids simultaneously to prevent them from drying out.
Day 4: Replace the mesoderm induction medium with 200 μL of endothelial induction medium (see Recipes) per well and culture EBs at 37 °C for 72 h.
Days 7–11: Replace the endothelial induction medium with 150 μL of endothelial maturation medium (see Recipes) for each well and culture at 37 °C for 48 h. Half-change the endothelial induction medium on days 9 and 11. Endothelial tissues (organoids) have a round and smooth morphology (Figure 2B).
Embedding organoids in Matrigel, day 12
Preparing dimpled Parafilm:
i. Cut the Parafilm into 5 cm × 5 cm squares, layer it on the gloved finger, and then press the broader round end of the 200 μL tips into the Parafilm to create a dimple. Repeat the same action to make a grid of dimples in the Parafilm (Figure 2D).
ii. Evenly spray 75% alcohol on the two sides of the dimpled Parafilm substrate; then, place the Parafilm in the biosafety cabinet for further sterilization by its own ultraviolet radiation for over 1 h. Place the sterilized Parafilm in sterile 60 mm dishes.
Note: Any other commercial product can be used to replace the dimpled Parafilm.
Preparing Matrigel.
Use growth factor-reduced Matrigel to embed organoids. Place the original concentrated Matrigel in the 4 °C fridge one day in advance to enable natural thawing. All Matrigel operations are conducted on ice.
Use a disposable Pasteur pipette to individually transfer each vessel organoid to each small dimple in the Parafilm, manually aspirate the extra medium with a 10 μL pipette tip, and then gently drip 10 μL of Matrigel onto each organoid. Gently move the organoid with the tip and adjust its position to the center of the Matrigel droplet.
Note: Ensure caution to draw as little liquid as possible with the organoids in the dimples and embed with the Matrigel as soon as possible after draining the liquid to avoid the drying out of organoids.
After all the organoids are embedded in the Matrigel, move them into a 37 °C incubator and solidify the Matrigel droplets for over 30 min.
Remove the embedded organoids from the Parafilm. Clamp the Parafilm with sterilized tweezers and place in the wells of a 6-well plate or dish with the expansion medium (see Recipes). Make the embedded organoids fall off the Parafilm sheet by gently dripping the medium onto the solidified Matrigel with a 1,000-μL tip.
Culture the Matrigel-embedded organoids in a 6-well plate or 60 mm dish for four days with 3 mL or 6 mL of expansion medium per well or dish.
Note: No more than 10 organoids in a well (6-well plate).
Optional step: Manually shake the dish daily to prevent the organoids from sticking to the bottom.
Vessel organoid maturation, days 16–40 (the morphology is shown in Figure 2C)
On day 16, replace the expansion medium with the maturation medium (see Recipes) and move the organoids onto a shaker in the CO2 incubator (the shaker speed is 25–30 revolutions per minute).
From day 16 onwards, half-change the medium every 2–3 days.
Note: Due to faster nutrient consumption as reflected by the color change to yellowish after half-changing the medium, the organoids at the late stages can be placed in a T25 flask that contains more culture medium. Please add fresh medium if the old medium evaporates to less than 3 or 6 mL per well or dish.
Brain organoid generation (schematic view shown in Figure 3)
Figure 3. Schematic procedure of generating brain organoids (BOrs) from hESCs
Making EBs, day 0
Culture H9 cells as described in section A; when cell density reaches 80% confluence and shows good growth status (flat and well-defined edges of colonies and high nucleus/cytoplasm ratio of cells), the stem cells can be used for organoid generation.
Detach H9 cells from the Matrigel-coated plate by adding Accutase solution into the wells. Incubate the cells for 10 min at 37 °C in the CO2 incubator and then add 2 mL of mTeSR into the well. Collect the medium into a 15 mL conical tube and centrifuge at 200× g for 2 min.
Cell counting:
Remove the supernatant and resuspend the cells with 1 mL of mTeSR containing 10 μM Y27632. Pipette cell suspension two to three times and then dilute cells with fresh aggregation medium at a reasonable ratio for cell counting. Only count live cells devoid of trypan blue labeling.
Add 20 μL of Lipidure® solution into each well of a 96-well V-bottom plate for 10 min; aspirate the Lipidure® and dry the plate in the biosafety cabinet for 5 min.
Dilute stem cells with the fresh and warm aggregation medium to 7,000–9,000 cells per 150 μL of medium and seed the cells into the V-bottom wells of a 96-well plate. Culture the cells in the CO2 incubator at 37 °C for 48 h to form the EBs.
Note: In general, 7,000 cells per well are used for generating brain organoids.
Neuroepithelium induction, days 2–10
Day 2: Replace the aggregation medium with 150 μL of the ectoderm induction medium (see Recipes) for each well and culture in the CO2 incubator at 37 °C for 48 h.
Day 4: Half-change the ectoderm induction medium in each well and continue the culturing at 37 °C for an additional 48 h.
Day 6: Replace the ectoderm induction medium with 150 μL of neuroepithelium induction medium (see Recipes). After 48 h, half-change the neuroepithelium induction medium on day 8 and then on day 10. Continue culturing at 37 °C for six days.
Embedding neuroepithelial tissues in Matrigel, day 12
In this step, the neuroepithelial tissues are ready for fusion with the vessel organoids. However, if the researcher wants to separately culture brain organoids, this embedding step is the same as described in step B3.
Brain organoid maturation, days 16–40
On day 16, replace the neuroepithelium induction medium with the maturation medium and move the Matrigel droplets to a shaker placed in the CO2 incubator (the shaker speed should be 25–30 rotations/min).
From day 16 onwards, half-change the medium every 2–3 days.
Note: Please add fresh medium if the old medium has evaporated to less than 3 or 6 mL per well or dish.
Vascularized brain organoid generation
Generate vessel organoids by following section B (days 0–10) of this protocol.
Generate neuroepithelial tissues by following section C (days 0–10) of this protocol.
Fusion of the vessel and brain organoids.
The preparation steps are the same as those for the culture of vessel organoids on day 12. Gently transfer two vessel organoids and one neuroepithelial tissue one by one to each dimple in the Parafilm; aspirate the excess medium from each tissue and then drip 25 μL of Matrigel onto the tissues.
Gently align the three tissues with the neural tissue in the middle of the two vessel organoids using a 10 μL pipette tip. Adjust the position to maintain the three tissues in the same horizontal plane and at the center of the droplet.
Place the droplets with embedded tissues on the Parafilm back into the 37 °C incubator to polymerize the Matrigel for over 40 min. Detach the solidified tissue droplets from the Parafilm as described above and then culture them in a 6-well plate or 60 mm dish for four days in the expansion medium, with each well or dish containing 3 or 6 mL of medium.
Note: We suggest the use of no more than five fusion tissues in one well. If more than five fusion tissues are cultured, transfer them into a T25 flask using a sterile Pasteur pipette (3 mL) with a broader cutting mouth.
On day 16, replace the expansion medium with the maturation medium and move the tissues to a shaker placed in the incubator.
As of day 16, half-change the medium every 2–3 days to facilitate the development of the cerebral brain organoid and its vascularization by vessel organoids. Observe the organoid morphology at various stages (Figure 2G).
Note: Check the color of culture medium and refresh it if it becomes yellow. Please add fresh medium if the old medium has evaporated to less than 3 or 6 mL per well or dish.
Data analysis
Immunofluorescence
The immunohistochemical images shown in this protocol were generated using whole-mount staining (Figure 4, Figure 5A) or cryo-section staining (Figure 5B). Briefly, the collected organoid samples were fixed in 4% paraformaldehyde (PFA) at 4 °C overnight and then washed three times with DPBS, followed by incubation in 0.5% Triton X-100 at RT (22–25 °C) for 1 h. After blocking with 5% bovine serum albumin (BSA) in 0.1% Triton X-100 at RT for 1 h, organoids were incubated with primary antibodies at 4 °C for over 48 h, washed with DPBS, and then incubated with secondary antibodies at 4 °C for over 48 h. Stained organoids were washed with DPBS three times before confocal imaging. For cryosection staining, the organoids were fixed in 4% PFA at 4 °C overnight and then washed three times with DPBS; then, the samples were dehydrated in 30% sucrose at 4 °C for 24–48 h. Subsequently, the organoids were embedded in OCT compound and cryo-sectioned into 30 μm thick slices. The sectioned slices were boiled in citrate-based antigen retrieval buffer for 10 min, followed by cooling for 60 min. Slices were washed with DPBS three times and incubated in 0.3% Triton X-100 at RT for 30 min, blocked with 5% BSA in 0.1% Triton X-100 at RT for 1 h, and incubated with the primary antibody at 4 °C for over 48 h, followed by washing with DPBS and incubation with the secondary antibody at 4 °C overnight. Secondary antibodies used were Alexa Fluor 488-, 555-, 594-, or 647-conjugated donkey anti-mouse, anti-rabbit, anti-rat, or anti-chicken immunoglobulin G (IgG) respectively (Invitrogen; all used at 1:1,000 dilution). DAPI (2.5 μg/mL) was used to stain the cell nuclei. Primary and secondary antibodies were prepared in DPBS containing 0.1% Triton X-100.
Figure 4. Identification by cell marker staining on vessel organoids (VOrs) and fusion vascularized brain organoids (fVBOrs). (A) Immunostaining of GFP and CD31 in D40 VOrs. Scale bar, 200 μm. Right: Imaris reconstruction of VOrs showing integrated vasculature structures. (B) Immunostaining of CD31 and DCX for labeling vessels and neurons, respectively, in D40 fVBOrs. Scale bar, 200 μm. Images in Figure 4 are quoted from the published article, Figure 1D and Figure 4C (Sun et al., 2022).
Figure 5. Microglial cells induced in vessel organoids (VOrs) and fusion vascularized brain organoids (fVBOrs). (A) Immunostaining of IBA1 for the labeling of microglial cells in D25 and D40 VOrs. Scale bar, 200 μm. (B) Immunostaining of IBA1 for the labeling of microglial cells in brain organoids (BOrs) and fVBOrs, respectively. Scale bar, 200 μm. Images in Figure 5 are quoted from the published article, Figure 5D and Figure 5F (Sun et al., 2022).
Analysis of the developmental process
During culturing, vessel organoids at various developmental stages were imaged from days 2 to 40, as shown in Figures 2B and C. The diameters of the vessel organoids were then quantified (Figure 2E).
As shown in Figure 4A, the vessel organoids exhibited robust vasculogenesis and complex vascular structures expressing the endothelial cell marker CD31. Furthermore, the fusion vascularized brain organoids showed that doublecortin-labeled neurons were surrounded by invaded vessels positively labeled with anti-CD31 antibodies, indicating a connection between neural and blood vessel structures (Figure 4B). Notably, microglial cells were induced in blood vessel organoids (Figure 5A), and these microglial cells invaded the vascularized brain organoids during fusion (Figure 5B). For other experiments on vessel organoids, such as the staining of cell type markers and single-cell analyses, please refer to Figures 1–3 of the published paper (Sun et al., 2022); for the downstream analysis of the fusion vascularized brain organoids, including the blood-brain-barrier, immune functional analyses, and neuro–vascular interactions, please refer to Figures 4–6 of the published paper (Sun et al., 2022).
Validation of protocol
This protocol has been used and validated in the following research article: Sun et al. (2022). Generation of Vascularized Brain Organoids to Study Neurovascular Interactions. eLife 11: e76707. DOI:10.7554/eLife.76707.
General notes and troubleshooting
Problem 1: Small EBs or scattered cells in V-bottom 96-well plate.
Possible cause: Unhealthy ESCs.
Solution: Culture the stem cells to a good state for generating organoids. (Good state means: high nucleus/cytoplasm ratio of cells, flat and well-defined edge of colonies, and approximately 80% cell confluent.)
Problem 1: Small EBs or scattered cells in V-bottom 96-well plate.
Possible cause: Insufficient cells or starting cells with an extensive number of dead cells.
Solution: Calculate the cells accurately with trypan blue.
Problem 1: Small EBs or scattered cells in V-bottom 96-well plate.
Possible cause: Omitted treatment of the wells with Lipidure® solution.
Solution: Treat the wells with Lipidure® solution.
Problem 2: The VOrs detached from the Matrigel when embedding.
Possible cause: Insufficient time for Matrigel solidification.
Solution: Prolong the time for Matrigel solidification in a 37 °C incubator.
Problem 3: Separation of organoid and Matrigel after embedding.
Possible cause: Strong pipetting during the medium change.
Solution: Gently add fresh medium along the wall of the well or dish; avoid touching the Matrigel with tips during medium renewal.
Problem 3: Separation of organoid and Matrigel after embedding.
Possible cause: Fast shaking speed.
Solution: Check the shaking speed daily and keep the shaker at a low speed.
Problem 4: VOrs show a bad state without strong vasculogenesis during the maturation stage.
Possible cause: The VOrs stick to the bottom of the dishes.
Solution: Put the VOrs in a rotating shaker to avoid sticking.
Acknowledgments
This work was partially supported by STI2030-Major Projects (2021ZD0202500), the National Natural Science Foundation of China (32130035 and 92168107 to Z.G.L., and 31871034 to X.C.J.), the Frontier Key Project of the Chinese Academy of Sciences (QYZDJ-SSW-SMC025), Shanghai Municipal Science and Technology Projects (2018SHZDZX05), and Shanghai Frontiers Science Center for Biomacromolecules and Precision Medicine at ShanghaiTech University. We thank the MultiOmics Core Facility, Molecular Imaging Core Facility, and Molecular and Cell Biology Core Facility at the School of Life Science and Technology, ShanghaiTech University, for providing technical support. The protocol described here was based on the previous original research paper (Sun et al., 2022).
Competing interests
The authors declare no competing interests.
Ethics
This study has not used human or animal subjects. The usage of commercially available H9 human embryonic cell line has been approved by the research ethics committee of the ShanghaiTech University.
References
Amin, N. D. and Paşca, S. P. (2018). Building Models of Brain Disorders with Three-Dimensional Organoids. Neuron 100(2):389–405.
Benito-Kwiecinski, S., Giandomenico, S. L., Sutcliffe, M., Riis, E. S., Freire-Pritchett, P., Kelava, I., Wunderlich, S., Martin, U., Wray, G. A., McDole, K., et al. (2021). An early cell shape transition drives evolutionary expansion of the human forebrain. Cell 184(8): 2084–2102.e19.
Cakir, B., Xiang, Y., Tanaka, Y., Kural, M. H., Parent, M., Kang, Y. J., Chapeton, K., Patterson, B., Yuan, Y., He, C. S., et al. (2019). Engineering of human brain organoids with a functional vascular-like system. Nat. Methods 16(11): 1169–1175.
Di Lullo, E. and Kriegstein, A. R. (2017). The use of brain organoids to investigate neural development and disease. Nat. Rev. Neurosci. 18(10): 573–584.
Lancaster, M. A. and Knoblich, J. A. (2014). Generation of cerebral organoids from human pluripotent stem cells. Nat. Protoc. 9(10): 2329–2340.
Lee, H. S., Han, J., Bai, H. J. and Kim, K. W. (2009). Brain angiogenesis in developmental and pathological processes: regulation, molecular and cellular communication at the neurovascular interface. FEBS J. 276(17): 4622–4635.
Mansour, A. A., Gonçalves, J. T., Bloyd, C. W., Li, H., Fernandes, S., Quang, D., Johnston, S., Parylak, S. L., Jin, X., Gage, F. H., et al. (2018). An in vivo model of functional and vascularized human brain organoids. Nat. Biotechnol. 36(5): 432–441.
Pham, M. T., Pollock, K. M., Rose, M. D., Cary, W. A., Stewart, H. R., Zhou, P., Nolta, J. A. and Waldau, B. (2018). Generation of human vascularized brain organoids. NeuroReport 29(7): 588–593.
Shi, Y., Sun, L., Wang, M., Liu, J., Zhong, S., Li, R., Li, P., Guo, L., Fang, A., Chen, R., et al. (2020). Vascularized human cortical organoids (vOrganoids) model cortical development in vivo. PLoS Biol. 18(5): e3000705.
Siegl, D., Kruchem, M., Jansky, S., Eichler, E., Thies, D., Hartwig, U., Schuppan, D. and Bockamp, E. (2023). A PCR protocol to establish standards for routine mycoplasma testing that by design detects over ninety percent of all known mycoplasma species. iScience 26(5): 106724.
Sun, X. Y., Ju, X. C., Li, Y., Zeng, P. M., Wu, J., Zhou, Y. Y., Shen, L. B., Dong, J., Chen, Y. J., Luo, Z. G., et al. (2022). Generation of vascularized brain organoids to study neurovascular interactions. eLife 11: e76707.
Tata, M., Wall, I., Joyce, A., Vieira, J. M., Kessaris, N. and Ruhrberg, C. (2016). Regulation of embryonic neurogenesis by germinal zone vasculature. Proc. Natl. Acad. Sci. U.S.A. 113(47): 13414–13419.
Wakui, T., Matsumoto, T., Matsubara., K., Kawasaki., T., Yamaguchi., H. and Akutsu, H. (2017). Method for evaluation of human induced pluripotent stem cell quality using image analysis based on the biological morphology of cells. J. Med. Imaging 4(4): 1.
Wimmer, R. A., Leopoldi, A., Aichinger, M., Wick, N., Hantusch, B., Novatchkova, M., Taubenschmid, J., Hämmerle, M., Esk, C., Bagley, J. A., et al. (2019). Human blood vessel organoids as a model of diabetic vasculopathy. Nature 565(7740): 505–510.
Wörsdörfer, P., Rockel, A., Alt, Y., Kern, A. and Ergün, S. (2020). Generation of Vascularized Neural Organoids by Co-culturing with Mesodermal Progenitor Cells. STAR Protoc. 1(1): 100041.
Xiang, Y., Tanaka, Y., Cakir, B., Patterson, B., Kim, K. Y., Sun, P., Kang, Y. J., Zhong, M., Liu, X., Patra, P., et al. (2019). hESC-Derived Thalamic Organoids Form Reciprocal Projections When Fused with Cortical Organoids. Cell Stem Cell 24(3): 487–497.e7.
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Stem Cell > Pluripotent stem cell
Biological Sciences > Biological techniques
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4,871 | https://bio-protocol.org/en/bpdetail?id=4871&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Studying Cell Migration (Random and Wound Healing) Parameters with Imaging and MATLAB Analysis
LY Ling-Yea Yu *
HL Hsuan-Chao Lin *
CH Chi-Lin Hsu *
TK Tuan-Yu Kao
FT Feng-Chiao Tsai
(*contributed equally to this work)
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4871 Views: 509
Reviewed by: Alka MehraAnaïs Panosa Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances Jul 2021
Abstract
Cell migration is an essential biological process for organisms, in processes including embryonic development, immune response, and cancer metastasis. To elucidate the regulatory machinery of this vital process, methods that mimic in vivo migration, including in vitro wound healing assay and random migration assay, are widely used for cell behavior investigation. However, several concerns are raised with traditional cell migration experiment analysis. First, a manually scratched wound often presents irregular edges, causing the speed analysis difficult. Second, only the migration speed of leading cells is considered in the wound healing assay. Here, we provide a reliable analysis method to trace each cell in the time-lapse images, eliminating the concern about wound shape and creating a more comprehensive understanding of cell migration—not only of collective migration speed but also single-cell directionality and coordination between cells.
Keywords: Live-cell imaging Single-cell tracing Wound healing assay Random migration
Background
Cell migration is an individual or collective cell movement that plays an important role in embryonic development (Weijer, 2009), immune response (Luster et al., 2005), and cancer metastasis (Friedl and Glimour, 2009). To investigate these essential physiological phenomena in vitro, wound healing assay or random migration assay are often applied (Jin et al., 2016).
Compared with traditional analyses of wound healing assay focusing on general migration speed, our script mainly focuses on precisely tracking each cell in each timeframe image, leading to speed, directionality, and coordination being analyzed simultaneously. Moreover, we can conquer the difficulties in analyzing the wound healing assay even when the edge of the wound is not smooth.
Since wound healing assay provides cell information like migration speed and directionality, and random migration provides cell coordination information, we applied both assays to elucidate the mechanisms of cell migration thoroughly. We found several pairs of genes and pathways that affect migration speed and cell coordination through these assays after performing two-hit inhibition: the knockdown of a migration-related gene using shRNA combined with the addition of an inhibitor of migration-related pathways at the same time. One of our best candidates, STK40 and MAPK, showed a synergistic effect on migration speed. By applying shSTK40 and MAPK inhibitor PD98059, we found that the absence of either of them has a mild effect on cell migration speed, while the speed significantly dropped when both of them were inhibited at the same time.
Our method is a half-automatic way to trace single cells in the time-lapse immunofluorescence images, which provides multiple parameters for a comprehensive analysis of cell migration, including speed, density, directionality, and cell–cell coordination. In sum, our method not only solves the aforementioned concern but provides a fast, cheap, and comprehensive analysis of cell migration.
Materials and reagents
96-well black/clear-bottom plate (Thermo Fisher Scientific, NuncTM, catalog number: 165305)
Collagen I, rat tail (Thermo Fisher Scientific, GibcoTM, catalog number: A1048301)
Hoechst 33342 (Thermo Fisher Scientific, InvitrogenTM, catalog number: H3570)
HEPES (Thermo Fisher Scientific, GibcoTM, catalog number: 15630106)
Bovine serum albumin (BSA) (BioShop, catalog number: ALB001)
Recombinant human EGF (PeproTech, catalog number: AF-100-15)
Human FGF-acidic recombinant protein (Thermo Fisher Scientific, GibcoTM, catalog number: PHG0014)
Heparin sodium salt from porcine intestinal mucosa (Merck, Sigma-Aldrich, catalog number: H3393)
Trametinib (LC laboratories, catalog number: T-8123)
PBS, pH 7.4 (Thermo Fisher Scientific, GibcoTM, catalog number: 10010023)
Cell migration supplements (see Recipes)
Cell lines
SAS cell line was used as indicated. Cells were grown in Dulbecco’s modified Eagle medium (DMEM) with 10% fetal bovine serum (FBS), and 1% penicillin and streptomycin.
Recipes
Cell migration supplements
20 mM HEPES
0.1% BSA
5 ng/mL EGF or 25 ng/mL FGF1
10 U heparin
Equipment
Fluorescence microscope (Nikon Eclipse Ti)
96-well scratcher (can be replaced by 200 μL tip or toothpick)
Software
MATLAB (MathWorks, https://www.mathworks.com/products/matlab.html) with the image processing toolbox (access date, 8/1/2017)
Nikon NIS-Elements Viewer (Nikon image acquisition software) (access date, 9/21/2017)
Fiji from ImageJ (open resource software for image analysis) (access date, 8/1/2017)
Excel (Microsoft) (access date, 8/1/2017)
Procedure
Plate coating and cell seeding
Add 100 μL of collagen I (30 μg/mL, diluted in PBS) to the 96-well black/clear-bottom plate and put the plate in a 37 °C, 95% humidity incubator for 30–40 min.
Rinse the plate with 100 μL of PBS and aspirate the liquid in the plate thoroughly before use.
After that, plate SAS cells onto the plate. (The cell number should be controlled to 90%–100% confluency on the day performing cell migration.)
(Optional) Cells were infected with lentiviruses for 24 h and selected with antibiotics for 24–48 h.
Nucleus staining
Stain the cells with 1 μg/mL of Hoechst 33342 at 37 °C for 1 h for SAS cells.
Scratching (for wound-healing assay only)
Change the medium to 100 μL of pre-warmed PBS.
Scratch the cells with a scratcher (move the scratcher back and forth twice). This method is for a full 96-well plate and can be replaced by using toothpicks or 200 μL tips if only part of the 96 well plate is used.
After wound-scratching, wash the cells with 100 μL of PBS twice to remove cell fragments.
Drug treatment
Change the medium to serum-free DMEM medium (for SAS cells) containing cell migration supplements and drugs (we used trametinib 100 nM or DMSO).
Imaging
Place the cells in an acrylic incubator at 32 °C for SAS cells. The acrylic incubator is a chamber with CO2 circulation [the temperature is lower than normal incubation (37 °C) to optimize the migration speed for analysis].
Use Nikon Eclipse Ti microscope for live-cell imaging. Take images every 10 min for a duration of 6–10 h. Our script is designed for pictures taken by 4× objective.
Data analysis
We process the image data through our MATLAB scripts. The schematic summary of how these images are processed is in Figure 1.
Figure 1. Schematic summary of data processing using MATLAB scripts
Description of MATLAB scripts and analysis
Define nuclei location: Z1_calcnuclei_adjustedPlateShift
This script defines the cell location in each image.
Open MATLAB, import data, and prepare the script.
Export images to working directory: an individual image is required for DAPI channel and time-lapse.
Note: The file name should begin with “well + row number + column number” to fit our script. If not, this could be changed in line 13. All images should be exported as tiff files (Figure 2).
Figure 2. Screenshot of Z1_calcnuclei_adjustedPlateShift user interface. Underlined is the path of exported tiff files. Note that the image file name in the folder should follow the rule: well + row number + column number + other details, e.g., WellA01_ChannelDAPI _Seq0000_ Z01_C1.tif (L12).
Open Z1_calcnuclei_adjustedPlateShift.m.
(Optional) Change the nuclear size in line 4. The nucleus size should be changed based on your cell line (in this case, we use 4 for SAS to best discriminate each cell).
Change the file path in line 3 and fill the row and column number based on your experimental setting in line 6 and 8. (In this case, we choose row B–G, and column 1–12, so we fill 2:7 for row and 1:12 for column, respectively.) (Figure 3)
Figure 3. Example of a 96-well plate setting corresponding to the row and column number in Z1_calcnuclei_adjustedPlateShift
This script defines nucleus based on a modified threshold selection method. By measuring fluorescence signals (DAPI; Channel 1) to define centroid of every nucleus, we can acquire the location information of each cell from all the acquired images. This allows users to get all the information of cell location.
We first retrieved the centroid of each cell for further calculation of cell movement. The highest signal in the DAPI channel within an estimated distance would be seen as cell centroids, and we labeled these signals as red dots in the image (Figure 4, left panel). To confirm the nucleus definition correctness, we examine the overlap of red dots and green dots (nuclei mask) in the representative figure (Figure 4, right panel). The original Hoechst signal is labeled as red, and our nuclei mask is labeled as green. The nuclei mask is based on Hoechst signal with extra restrictions on size and shape to eliminate noise (such as cell debris) from our analysis. The higher the overlapped ratio of red and green dots, the higher the precision of recognizing the cells. Then, we compare the images between centroids and nuclei masks to double-check if our centroids are in the right place.
Figure 4. Example of capturing cell nuclei after running Z1_calcnuclei_adjustedPlateShift. (A) The red dots are the highest Hoechst signal compared with its surrounding signal as our centroids; this labeled centroid area is magnified in (C). (B) Overlap of the image in the panel with the nuclei mask (green) generated by the processed Hoechst signal and the original Hoechst signal (red), which is magnified in (D). The white arrows in (A) and (B) indicate the direction of cell migration, and the white squares are the cells chosen for magnification.
Monitoring cell movement: Z2_trace nuclei
This script calculates changes of cell location between each time frame.
Open Z2_trace nuclei.m.
Change the file path in line 3 and fill in the row and column number based on your experimental setting in line 7 and 9 (Figure 5).
(Optional) Change the nuclear size (line 4). The nucleus size should be changed based on your cell line (in this case, we use 4 for SAS to best discriminate each cell).
Click Run.
Figure 5. Screenshot of the Z2_trace nuclei user interface. The underlined and circled areas need to be adjusted in each experiment.
Get cell information: Z3_getcellparameters062945
Parameters such as speed, coordination, and directionality were analyzed using MATLAB.
Open Z3_getcellparameters062945.m.
Change the file path in line 3 and fill the row and column number based on your experimental setting in line 7 and 9 (Figure 6A).
(Optional) Change the nuclear size in line 4. The nucleus size should be changed based on your cell line (in this case, we use 4 for SAS to best discriminate each cell).
Click Run.
Define the number of analyzed cells:
Fill out the cell number in line 16 that you would like to analyze (Figure 6A). Selected cells should be marked as red. Below is an example of analyzing 1,000 and 10,000 cells (Figure 6B). Note that for random migration, we select all the cells in the frame.
Figure 6. Example of different parameter settings for cell numbers used for migration analysis. (A) User interface of Z3_getcellparameters062945. The circled area defines the number of cells being analyzed in tracking. (B) The left panel shows the analysis of 1,000 cells, and the right panel shows the analysis of 10,000 cells. The black dots represent the cells in the entire image, and the red dots represent the cells going to be analyzed. We also show the XY axis from 0 to 1,600 to better spot the exact cell location from the figure and exported data.
Data will be saved under the file called cell data. File name: Well name + bestsp.
Parameters will be saved into several spreadsheets (Figures 7 and 8, Table 1); each spreadsheet is a parameter of migration behavior. An example of exported data is shown in Figure 8. Note that the angle data (wellang) need to be further calculated through the formula: π/2 - θ. θ is the original value shown on the spreadsheet.
Figure 7. Screenshot of Z3_getcellparameters062945 in workspace after running this script. MATLAB files have been generated after running Z3_getcellparameters062945, and the circled files are exported to Excel for further analysis.
Figure 8. Graphical schematic of variables used in cell migration analysis. The directionality is the angle of cell movement relative to the wound direction. Retrieved from Yu et al. (2021).
Table 1. Migration statistics
Variable Calculation Description
Wellang π/2-α Average angle of cell movement relative to the wound direction in the well
wellcoorCos Cos θ Average cell–cell coordination in the well
wellden Total cell number in each frame Cell number of this frame. Note that cell migration speed could be influenced by cell density, so this data has to be quite consistent among each well
wellspeed Distance/time Average cell migration speed in the well. The unit is μm/h.
wellper Displacement/distance Average consistency of directionality of the cell movement in the well
To make the definition clear, Figure 7 explains the variables being used.
Although our scripts generate all variables, we use different parameters for analyzing different cell migrations. For wound healing assay, we analyzed the speed, angle, coordination, and persistence of cell migration. However, we do not analyze cell angles for random migration because we define cell angle as how cells deviate from the direction of the wound. Here, we provide an example of the data we used for analyzing the wound healing assay result (Figure 9).
Figure 9. xample of the population of cell data. Each data point corresponds to the mean value of each well. These data are from row B–G, column 2–9 of a 96-well plate.
These data could be plotted into bar graphs for visualization (Figure 10).
Figure 10. Example of the bar graph for cell migration analysis. Retrieved from Yu et al. (2021).
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Yu et al. (2021). Synthetic dysmobility screen unveils an integrated STK40-YAP-MAPK system driving cell migration. Sci. Adv. 7(31): eabg2106 (Figure 1, panel C and G, and Figure 2, panel B and C).
Acknowledgments
Funding: This work was supported by grants from The Ministry of Science and Technology in Taiwan (MOST 107-2320-B-002-038-MY3 and MOST 108-2926-I-002-002-MY4), National Taiwan University Hospital (NTUH 106-T02, NTUH 107-T13, NTUH 108-T13, and VN 109-14), and The Liver Disease Prevention and Treatment Research Foundation in Taiwan. This protocol was adapted from Yu et al. (2021).
Competing interests
The authors declare that they have no competing interests.
References
Friedl, P. and Gilmour, D. (2009). Collective cell migration in morphogenesis, regeneration and cancer. Nat. Rev. Mol. Cell Biol. 10(7): 445–457.
Jin, W., Shah, E. T., Penington, C. J., McCue, S. W., Chopin, L. K. and Simpson, M. J. (2016). Reproducibility of scratch assays is affected by the initial degree of confluence: Experiments, modelling and model selection. J. Theor. Biol. 390: 136–145.
Luster, A. D., Alon, R. and von Andrian, U. H. (2005). Immune cell migration in inflammation: present and future therapeutic targets. Nat. Immunol. 6(12): 1182–1190.
Weijer, C. J. (2009). Collective cell migration in development. J. Cell Sci. 122(18): 3215–3223.
Yu, L. Y., Tseng, T. J., Lin, H. C., Hsu, C. L., Lu, T. X., Tsai, C. J., Lin, Y. C., Chu, I., Peng, C. T., Chen, H. J., et al. (2021). Synthetic dysmobility screen unveils an integrated STK40-YAP-MAPK system driving cell migration. Sci. Adv. 7(31): eabg2106.
Supplementary information
The following supporting information can be downloaded here:
Z1_calcnuclei_adjustedPlateShift.m
Z2_trace nuclei.m
Z3_getcellparameters062945.m
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Cancer Biology > Invasion & metastasis > Cell biology assays
Cell Biology > Cell signaling > Intracellular Signaling
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In the Z2_trace nuclei.m script, what is the relocateD function for? What is the definition of this function? Is it an in-house or matlab function?
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Peer-reviewed
Spot Assay and Colony Forming Unit (CFU) Analyses–based sensitivity test for Candida albicans and Saccharomyces cerevisiae
SS Satya Ranjan Sahu
BU Bhabasha Gyanadeep Utkalaja
SP Shraddheya Kumar Patel
NA Narottam Acahrya
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4872 Views: 761
Reviewed by: Alba BlesaFernando A Gonzales-ZubiateThibaud T. Renault
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Original Research Article:
The authors used this protocol in Gut Microbes Jan 2023
Abstract
Cellular sensitivity is an approach to inhibit the growth of certain cells in response to any non-permissible conditions, as the presence of a cytotoxic agent or due to changes in growth parameters such as temperature, salt, or media components. Sensitivity tests are easy and informative assays to get insight into essential gene functions in various cellular processes. For example, cells having any functionally defective genes involved in DNA replication exhibit sensitivity to non-permissive temperatures and to chemical agents that block DNA replication fork movement. Here, we describe a sensitivity test for multiple strains of Saccharomyces cerevisiae and Candida albicans of diverged genetic backgrounds subjected to several genotoxic chemicals simultaneously. We demonstrate it by testing the sensitivity of DNA polymerase defective yeast mutants by using spot analysis combined with colony forming unit (CFU) efficiency estimation. The method is very simple and inexpensive, does not require any sophisticated equipment, can be completed in 2–3 days, and provides both qualitative and quantitative data. We also recommend the use of this reliable methodology for assaying the sensitivity of these and other fungal species to antifungal drugs and xenobiotic factors.
Keywords: Spot assays Colony forming units Fungi Saccharomyces Candida Genotoxic stress Temperature sensitivity DNA damaging agents DNA replication inhibitors Hydroxyurea Methyl methanesulfonate Antifungal drugs Cell survival DNA polymerase knockouts
Background
Sensitivity assays are crucial in determining the susceptibility of a cell to a particular stress and are used to gain insights into diverse cellular processes in both prokaryotic and eukaryotic cells, including yeasts like Saccharomyces cerevisiae, Candida albicans, or Schizosaccharomyces pombe (Kwolek-Mirek and Zadrag-Tecza, 2014). S. cerevisiae and C. albicans are genetically related fungal species and are considered the best model organisms for sensitivity studies in yeast research due to their ease of manipulation, rapid growth, and well-established genetic modification strategies (Botstein et al., 1997; Berman, 2012). The growth curve analysis based on the optical density (OD) measurement in the presence and absence of a cytotoxic chemical is routinely used to monitor the sensitivity of a strain. Here, we describe an alternative protocol that uses a combination of spot and CFU analyses to obtain both qualitative and quantitative sensitivity results of several strains in a single experiment in a short time. We have been using this methodology repeatedly and successfully in our studies for many years now (Manohar et al., 2018; Kumari et al., 2021 and 2023; Manohar et al., 2022; Patel et al., 2023).
In the spot assay, yeast cultures are spotted on agar plates with or without different concentrations of any cytotoxic agents in a dilution series; sensitivity is analyzed based on the density of the cells present in a given spot. Comparison of growth inhibition in a specific spot with and without drugs allows to assess the sensitivity of a strain to that particular drug. Similarly, relative sensitivity to a particular drug can be assessed by comparing various isogenic wildtype and mutant strains. Since spotting will only determine the cumulative qualitative behavior of a group of colonies present in a particular spot, we support this assay by enumerating colony forming unit (CFU) efficiency of a yeast strain by spreading different dilutions of the culture on solid media plates with or without different concentrations of drugs. Estimating the number of colonies and their comparison across drug concentrations or strains makes it a quantifiable sensitivity test. The combination of both assays provides us with a more comprehensive sensitivity profile of a strain. The complete assay involves spotting and spreading serially diluted yeast cells of culture on agar plates; the resulting growth patterns in both types of plates are then used to assess the sensitivity of each strain (Figure 1).
Figure 1. Schematic representation of the methodology to determine sensitivity of isogenic strains to a specific drug or reagent
This protocol shows the steps involved in testing four strains in their sensitivity to the presence of hydroxyurea (HU), a DNA replication inhibitor, and methyl methanesulfonate (MMS), a DNA methylating agent. However, the sensitivity of 12 different strains or drugs can be verified using a 96-well plate. We used wildtype strains of S. cerevisiae and C. albicans and their pol32 null strains for sensitivity tests. Pol32 is the smallest subunit of DNA polymerase delta, a DNA polymerase involved in both lagging and leading strand synthesis (Acharya et al., 2011; Khandagale et al., 2019). In its absence, cells exhibit high temperature sensitivity and growth retardation in the presence of HU, MMS, and other DNA damaging agents (Patel et al., 2023). Here, we repeated the experiment to show the effect of HU and MMS on the growth of these strains by simultaneously using spot and CFU analyses (Figure 2). The differential cell density among the strains in the spot assay and the statistical estimation of the number of colonies in the CFU assay confirmed that pol32-deficient strains are more sensitive to these tested drugs than their respective wildtype strains.
Figure 2. Sensitivity tests for pol32 deficient strains of C. albicans and S. cerevisiae. (A) Cells of wildtype (WT) and pol32 knockout strains of C. albicans (SC5314, diploid) and S. cerevisiae (EMY74.7, haploid) were spotted on YPD-agar media containing different concentrations of hydroxyurea (HU) and methyl methanesulfonate (MMS). (B) Dilutions of these cells were spread to get isolated colonies; colonies were counted and plotted in a line graph as obtained from Table 1 and Table S1. ns = nonsignificant and *** p value < 0.001.
This protocol is simple, cost effective, and does not require any expensive equipment. This assay can be used to assess multiple parameters such as growth phenotypes, viability, stress or drug resistance, and genotoxicity of multiple strains at a time (Kumari et al., 2023). Altogether, this procedure provides a well-standardized, sensitive, and reproducible protocol to assess yeasts’ sensitivity.
Materials and reagents
Yeast strain of interest
Round-bottom glass culture tubes (Borosil, catalog number: 9900006)
250 mL conical flask (Schott Duran, catalog number: 1006940)
Yeast extract peptone dextrose (YPD) (Himedia, catalog number: M1363)
Yeast extract peptone dextrose agar (YPDA) (Himedia, catalog number: G038)
Ethanol (100% and 70%) (Fisher Chemical, catalog number: 2051537)
Hydroxyurea (MP Biomedical, catalog number: 102023)
MMS (SRL, catalog number: 74384)
Sterile 1.5 mL microcentrifuge tube (Axygen, catalog number: MCT150LC)
Round U-bottom 96-well plate (Thermo Fisher Scientific, catalog number: 165306)
Sterile Petri dish 100 mm × 15 mm (Falcon, catalog number: 351008)
Glass Petri dish 100 mm × 15 mm (Borosil, catalog number: 3165077)
Sterile spreader (Tarson, catalog number: 920081)
Cuvettes 200–1,600 nm (Eppendorf, catalog number: 0030106300)
Aluminum foil (Rolias)
Gloves (Blue Shield)
Equipment
Laminar airflow (Thermo Scientific Biological Safety Cabinets, catalog number: 41346502)
Sterile pipette sets (Rainin, catalog number: E1338890T-SL1000, B637036181-SL200, B641141423-SL20)
Laboratory spirit lamp or Bunsen burner (VWR, catalog number: 17822-605)
Spectrophotometer (Eppendorf, Bio Photometer Plus, catalog number: 6132)
48-pin spotter (Sigma-Aldrich, catalog number: R2383)
Refrigerated shaker incubator (Scigenic Biotech, catalog number: LE-4676-AH)
Refrigerated incubator (Scigenic Biotech, catalog number: C-1NC-100)
Chemidoc XRS gel imager (Bio-Rad, catalog number: 1708370)
White Light box (Cole-Parmer: NC1851470)
Software
Bio-Rad Image lab (2017, version number: 6.0.1)
GraphPad prism v8.0
Procedure
These steps must be carried out in a sterile environment, preferably using a biosafety cabinet. Before starting the experiment, wipe the laminar airflow hood with 70% ethanol, place all necessary equipment (Petri dish, 96-well plates, spirit lamp, spotter, pipette set, sterile tip box, 1.5 mL MCT tubes) inside, and switch on the UV for 30 min. Use proper personal protective accessories such as laboratory jackets and gloves to prevent contamination and maintain your own safety.
Strain inoculation
Freshly streak all the yeast strains of interest from glycerol stock on a sterile YPD agar plate and incubate for 48 h at 30 °C.
Inoculate a single colony of each strain in 5 mL of autoclaved YPD broth medium and allow it to grow overnight (14–16 h) at 30 °C in a shaker incubator at 200 rpm.
YPD agar plates pouring
Take freshly autoclaved YPD agar media and swirl gently until it cools enough to handle. Once you are able to hold the flask comfortably for a few seconds, pour approximately 25 mL of media per plate (see Note 1).
To prepare YPD plates with HU or MMS (or any other reagents), take freshly autoclaved YPD agar media and swirl gently until it cools down to a temperature comfortable to handle. At this point, add the required volume of the chemicals’ stock solution (e.g., 100 mM HU or 99% MMS) to individual flasks, to obtain the final concentrations. Mix well by swirling and then pour. For example: for our strains, the final concentration of HU and MMS will be 5, 10, and 20 mM, and 0.001%, 0.002%, and 0.003%, respectively. A total of four plates (one for spotting and three for spreading) for each reagent concentration is required (see Note 2).
Allow the plates to cool down completely for 45–60 min; this is necessary for efficient spotting and spreading. Afterward, cover the plates and keep them in an inverted position. Critical: Before pouring the plates with any reagents, strictly follow each toxic reagent’s user manual regarding safety, light sensitivity, thermal stability, and solubility. While pouring the plates with light-sensitive reagents, switch off the laminar hood light and cover the plates with aluminum foil. MMS and other mutagenic reagents have to be handled carefully; detoxify the flasks, plates, etc. with MMS contamination as per the supplier’s recommendation as soon as the experiment is completed. Label the plates according to the concentration before pouring the plates. Put different arrow marks to show the direction of strains and dilutions. Pause point: You can store plates at 4 °C by wrapping them with parafilm and aluminum foil for next-day spotting/spreading. Long-term storage is not encouraged, as the plates will dry up and the local concentration of the drugs will increase (see Additional note 1).
Serial dilution
Dilute the overnight grown culture to 1:10 ratio with YPD broth in a 1.5 mL microcentrifuge tube and measure the optical density (OD) in a spectrophotometer to determine culture density. Critical: The 600 nm OD of the undiluted culture is erroneous if it goes beyond 3; therefore, it is recommended to dilute the culture with YPD to keep OD around 1.
The concentrations of different cytotoxic chemicals need to be standardized based on the strains, which may vary from experiment to experiment when more strains are included in sensitivity tests. Prior information based on published literature will be helpful.
Make an equal number of cells of all strains by maintaining the OD 1.0 at 600 nm in the spectrophotometer. Double-check the OD to avoid variability in cell number.
Add 200 μL of 1.0 OD culture of each strain to the first row of a 96-well plate (S1–S12); one set of samples will be for spotting and the other for the spreading CFU assay.
Fill the five successive downstream wells in the columns (B1, C1, D1, E1, and F1, and similar) with 180 μL of YPD broth for serial dilution.
Perform serial dilution by pipetting out 20 μL of culture from the main culture (A1) to the downstream well and continue up to the sixth well (F1) (10-5 dilution) for the first set of samples. By doing so, 6 × 106 cells in A1 get diluted to 60 cells in F1. The same dilution will be performed for the rest of the strains (see Note 3). Critical: Before pipetting out 20 μL, mix it evenly for uniform cell density and efficient serial dilution.
Spotting of dilutions
Sterilize the spotter by dipping the pins in 100% ethanol kept in a glass Petri dish, followed by brief flaming three times. Allow the spotter to cool down (5–10 min) (see Note 4). Caution: Keep the spirit lamp or Bunsen burner away from the vertical airflow well inside the laminar hood. Placing the lamp too close to the airflow may cause over-flaming due to the introduction of air, which could potentially lead to an explosion of the laminar hood. Place the spirit lamp and the Petri dish with 100% ethanol in two separate corners of the laminar hood. Immediately close the lid of the spirit lamp after flaming.
Once the spotter cools down completely, dip the spotter into the 96-well plate with the serial dilutions of cultures and spot gently on the agar plates with or without cytotoxic agents. Take out the spotter vertically without disturbing the plate (see Note 5). Spotting has to be done separately for each plate.
After spotting, leave the plates undisturbed for 10–15 min to ensure complete absorption of spotted culture; keep the plates at 30 °C for 24–48 h depending upon the growth on the plates. Critical: If the additive reagent is light-sensitive, perform spotting by switching off the laminar hood light and cover with aluminum foil during incubation. After 24 h, monitor plates at regular intervals to avoid overgrowing of spots.
Spreading of dilutions for estimating CFU
Pipette out 100 μL of 10-4 diluted samples from the fifth-row wells (E1–E12), transfer into a 1.5 mL microcentrifuge tube, and make up the volume to 600 μL with YPD broth. Critical: Depending upon the number of plates required to spread, select the dilution accordingly from the 96-well plates (10-3 or 10-4) for further dilution in 1 mL microcentrifuge tubes. If the number of plates is > 6, one can select a lower dilution and continue.
Thoroughly mix the culture in a 1.5 mL microcentrifuge tube by pipetting with a 1 mL pipette or by vortexing for a few seconds. Spread 100 μL of the resultant samples on the plates with or without reagents. Three plates containing the same concentration of HU and one YPD agar control plate will be used for spreading to get statistical significance.
Incubate all plates at 30 °C for 24–48 h depending upon the growth of isolated colonies on the plates (see Additional note 2). Critical: Too few or too many colonies on the plates may give a false estimation; therefore, the dilution has to be selected in such a way that approximately 50–80 isolated colonies will appear on a plate. Pause point: Prior to imaging/counting of colonies, one can keep the plates at 4 °C by wrapping them in parafilm and aluminum foil (Figure S1).
Place the plate on a light box and count the colonies manually. Note down the information with respect to each plate immediately (Additional note 3).
Plates imaging
Place the YPD plate from both spot and CFU analyses in an upright position on a white plate of Bio-Rad gel doc and remove the plate cover to prevent reflection. Open the Image lab software, select the stain-free blot in the blot section, and acquire the image. For best imaging, place two plates at a time (see Note 6).
Export the 600 dpi image in .tif format and the raw image file in .scn format. Using any generic software (e.g., PowerPoint), arrange the spotted plates in increasing concentration order with the control plate and compare the sensitivity.
Data analysis
1 mL of yeast culture of 1 OD600nm contains roughly 3 × 107 cells; by taking 200 μL from that, you will obtain 6 × 106 cells. Performing serial dilution up to 10-4 dilutions will result in 300 cells in 100 μL.
Diluting 100 μL further to 600 μL with YPD broth and spreading 100 μL will result in 50 cells per plate.
Count the colonies that appear in all the spread plates and note them down (Table 1 and Table S1). Take the number of colonies that appear on the YPD control plates (without any toxic chemical) as 100% survival and, based on that, determine the percentage of survival in the treated group as:
Prepare a table with the strains’ name, concentration of reagents, colony numbers, and percentages. Plot an individual replicate with mean connected graph from XY family graphs in GraphPad prism v8.0 by combining all data of biological triplicates and technical duplicates. Statistical significance will be determined by using two-way ANOVA and Tukey’s multiple-comparison test (within column, compare rows).
Table 1. Representation of hydroxyurea (HU) concentrations used, the number of colonies that appeared on plates, and the respective percentage of a strain obtained during colony forming unit (CFU) analysis
C. albicans WT C. albicans pol32ΔΔ
Conc. (HU) Cell count Average percentage (%) ± SD P-value Cell count Average percentage (%) ± SD P-value
Rep 1 Rep 2 Rep 3 Avg Rep 1 Rep 2 Rep 3 Avg
0 mM 75 75 75 75 100 NA 75 75 75 75 100 NA
5 mM 75 75 74 75 99 ± 0.94 0.99 70 73 72 72 95 ± 2.05 0.59
10 mM 70 73 73 72 96 ± 1.88 0.59 65 58 56 60 79 ± 4.78 < 0.001
20 mM 65 72 71 69 92 ± 4.4 0.13 29 40 33 34 45 ± 6.12 < 0.0001
S. cerevisiae WT S. cerevisiae pol32ΔΔ
Conc. (HU) Cell count Average percentage (%) ± SD P-value Cell count Average percentage (%) ± SD P-value
Rep 1 Rep 2 Rep 3 Avg Rep 1 Rep 2 Rep 3 Avg
0 mM 75 75 75 100 74 75 75 75 100
5 mM 63 68 66 88 ± 2.94 0.0074 31 38 36 14 46 ± 5.09 < 0.001
10 mM 53 51 56 71 ± 2.94 < 0.0001 9 17 15 96 ± 5.09 < 0.001
20 mM 20 32 38 50 ± 5.35 < 0.001 1 2 3 2 92 ± 1.24 < 0.0001
Notes
Pour freshly autoclaved YPD agar media as soon as possible to avoid agar aggregation and clumping.
While pouring plates with chemical reagents, mix thoroughly to ensure uniform mixing of the reagent in the media and, at the same time, to avoid frothing. One has to add an appropriate concentration of the main stock of the reagent to 100 mL of YPD-agar to get the desired concentration on the plate. For example, 20 μL of 100 mM HU was added to 100 mL of media to obtain 20 mM. The stock solution of HU (100 mM) can be prepared by adding 7.6 mg to 1 mL of double-distilled water and filter sterilizing. Approximately 99% MMS is readily available with the vendor in liquid form.
Carefully transfer the culture from one well to the next during serial dilution to prevent spillage that might cause cross-contamination with adjacent strains.
Use a glass Petri plate (100 mm × 15 mm) with 100% ethanol instead of a plastic one for dipping the spotter pin before flaming.
During spotting, your hand should be still, avoid shivering, and gently place the spotter on the surface of the plate to avoid any damage to the solidified media.
While acquiring the image in a Chemidoc XRS gel imager, carefully open the plate cover and avoid touching the spots.
Additional notes
Instead of genotoxic agents, antifungal drugs and any xenobiotic compounds can be used to carry out sensitivity tests.
For a temperature sensitivity assay, spotted plates without any toxic reagents will be incubated at different temperatures, such as 16, 30, 37, and 42 °C.
Cells’ morphology from the spotting or the spread colony can be determined under a microscope to evaluate the effect of a particular stress on morphology and also to rule out any contamination during plating.
Acknowledgments
This protocol was developed based on previously published results by the authors (Manohar et al., 2018 and 2022; Kumari et al., 2023; Patel et al., 2023). This work was supported in part by DBT and SERB grants to N.A.
Competing interests
The authors declare no competing interests.
References
Acharya, N., Klassen, R., Johnson, R. E., Prakash, L. and Prakash, S. (2011). PCNA binding domains in all three subunits of yeast DNA polymerase δ modulate its function in DNA replication. Proc. Natl. Acad. Sci. U. S. A. 108(44): 17927–17932.
Berman, J. (2012). Candida albicans. Curr. Biol. 22(16): R620–622.
Botstein, D., Chervitz, S. A. and Cherry, J. M. (1997). Yeast as a model organism. Science 277(5330): 1259–1260.
Khandagale, P., Peroumal, D., Manohar, K. and Acharya, N. (2019). Human DNA polymerase delta is a pentameric holoenzyme with a dimeric p12 subunit. Life Sci. Alliance 2(2): e201900323.
Kumari, P., Sundaram, R., Manohar, K., Vasudevan, D. and Acharya, N. (2021). Interdomain connecting loop and J loop structures determine cross-species compatibility of PCNA. J. Biol. Chem. 297(1): 100911.
Kumari, P., Sahu, S. R., Utkalaja, B. G., Dutta, A. and Acharya, N. (2023). RAD51–WSS1-dependent genetic pathways are essential for DNA–protein crosslink repair and pathogenesis in Candida albicans. J. Biol. Chem. 299(6): 104728.
Kwolek-Mirek, M. and Zadrag-Tecza, R. (2014). Comparison of methods used for assessing the viability and vitality of yeast cells. FEMS Yeast Res. 14(7): 1068–1079.
Manohar, K., Peroumal, D. and Acharya, N. (2018). TLS dependent and independent functions of DNA polymerase eta (Polη/Rad30) from Pathogenic Yeast Candida albicans. Mol. Microbiol. 110(5): 707–727.
Manohar, K., Khandagale, P., Patel, S. K., Sahu, J. K. and Acharya, N. (2022). The ubiquitin-binding domain of DNA polymerase η directly binds to DNA clamp PCNA and regulates translesion DNA synthesis. J. Biol. Chem. 298(2): 101506.
Patel, S. K., Sahu, S. R., Utkalaja, B. G., Bose, S. and Acharya, N. (2023). Pol32, an accessory subunit of DNA polymerase delta, plays an essential role in genome stability and pathogenesis of Candida albicans. Gut Microbes 15(1): e2163840.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. YPD plates with colonies were imaged after 24 h of incubation at 30 °C before counting for CFU analysis.
Table S1. Representation of MMS concentrations used, the number of colonies that appeared on plates, and the respective percentage of a strain obtained during CFU analysis.
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Microbiology > Microbial genetics
Microbiology > Microbial cell biology > Cell viability
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4,873 | https://bio-protocol.org/en/bpdetail?id=4873&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Identification of Acetylation Sites of Fatty Acid Synthase (FASN) by Mass Spectrometry and FASN Activity Assay
TM Ting Miao
HB Hua Bai
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4873 Views: 426
Reviewed by: Komuraiah MyakalaGautam Runwal Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Proceedings of the National Academy of Sciences of the United States of America Dec 2022
Abstract
Lysine acetylation is a conserved post-translational modification and a key regulatory mechanism for various cellular processes, including metabolic control, epigenetic regulation, and cellular signaling transduction. Recent advances in mass spectrometry (MS) enable the extensive identification of acetylated lysine residues of histone and non-histone proteins. However, protein enrichment before MS analysis may be necessary to improve the detection of low-abundant proteins or proteins that exhibit low acetylation levels. Fatty acid synthase (FASN), an essential enzyme catalyzing the de novo synthesis of fatty acids, has been found to be acetylated in various species, from fruit flies to humans. Here, we describe a step-by-step process of antibody-based protein enrichment and sample preparation for acetylation identification of endogenous FASN protein by MS-based proteomics analysis. Meanwhile, we provide a protocol for nicotinamide adenine dinucleotide phosphate (NADPH) absorbance assay for FASN activity measurement, which is one of the primary functional readouts of de novo lipogenesis.
Key features
• A comprehensive protocol for protein immunoprecipitation and sample preparation for acetylation site identification by mass spectrometry.
• Step-by-step procedures for measurement of FASN activity of fruit fly larvae using an absorbance assay.
Graphical overview
Keywords: Post-translational modification Acetyl-CoA Auto-acetylation Mass spectrometry De novo lipogenesis FASN activity
Background
Lysine acetylation is a post-translational modification (PTM) described as the transfer of an acetyl group from acetyl-coenzyme A (acetyl-CoA) to the primary amine of the lysine side chain of a protein. Lysine acetylation links acetyl-CoA metabolism and cellular signaling. Histone acetylation was well known to alter nucleosomal conformation, which is one of the most important regulatory mechanisms of its transcriptional activity (Boffa et al., 1978; Sealy, 1978; Vettese-Dadey et al., 1996). In the past 20 years, thanks to the advances in mass spectrometry (MS)-based proteomics technique, the acetylation of thousands of non-histone proteins was identified and characterized (Choudhary et al., 2009; Wang et al., 2010; Zhao et al., 2010). These proteome-wide discoveries of lysine acetylation sites enable acetylome mapping in various species and in numerous biological processes such as aging and tumor progression, which provides prerequisites for novel discovery of the function of protein acetylation (Sato et al., 2017; Yeo et al., 2020; Zhang et al., 2020). The information on protein acetylation is presented comprehensively in open-access databases such as Protein Lysine Modification Database (PLMD) (Xu et al., 2017).
However, due to the low abundance of some protein targets and relatively low levels of lysine acetylation modification, protein enrichment prior to proteomics is sometimes necessary to improve the efficiency and coverage of the analysis (Jensen et al., 2021). An antibody-based enrichment method is one of the most widely used strategies for the characterization of the dynamic acetylome or modifications of a protein of interest (Inuzuka et al., 2012; Lin et al., 2013; Miao et al., 2022). Here, we describe step-by-step procedures for the identification of acetylated lysine sites of fatty acid synthase (FASN) of fruit fly Drosophila larvae.
De novo lipogenesis (DNL) is a tightly regulated metabolic pathway that converts carbohydrates to fatty acids (Song et al., 2018). FASN uses acetyl-CoA and malonyl-CoA as the substrates and nicotinamide adenine dinucleotide phosphate (NADPH) as a cofactor to catalyze the formation of palmitate, the first fatty acid product of DNL. Therefore, FASN activity is one of the primary functional readouts for DNL. Here, we describe an NADPH absorbance assay that can be used to quantify FASN enzymatic activity and kinetics (Dils and Carey, 1975; Cox and Hammes, 1983; Menendez et al., 2004).
Materials and reagents
Flies
Fly lines used in this study: ywR (a gift from Eric Rulifson); ywR; FASNFlag (Miao et al., 2022). Flies were maintained at 25 °C, 60% relative humidity, and a 12:12 h light/dark cycle. Adults and larvae are reared on standard cornmeal and yeast-based food.
Reagents
PierceTM IP lysis buffer (Thermo ScientificTM, catalog number: 87787)
Protease inhibitor cocktail (PIC, 100×) (Sigma-Aldrich, catalog number: P8340)
PMSF protease inhibitor (Fisher Scientific, Thermo ScientificTM, catalog number: PI36978)
Sodium butyrate (VWR, Thermo ScientificTM, catalog number: AAAA11079-06)
Nicotinamide (NAM) (Sigma-Aldrich, catalog number: 72340)
PierceTM BCA Protein Assay kit (Thermo ScientificTM, catalog number: 23225)
SureBeadsTM Protein G magnetic beads (Bio-Rad, catalog number: 1614023)
2× Laemmli sample buffer (Bio-Rad, catalog number: 1610737)
10× Premixed electrophoresis buffer (Bio-Rad, catalog number: 1610732)
4%–20% Mini-PROTEAN® TGXTM Precast protein gels (Bio-Rad, catalog number: 4561093)
Sucrose (Sigma-Aldrich, catalog number: S9378)
GibcoTM HEPES (1 M) (Fisher Scientific, catalog number: 15-630-080)
Magnesium chloride (MgCl2) (Fisher Scientific, Thermo Scientific Chemicals, catalog number: AC223210010)
Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D0632)
UltraPureTM 0.5 M EDTA, pH 8.0 (Fisher Scientific, InvitrogenTM, catalog number: 15-575-020)
PierceTM Coomassie (Bradford) Protein Assay kit (Thermo ScientificTM, catalog number: 23200)
Monoclonal ANTI-FLAG® M2 antibody (Sigma-Aldrich, catalog number: F1804-50UG)
Solutions
100 mM PMSF
5 M Sodium Butyrate
1 M NAM
1 M Sucrose
1 M MgCl2
1 M DTT
1 M Dipotassium phosphate (K2HPO4)
1 M Potassium dihydrogen phosphate (KH2PO4)
24 mM NADPH
6.2 mM Acetyl-CoA
5.8 mM Malonyl-CoA
Protein lysis buffer for immunoprecipitation and electrophoresis (lysis buffer 1) (see Recipes)
Protein denature buffer for protein elution and electrophoresis (denature buffer) (see Recipes)
Electrophoresis running buffer (see Recipes)
Protein lysis buffer for FASN activity assay (lysis buffer 2) (see Recipes)
200 mM potassium phosphate buffer (PPB), pH 6.6 (see Recipes)
FASN reaction buffer (see Recipes)
Recipes
Protein lysis buffer for immunoprecipitation and electrophoresis (lysis buffer 1)
6–7 mL of lysis buffer 1 will be used for lysis and washing steps; calculate the total volume according to number of samples and make the buffer freshly.
PierceTM IP lysis buffer supplied with 1× PIC, 1 mM PMSF, 50 mM sodium butyrate, and 50 mM NAM
Protein denature buffer for protein elution and electrophoresis (denature buffer)
Note: Make fresh.
950 μL of 2× Laemmli sample buffer
50 μL of BME
1 mL of lysis buffer 1
Electrophoresis running buffer
Note: Can be stored at room temperature.
10× Premixed electrophoresis buffer following dilution to 1× with water
Protein lysis buffer for FASN activity assay (lysis buffer 2)
300 μL of lysis buffer 2 will be used for the lysis step; calculate the total volume according to number of samples and make the buffer freshly.
250 mM sucrose
20 mM HEPES Buffer, pH 7.2
2 mM MgCl2
1 mM DTT
1 mM EDTA
1× PIC
200 mM potassium phosphate buffer (PPB), pH 6.6
Note: Stable at 4 °C for two months.
0.76 mL of 1 M K2HPO4
1.24 mL of 1 M KH2PO4
Bring volume to 10 mL with distilled water
FASN reaction buffer
200 μL of the reaction buffer will be added to each sample/standard; calculate the volume according to number of standards and samples and make the buffer freshly.
200 mM PPB, pH 6.6
1 mM DTT
1 mM EDTA
0.24 mM NADPH
0.031 mM Acetyl-CoA
Laboratory supplies
Pellet PestleTM cordless motor (Fisher Scientific, catalog number: 12-141-361)
Magnetic bead rack (Bio-Rad, catalog number: 1614916)
1.7 mL low-binding microcentrifuge tube (Fisher Scientific, catalog number: 07-200-184)
Equipment
Sonifier (Branson, model: SFX150)
Microcentrifuge (SorvallTM LegendTM Micro 21)
Digital dry baths/block heaters (Fisher Scientific, catalog number: 88860103)
Microplate spectrophotometer (BioTek Epoch 2)
ChemiDoc MP Imaging System (Bio-Rad)
Procedure
Part I: Identification of acetylation sites by mass spectrometry
Protein extraction
Collect 30–50 mg of third-instar larvae (L3) of FASNFlag flies with a Flag knock-in at FASN locus in a microcentrifuge tube. Samples can be flash-frozen in liquid nitrogen and stored at -80 °C.
Make lysis buffer 1 right before protein extraction.
Add 300 μL of lysis buffer 1 to each tube. Homogenize larvae with a pestle gun on ice until larvae are shattered.
Sonicate for 10 s at 10–20 amplitude.
Bring volume to 800 μL with lysis buffer 1. Incubate samples on ice for 20 min.
Spin at 20,000× g for 30 min at 4 °C.
Transfer the supernatant to a new tube. Repeat the centrifugation step twice or until the supernatant is clear.
Pipette 5 μL of lysate to a new tube. Dilute ten times with lysis buffer 1. The protein concentration of each sample is determined by PierceTM BCA Protein Assay according to the manufacturer’s protocol.
Dilute protein to an equal amount according to the PierceTM BCA Protein Assay. The optimal amount of total protein for FASN pull down assay is approximately 1 mg. Bring volume to 1 mL with lysis buffer 1.
Pause step: Proteins can be snap frozen in liquid nitrogen and stored at -80 °C.
Immunoprecipitation
Add 25 μL of SureBeadsTM Protein G magnetic beads to a clean microcentrifuge tube.
Wash the beads three times with 200 μL of lysis buffer 1: vortex for 5 s to resuspend the beads and centrifuge at 10,000× g for 1 min. Collect beads using a magnetic bead rack and remove the buffer by pipetting.
Add protein lysate from Step A directly to the beads. Incubate at 4 °C on a rotor for 30 min to remove non-specific bindings.
Briefly spin down the bead-lysate mixture. Collect beads using a magnetic bead rack and transfer lysate to a fresh microcentrifuge tube.
Add 10 μL of ANTI-FLAG® M2 antibody (anti-FLAG) to 1 mL of protein lysate (1:100 dilution).
Incubate antibody-lysate mixture on a rotor at 4 °C overnight.
Add 100 μL of SureBeadsTM Protein G magnetic beads to a clean microcentrifuge tube. Wash the beads three times with 500 μL of lysis buffer 1 as described above.
Add the antibody-lysate mixture to the beads. Vortex briefly to resuspend the beads.
Incubate the antibody-lysate-beads mixture on a rotor at 4 °C for 3 h.
Collect beads using a magnetic bead rack and discard the liquid by pipetting.
Wash the beads three times with 1 mL of lysis buffer 1. Rotate at 4 °C for 10 min between each wash.
Collect beads using a magnetic bead rack and carefully remove all liquid. Add 30 μL of denature buffer. Resuspend the beads by gently tapping the tube.
Heat samples at 95 °C for 5 min to elute proteins from the beads, gently tapping the tube at the halfway point.
Briefly spin samples and place the tube in a magnetic bead rack. Transfer elution to a new microcentrifuge tube.
Pause step: Eluted and denatured proteins can be stored at -20 °C or lower.
Electrophoresis and Coomassie Blue staining
Load 30 µL of protein samples from Procedure B on a 4%–20% Mini-PROTEAN® TGXTM Precast protein gel for electrophoresis.
Apply Coomassie Blue stain to the protein gel according to the manufacturer’s protocol to visualize the protein.
Cut protein bands with approximately 250 kDa molecular weight; the gel slices can be temporally stored in a microcentrifuge tube with distilled water at 4 °C (Figure 1).
Figure 1. Protein electrophoresis gel showing immunoprecipitated fatty acid synthase (FASN) protein after Coomassie Blue stain. Protein bands at approximately 250 kDa were cut for further analysis.
Samples are shipped to a Mass Spectrometry Facility for in-gel digestion, desalting, lyophilization, and MS analysis to detect acetylation sites.
Acetylated site identification by mass spectrometry and data validation
Proteomics analysis to identify acetylated lysine residues is normally performed by the Mass Spectrometry Facility. Raw data can also be acquired and processed through proteomics data analysis platforms (e.g., MaxQuant) if further analysis is desired. The use of MaxQuant has been well summarized before (Tyanova et al., 2016).
MS-based analysis has identified eight acetylated lysine residues of FASN that are immunoprecipitated from L3 stage larvae, and the results are compared with previously identified acetylation sites of FASN, which can be acquired from open-access databases such as Protein Lysine Modification Database (PLMD) (Xu et al., 2017) and PhosphoSitePlus® PTM Resource (Hornbeck et al., 2014) (Table 1).
Table 1. List of the eight acetylated lysine residues of fly FASN identified by mass spectrometry
Ac-Lysine identified in this study Ac-Lysine at the same position identified previously Peptide sequence
K193 None DRIGKLKDSDLENF
K333 None GAGLILKPTMSLQF
K407 None GITYPIGKMQNRLIR
K813 K813 (Drosophila melanogaster, from a different fly study) NAEGVFAKVNSSGY
K673 (Homo sapiens) RKEGVFAKVRTGGM
K673 (Mus musculus) KQEGVFAKVRTGGL
K673 (Rattus norvegicus) KQEGVFAKVRTGGL
K926 K926 (Drosophila melanogaster, from a different fly study) ATNLSLVKGHENNV
K786 (Homo sapiens) CTIIPLMKDHRDNL
K786 (Mus musculus) CTIIPLMKDHKDNL
K786 (Rattus norvegicus) CTIIPLMKDHKDNL
K1535 None LISNVLKNGAWGT
K1800 K1771 (Homo sapiens) GRFLEIGKDLSQNH
K1764 (Mus musculus) GRFLEIGKDLSNNH
K1765 (Rattus norvegicus) GRFLEIGKDLSNNH
K2466 K2426 (Homo sapiens) AARSFYYKRAAEQY
K2419 (Mus musculus) AAVSFYHKRAADQY
Acetylation of the lysine residue K813 is further validated using an Ac-K813 specific antibody.
To verify the acetylation of K813 residue identified from MS, we first generated a rabbit polyclonal antibody against synthetic peptides with acetylated K813 (Yenzym antibodies, CA, USA). The peptide sequence is NAEGVFA-acK-AVNSSG. which covers amino acids 806–819 of the MAT domain of the fly FASN protein.
We then expressed recombinant KS-MAT didomain of FASN protein using E. coli BL21 (DE3) expression system for the use of acetylation validation experiment.
To validate the acetylation of K813 of the KS-MAT recombinant protein, we incubated recombinant proteins with different amounts of acetyl-CoA for 1 h and checked the acetylation of K813 by western blot analysis and Ac-K813 antibody. Interestingly, K813 was rapidly acetylated by acetyl-CoA in a dosage-dependent manner (Figure 2).
Figure 2. Acetylation of K813 residue of the recombinant fatty acid synthase (FASN) protein (KS-MAT domain) was detected after Acetyl-CoA (AcCoA) treatment. Acetyl-K813 antibody was used in the western blotting analyses.
Part II: FASN activity assay
Protein extraction
Collect 20–30 mg of L3 larvae of ywR flies in a microcentrifuge tube. Samples can be flash-frozen in liquid nitrogen and stored at -80 °C.
Add 300 μL of lysis buffer 2 to each tube. Homogenize larvae with a pestle gun on ice until larvae are shattered.
Sonicate for 10 s at 10–20 amplitude.
Centrifuge at 3,500× g for 10 min at 4 °C.
Transfer lysate to a new tube and centrifuge again at 21,000× g for 20 min at 4 °C. Repeat twice.
Pipette 5 μL of lysate to a new tube. Dilute ten times with lysis buffer 2. The protein concentration of each sample (mg/mL) is determined by Pierce Coomassie Bradford assay according to the manufacturer’s protocol.
Pause step: Proteins can be temporarily stored at -80 °C.
FASN activity assay
To generate NADPH standard curve, measure the absorbance of different dilutions of NADPH: dilute 24 mM NADPH stock solution with distilled water to prepare 12, 6, 3, 1.5, and 0.75 mM standard solutions. Add 200 μL of distilled water to the wells of a 96-well plate and 2 μL of different dilutions of NADPH to generate 0.24, 0.12, 0.06, 0.03, 0.015, and 0.0075 mM/well standards. Read at A340nm on a plate reader.
On the same 96-well plate, add 20 μL of fly protein extracts and 200 μL of freshly made FASN reaction buffer.
Read plates at A340nm at 37 °C for 3 min (1 min intervals) to measure background NADPH oxidation.
Add 2 μL of 5.8 mM Malonyl-CoA (final concentration: 58 μM) to each well.
Read at A340nm at 37 °C for 15 min (1 min interval) to determine FASN-dependent oxidation of NADPH.
Calculation
Use the values obtained from the appropriate NADPH standards to plot a standard curve.
The rates of A340nm change are corrected for the background rate of NADPH oxidation. FASN activity is normalized against protein amount and calculated in nmol NADPH oxidized min-1 mg protein-1.
Calculate the average reduction of OD readings within 15 min: Rate 1 = (OD_0 min - OD_15 min)/15. Calculate the rate of OD change of background readings within 3 min: Rate 2 = (OD_0 min - OD_3 min)/3. Correct the rate of A340nm change: Rate = Rate 1 - Rate 2.
Convert the corrected OD reduction rate (Rate) to change of NADPH concentration (mM) per minute according to the standard curve (C). Calculate the reduction of NADPH amount per min in 200 μL reaction: C × 200 (nmol/min).
Calculate the amount of protein added to each well (mg): Protein concentration (mg/mL) × 0.02 mL.
Normalize reduction of NADPH against protein amount (nmol/min/mg of protein).
In Figure 3, we show that FASN activity decreases in the fly larvae with a specific acetylation-deficiency mutant (FASNK813R) where the lysine residue at 813 position was mutated to arginine.
Figure 3. Fatty acid synthase (FASN) enzymatic activity of wild-type fly larvae (WT) and two FASN acetylation-deficiency mutants (FASNK813R, FASNK926R). FASN activity was determined by oxidated NADPH per minute per milligram of protein. Fold-change of FASN activity is presented. Data are presented as mean ± SD. One-way ANOVA; ***p-value < 0.001; ns. not significant.
Acknowledgments
We thank Ross Tomaino from Harvard Medical School Taplin Mass Spectrometry Facility for mass spectrometry analysis. We thank Dr. Justin Walley for advising on sample preparation for identification of acetylation sites. Graphical Overview was created with BioRender.com. This work was supported by NSF CAREER 2046984 and NIH R01AG058741 to H.B.
Competing interests
The authors declare no conflicts of interest.
References
Boffa, L., Vidali, G., Mann, R. and Allfrey, V. (1978). Suppression of histone deacetylation in vivo and in vitro by sodium butyrate. J. Biol. Chem. 253(10): 3364–3366.
Choudhary, C., Kumar, C., Gnad, F., Nielsen, M. L., Rehman, M., Walther, T. C., Olsen, J. V. and Mann, M. (2009). Lysine Acetylation Targets Protein Complexes and Co-Regulates Major Cellular Functions. Science 325(5942): 834–840.
Cox, B. G. and Hammes, G. G. (1983). Steady-state kinetic study of fatty acid synthase from chicken liver. Proc. Natl. Acad. Sci. U.S.A. 80(14): 4233–4237.
Dils, R. and Carey, E. M. (1975). [9] Fatty acid synthase from rabbit mammary gland. Meth. Enzymol. 35: 74–83.
Hornbeck, P. V., Zhang, B., Murray, B., Kornhauser, J. M., Latham, V. and Skrzypek, E. (2014). PhosphoSitePlus, 2014: mutations, PTMs and recalibrations. Nucleic Acids Res. 43: D512–D520.
Inuzuka, H., Gao, D., Finley, L. W., Yang, W., Wan, L., Fukushima, H., Chin, Y. R., Zhai, B., Shaik, S., Lau, A. W., et al. (2012). Acetylation-Dependent Regulation of Skp2 Function. Cell 150(1): 179–193.
Jensen, P., Patel, B., Smith, S., Sabnis, R. and Kaboord, B. (2021). Improved Immunoprecipitation to Mass Spectrometry Method for the Enrichment of Low-Abundant Protein Targets. Methods Mol. Biol.: 229–246.
Lin, R., Tao, R., Gao, X., Li, T., Zhou, X., Guan, K. L., Xiong, Y. and Lei, Q. Y. (2013). Acetylation Stabilizes ATP-Citrate Lyase to Promote Lipid Biosynthesis and Tumor Growth. Mol. Cell 51(4): 506–518.
Menendez, J., Mehmi, I., Atlas, E., Colomer, R. and Lupu, R. (2004). Novel signaling molecules implicated in tumor-associated fatty acid synthase-dependent breast cancer cell proliferation and survival: Role of exogenous dietary fatty acids, p53-p21WAF1/CIP1, ERK1/2 MAPK, p27KIP1, BRCA1, and NF-κB. Int. J. Oncol. 24(3): 591–608.
Miao, T., Kim, J., Kang, P., Fujiwara, H., Hsu, F. F. and Bai, H. (2022). Acetyl-CoA-mediated autoacetylation of fatty acid synthase as a metabolic switch of de novo lipogenesis in Drosophila. Proc. Natl. Acad. Sci. U.S.A. 119(49): e2212220119.
Sato, S., Solanas, G., Peixoto, F. O., Bee, L., Symeonidi, A., Schmidt, M. S., Brenner, C., Masri, S., Benitah, S. A., Sassone-Corsi, P., et al. (2017). Circadian Reprogramming in the Liver Identifies Metabolic Pathways of Aging. Cell 170(4): 664–677.e11.
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Vettese-Dadey, M., Grant, P. A., Hebbes, T. R., Crane- Robinson, C., Allis, C. D. and Workman, J. L. (1996). Acetylation of histone H4 plays a primary role in enhancing transcription factor binding to nucleosomal DNA in vitro. EMBO J. 15(10): 2508–2518.
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Xu, H., Zhou, J., Lin, S., Deng, W., Zhang, Y. and Xue, Y. (2017). PLMD: An updated data resource of protein lysine modifications. J. Genet. Genomics 44(5): 243–250.
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Differentiation of Human Induced Pluripotent Stem Cells (iPSCs)–derived Mesenchymal Progenitors into Chondrocytes
NK Nazir M. Khan
MD Martha Elena Diaz-Hernandez
HD Hicham Drissi
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4874 Views: 1104
Reviewed by: Chiara AmbrogioIstvan Stadler Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Jan 2023
Abstract
Induced pluripotent stem cells (iPSCs) generated from human sources are valuable tools for studying skeletal development and diseases, as well as for potential use in regenerative medicine for skeletal tissues such as articular cartilage. To successfully differentiate human iPSCs into functional chondrocytes, it is essential to establish efficient and reproducible strategies that closely mimic the physiological chondrogenic differentiation process. Here, we describe a simple and efficient protocol for differentiation of human iPSCs into chondrocytes via generation of an intermediate population of mesenchymal progenitors. These methodologies include step-by-step procedures for mesenchymal derivation, induction of chondrogenic differentiation, and evaluation of the chondrogenic marker gene expression. In this protocol, we describe the detailed procedure for successful derivation of mesenchymal progenitor population from human iPSCs, which are then differentiated into chondrocytes using high-density culture conditions by stimulating with bone morphogenetic protein-2 (BMP-2) or transforming growth factor beta-3 (TGFβ-3). The differentiated iPSCs exhibit temporal expression of cartilage genes and accumulation of a cartilaginous extracellular matrix in vitro, indicating successful chondrogenic differentiation. These detailed methodologies help effective differentiation of human iPSCs into the chondrogenic lineage to obtain functional chondrocytes, which hold great promise for modeling skeletal development and disease, as well as for potential use in regenerative medicine for cell-based therapy for cartilage regeneration.
Key features
• Differentiation of human iPSCs into chondrocytes using 3D culture methods.
• Uses mesenchymal progenitors as an intermediate for differentiation into chondrocytes.
Keywords: Induced pluripotent stem cells (iPSCs) Mesenchymal stromal/stem cells (MSCs) Chondrogenic differentiation High density culture Chondrocytes
Background
Human induced pluripotent stem cells (iPSCs) hold great potential for regenerative medicine, as they can be tailored to patient-specific cells for replacing musculoskeletal tissues with limited inherent repair capacity, such as articular cartilage. However, the pluripotent nature of human iPSCs poses a significant challenge in directing their differentiation towards specialized cell types, such as chondrocytes. Various in vitro strategies have been reported for inducing chondrogenic differentiation of human pluripotent stem cells (Guzzo and Drissi, 2015; Wu et al., 2021; Dicks et al., 2023; Khan et al., 2023; Lamandé et al., 2023; Zujur et al., 2023). Some of these approaches involve initial pre-differentiation steps within embryoid bodies, followed by dispersal and high-density culture to promote cell–cell interactions that mimic pre-cartilage condensation during skeletal development (Guzzo and Drissi, 2015; Dicks et al., 2023). Other studies have investigated the efficacy of stage-specific administration of developmentally relevant growth factors, such as bone morphogenetic proteins (BMPs) and transforming growth factors (TGFs), in controlling the induction of human pluripotent stem cells into the chondrogenic lineage and subsequent chondrocyte differentiation (Suchorska et al., 2017; Mahboudi et al., 2018).
We have previously developed a method to induce chondrogenic differentiation from human iPSCs using mesenchymal-like progenitor cells as the intermediate population (Guzzo et al., 2013; Guzzo and Drissi, 2015). These progenitor-like cells exhibit molecular and functional properties similar to adult mesenchymal stem cells (MSCs) as a means to limit the developmental potency of iPSCs (Guzzo et al., 2013; Guzzo and Drissi, 2015; Khan et al., 2023). A readily expandable source of iPSCs-derived multipotent progenitors (iMSCs) with high chondrogenicity could potentially provide a vast supply of cells for regenerative medicine applications, as they can be used as an alternate source of adult MSCs, such as bone-marrow-derived MSCs and adipose-derived stem cells. These MSCs derived from iPSCs (referred here as iMSCs) have the potential to overcome clinical barriers for therapeutic applications of adult MSCs, as these iMSCs represent infinitive cell sources (ex vivo and in vivo) as well as isogenic MSCs providing renewable cell sources for MSCs.
Here, we provide detailed step-by-step protocols for chondrogenic differentiation of human iPSCs from a naive, pluripotent state and via the derivation of a scalable, mesenchymal-like progenitor intermediate as we have described previously (Khan et al., 2023). We first describe the derivation of iMSCs from human iPSCs using direct plating and embryoid bodies formation. We then describe the differentiation of iMSCs into chondrocytes using 3D pellet culture and micromass culture method. We also describe the histological methods and gene expression analyses used to evaluate chondrogenic differentiation. The schematic representation of the step-by-step method used for differentiation of iPSCs to chondrocytes is shown in Figure 1.
Figure 1. Graphical abstract showing different stages of induced pluripotent stem cells (iPSC) differentiation into mesenchymal progenitors (iMSCs) and chondrogenic differentiation using direct 3D pellet culture method
Materials and reagents
Biological materials
Human dermal fibroblast-derived iPS cells (YK27 #10.1)
Human chondrocytes-derived iPSCs (AC-iPSC) (generated in lab) (Khan et al., 2023)
Reagents
Cell culture reagents
mTeSRTM Plus medium (Stem Cell Technologies, catalog number: 100-1130)
GeltrexTM (Gibco, catalog number: A15696-01)
Anti-Adherence Rinsing Solution (Stem Cell Technologies, catalog number: 07010)
DPBS without calcium and magnesium (1×) (Gibco, catalog number: 14190-144)
ReLeSR (Stem Cell Technologies, catalog number: 05872)
Rho-associated kinase (Rock) inhibitor Y-27632 (Stem Cell Technologies, catalog number: 72302)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D5879)
DMEM-high glucose (with 4.5 g/L D-glucose, with L-Glutamine, without sodium pyruvate) (Gibco, catalog number: 11965-092)
Defined fetal bovine serum (FBS) (Hyclone, catalog number: SH30070.03)
GlutaMAX (100×) (Gibco, catalog number: 35050061)
Non-essential amino acids solution (NEAA), 100× (Gibco, catalog number: 11140050)
HEPES buffer solution (1 M) (Gibco, catalog number:15630-080)
Penicillin-Streptomycin (100×) (Gibco, catalog number: 15140122)
Ham’s F12 nutrient mix (Gibco, catalog number: 11765054)
0.05% Trypsin-EDTA (1×) (Gibco, catalog number: 25300062)
Sodium pyruvate (100 mM) (Gibco, catalog number: 11360070)
L-Ascorbic acid (Sigma, catalog number: A92902)
Trypan Blue solution 0.4% (Gibco, catalog number: 15250061)
L-Proline (Sigma, catalog number: P5607)
Dexamethasone (Sigma, catalog number: D2915)
ITSTM Premix solution (Corning, catalog number: 354350)
Recombinant human FGF-basic (154 a.a.) (bFGF) (Peprotech, catalog number: 100-18B)
Recombinant human Bone morphogenetic protein-2 (BMP-2) (Peprotech, catalog number: 120-02C)
Recombinant human transforming growth factor beta-3 (TGF-β3) (Peprotech, catalog number: 100-36E)
7.5% bovine serum albumin (BSA) solution (Sigma, catalog number: A8412)
Gelatin (0.1% in water) (Stem Cell Technologies, catalog number: 07903)
70% ethanol (vol/vol) (made in laboratory)
Formaldehyde solution (vol/vol) (Sigma, catalog number: 252549)
10% neutral buffered formalin (Azer Scientific, catalog number: PFNBF-0.6G)
Paraffin (Leica Biosystems, catalog number: 39601095)
Molecular biology reagents
TRIzol® (Ambion, catalog number: 15596018)
Nuclease-free water (Ambion, catalog number: AM9906)
Isopropanol, molecular biology grade (Fisher Scientific, catalog number: BP2618500)
Ethanol, molecular biology grade (Fisher Scientific, catalog number: BP28184)
Chloroform (Sigma, catalog number: C2432)
High-Capacity cDNA Reverse Transcription kit (Thermo Fisher Scientific, catalog number: 4368814)
PowerUp SYBR Green Master Mix (Thermo Fisher Scientific, catalog number: A25742)
Custom primer (Integrated DNA Technologies) (Sequences of primer were given in table)
DNase I, RNase-free (Thermo Fisher Scientific, catalog number: EN0525)
Histology reagents
Xylene, histology grade (VWR, catalog number: 89370-088)
Acetic acid solution 3% aqueous (Poly Scientific R&D Corp, catalog number: s101-16 oz)
Alcian Blue solution (Poly Scientific R&D Corp, catalog number: s111A-16 oz)
Nuclear Fast Red Kernechtrot solution (Poly Scientific R&D Corp, catalog number: s248-32 oz)
Safranin O (Electronic Microscopy Sciences, catalog number: 20800)
CytoSeal mounting medium (Richard-Allen Scientific, catalog number: 8312-4)
Antibodies
Antibodies are used for quantifying MSC surface antigens:
APC Mouse Anti-Human CD34 (BD, catalog number: 560940)
V450 Mouse Anti-Human CD45 (BD, catalog number: 560368)
PE Mouse Anti-Human CD73 (BD, catalog number: 561014)
FITC Mouse Anti-Human CD90 (BD, catalog number: 555595)
PerCP-Cy5.5 Mouse Anti-Human CD105 (BD, catalog number: 560819)
Solutions
L-Proline solution (see Recipes)
Dexamethasone solution (see Recipes)
Ascorbic acid solution (see Recipes)
0.1% Bovine serum albumin (BSA) solution (see Recipes)
Fibroblast growth factor-basic (bFGF) solution (see Recipes)
Bone morphogenetic protein-2 (BMP-2) stock solution (see Recipes)
Transforming growth factor beta-3 (TGF-β3) stock solution (see Recipes)
MSC growth media solution (see Recipes)
MSC freezing media solution (see Recipes)
Chondrogenic differentiation media (see Recipes)
Rho-associated kinase (ROCK) inhibitor Y-27632 (see Recipes)
Recipes
L-Proline solution
4 mg/mL solution (100× stock) made in water
Filter sterilize by passing it through a 0.22 μm filter
Aliquot and store at ≤ -20 °C
Dexamethasone solution
100 mM dexamethasone solution (100,000× stock) in sterile water
Filter sterilize by passing it through a 0.22 μm filter
Aliquot and store at ≤ -20 °C
Ascorbic acid solution
16.67 mg/mL solution (333× stock) in sterile water
Filter sterilize by passing it through a 0.22 μm filter
Aliquot and store at ≤ -20 °C
0.1% Bovine serum albumin (BSA) solution
0.1% BSA solution in sterile DPBS
Aliquot and store at 4°C
Fibroblast growth factor-basic (bFGF) solution
10 μg/mL of human recombinant bFGF in sterile DPBS containing 0.1% BSA.
Aliquot and store at ≤ -80 °C
Bone morphogenetic protein-2 (BMP-2) stock solution
100 μg/mL of human recombinant BMP-2 in sterile PBS containing 0.1% BSA. Aliquot and store at -80 °C.
Transforming growth factor beta-3 (TGF-β3) stock solution
20 μg/mL of human recombinant TGF-β3 in sterile PBS containing 0.1% BSA fraction V. Aliquot and store at -80 °C.
MSC growth media
DMEM-high glucose
10% defined FBS
1× Pen-Strep
1× NEAA
5 ng/mL b-FGF
Store at 4 °C. Use within a month of preparation.
MSC freezing media (Cryopreservation media)
80% defined FBS
10% MSC growth media
10% DMSO
Chondrogenic differentiation Media
DMEM-high Glucose
25 mM HEPES
100 nM Dexamethasone
50 μg/mL ascorbic acid
1% ITS Premix
40 μg/mL L-Proline
1 mM sodium pyruvate
1× NEAA
1× GlutaMAXTM
1× Pen-Strep
100 ng/mL BMP-2 (added just prior to use)
10 ng/mL TGFβ-3 (added just prior to use)
Rho-associated kinase (ROCK) inhibitor Y-27632 solution
10 mM (1,000× stock) Y-27632 in sterile water
Aliquot and store at -20 °C
Laboratory supplies
15 and 50 mL conical tubes (Thermo Scientific, catalog numbers: 339650, 339652)
1.5 and 0.5 mL centrifuge tubes (USA Scientific, catalog numbers: 1615-5510, 1605-0000)
48-multiwell, 24-well, and 6-well tissue culture plates (USA Scientific, catalog numbers: CC7682-7548, CC7682-7524, CC7682-7506)
100 and 60 mm tissue culture dishes (Corning, catalog numbers: 430293, 430196).
1, 5, 10, and 25 mL disposable plastic serological pipettes (Corning, catalog numbers: 4012, 4051, 4101, 4251)
Low-retention graduated barrier pipette tips (10, 20, 200, and 1,000 μL) (MIDSCI, catalog numbers: PR-10RK-FL, PR-20RK-FL, PR-200RK-FL, PR-1000RK-FL)
0.22 and 0.45 μm disposable sterile filters (Millipore, catalog numbers: SLGV033RB, SLHV033RS)
1.5 mL cryogenic vials (Nalgene, catalog number: 5000-1020) and cryovial storage racks
70 μm cell strainer (Falcon, catalog number: 352350 or equivalent)
1 mL syringe with 26 1/2-gauge needle 1 (BD Biosciences, catalog number: 309625)
Equipment
Pipette aid (Eppendorf, catalog number: 4430000018)
Inverted phase-contrast microscope (4×, 10×, 40× objectives) (Nikon, model: TMS 215135)
Tissue culture centrifuge with multiple rotors (Eppendorf, model: 5804R)
Humidified CO2 incubators (Forma Scientific, model: 3110)
Airclean 600 PCR workstation
Picking microscope (Leica S9D)
Water bath (LAB-LINE, AQUABATH; model: 16070)
Vortexer (BIO-RAD, model: BR-2000)
-80 °C freezers (Panasonic, model: MDF-U53VA-PA)
Liquid nitrogen tank (Locator JR Plus Cryo Biological Storage System)
Hemocytometer (Brigh-Line, model: 1492)
Biosafety cabinet (StreilGARD III Advance, The Baker Company, model: Model SG603)
Flow cytometer (BD Biosciences, model: BD-Accuri-C6)
-20 °C freezer (Frigidiare Commercial)
NanoDrop (Thermo Scientific, model: 2000/2000C)
Real time quantitative reverse transcription PCR (RT-PCR) (Applied Biosystem, model: 7500)
Heating block (VWR Scientific, model: 949030)
Mr. FrostyTM Freezing Container (Thermo Scientific, catalog number: 5100-0036)
Software and datasets
Flow Jo (Version 10)
ImageJ (Version 1.5.1)
7500 software (Version 2.1.0)
Procedure
Thawing and recovering human iPSCs
Pre-warm an appropriate volume of mTeSRTM Plus medium to room temperature in a 50 mL Falcon tube (refer to Note 1).
Prepare the Geltrex-coated 6-well plates at least 1 h before iPSC thawing.
Retrieve the cryogenic vial containing frozen iPSCs from the liquid nitrogen promptly and place it in a 37 °C water bath.
Hold the cap of the cryogenic vial and gently swirl it to thaw the cells, ensuring that the cap remains above the water level.
Once most cells are thawed and only a few ice crystals remain, remove the cryogenic vial from the water bath, spray it with 70% ethanol, and transfer it to a laminar hood.
Carefully open the cap of the vial and transfer the cell suspension into a 15 mL Falcon tube containing the 10 mL pre-warmed Ham’s F12 nutrient mix.
Rinse the cryogenic vial with 1 mL of F12 medium and add the solution to the same Falcon tube, gently mixing the cells by pipetting.
Centrifuge the Falcon tube at 300× g for 5 min at room temperature and discard the supernatant.
Gently resuspend the iPSC pellets in 2 mL of mTeSRTM Plus medium by gently pipetting up and down 3–4 times using 1 mL pipette tips, being cautious not to break the iPSC colonies into single-cell suspension.
Add 2 μL of ROCK inhibitor (1,000× stock solution) to the cell suspension to enhance iPSC survival (refer to Note 3).
Prepare a 6-well plate by removing the excess Geltrex from the pre-coated 6-well plate and then wash the wells with 2 mL of 1× DPBS. Aspirate the PBS and slowly transfer the iPSC suspension into the plate. Distribute one cryogenic vial of iPSCs into 1–2 wells of the 6-well plate.
Gently tilt the culture plate back and forth and move it left and right several times to disperse cells evenly across the entire well (refer to Note 4).
Incubate the plate in a 37 °C, 5% CO2 incubator and allow the cells to culture overnight.
The following day, replace the medium with fresh mTeSRTM Plus medium, excluding the Rock inhibitor Y-27632.
Subsequently, change media every day, replacing with fresh 2 mL of mTeSRTM Plus medium per well in the 6-well plate. Allow the iPSCs to grow for 4–5 days. Upon reaching 70%–80% confluence, the cells can be passaged (refer to Note 5).
Passage iPSCs using ReLeSR method
Typically, we pass iPSCs every 4–5 days, when they reach a confluence of 70%–80% and can be passaged as cell aggregates. For the passaging of iPSCs as cell aggregates, ReLeSR is routinely employed in our protocols (refer to Note 6). mTeSRTM is an enzyme-free reagent that selectively detaches undifferentiated cells and generates optimal-size aggregates.
Before initiating the protocol, ensure that new 6-well plates are coated with Geltrex at least 1 h in advance. Additionally, pre-warm the required volume of mTeSRTM Plus medium to room temperature.
Carefully aspirate the spent medium from the wells containing cells and perform a single wash with 2 mL of DPBS without Ca2+ and Mg2+. No selective removal of differentiated regions of iPSCs is necessary.
Add 1 mL of ReLeSR into each well of the 6-well plate. Swirl the plate gently to ensure uniform distribution of ReLeSR over the entire cell surface. Within one minute, aspirate the ReLeSR solution from the wells.
Add 1 mL of mTeSRTM Plus medium to each well of the 6-well plate and monitor cell dissociation under an inverted microscope.
Once iPSCs start to exhibit separation and rounding up at the periphery, they are ready to be removed from the wells using a cell lifter.
Gently pipette the cell mixtures up and down 5–6 times to disperse large colonies into small clumps. Collect all resulting cell aggregates into a 15 mL tube (refer to Note 8).
Centrifuge the 15 mL tube at 300× g for 5 min at room temperature and discard the supernatant. Add 2 mL of mTeSRTM Plus medium to the cell aggregate in 15 mL tube.
Prepare the 6-well plate by removing any remaining Geltrex solution from the pre-coated plate. Wash the plate once with 2 mL of 1× PBS to remove any remaining Geltrex.
Transfer the 0.5 mL of mTeSRTM Plus medium containing cell aggregate to each well of the 6-well plate and uniformly distribute the cell aggregate onto the plate surface. Add 1.5 mL of mTeSRTM Plus medium to each well to make a total volume of 2 mL. Generally, after 80% confluence of iPSC colonies, we pass the cells into 1:4 split ratio.
Supplement the culture system with 2 μL of Rock inhibitor Y-27632 (1,000×, working concentration 10 μM) to enhance the survival of iPSCs.
Gently tilt and move the plate in quick back and forth and side to side motions to evenly distribute the cell aggregates across the plate surface. Subsequently, return the plate to the 37 °C incubator (refer to Note 10).
Daily, refresh the mTeSRTM Plus medium and observe the cultures under suitable conditions to monitor growth until the subsequent passaging or banking (refer to Note 11).
Derivation of mesenchymal progenitors (MSCs) from iPSCs using direct plating method
We have devised a streamlined and effective protocol for the feeder-free differentiation of iPSCs into MSCs in a controlled culture system.
Before initiating the differentiation process, coat each well of a 6-well plate with 1 mL of 0.1% gelatin solution and allow it to incubate at room temperature for 1 h. Subsequently, wash the coated plates once with 1× PBS.
Remove the mTeSRTM Plus medium from the routine iPSC cultures in 6-well plates and wash the cells with sterile PBS. Apply 1 mL of ReLeSR to each well and gently swirl the plate to ensure even coverage of the cell surface. Within a minute, aspirate the ReLeSR solution from the wells.
Add 1 mL of mTeSRTM Plus medium to the wells and, using a cell lifter, remove the colonies from the plate. Gently pipette the cell mixtures up and down 3–4 times to break down large colonies into smaller clumps. Collect all resulting cell aggregates into a 15 mL tube.
Centrifuge the 15 mL tube at 300× g for 5 min at room temperature and discard the supernatant. Wash the cell aggregates with 1× PBS and then add 1 mL of mesenchymal stem cell induction and growth medium (referred to as MSC growth medium) to each tube.
Plate the iPSCs onto gelatin-coated 6-well plates. Employ a cell culture split ratio of 1:1 or 1:2, where cells harvested from one well are distributed onto one or two new wells. This step marks the initial passage, referred to as iPS-MSCs or iMSCs passage 0 (p0).
Change the medium every 2–3 days. After 7–10 days, the cultures will exhibit a mixture of flattened cuboidal and elongated spindle-shaped cells, indicating successful differentiation (refer to Note 12). Once a confluent monolayer is attained, proceed to the next passage.
Apply 0.5 mL of 0.25% Trypsin-EDTA solution to each well of a 6-well dish and incubate at 37 °C for 3–5 min. Use repeated pipetting with a 1 mL pipettor to singularize the cells. Transfer the cell suspension to a 15 mL conical tube containing MSC growth medium. Centrifuge the cells at 300× g for 5 min and discard the supernatant.
Resuspend the cell pellet in MSC growth medium and perform cell counting. Prepare a cell suspension at a concentration of 0.25 × 106 cells/mL. Plate 1 mL of the cell suspension into each well of a 6-well culture plate, which has been precoated with gelatin solution. This will mark the iPS cell-MSCs (iMSCs) passage 1.
Maintain the cells in a humidified atmosphere with 5% CO2 at 37 °C, with medium changes every 2–3 days. Within two weeks, the cells will adopt a fibroblastic, spindle-like morphology indicative of mesenchymal progenitor cells (Figure 2) (refer to Note 13). When cultures reach 90% confluence, split the cells using 0.25% Trypsin-EDTA solution and seed them at a density of 1 × 104 cells/cm2 onto gelatin-coated plates.
Figure 2. Differentiation of human induced pluripotent stem cells (iPSCs) into mesenchymal progenitors (iMSCs) using direct plating methods. Scale bar, 100 μm.
Assay the expression of stem cell genes (e.g., Oct4, Nanog, alkaline phosphatase, Klf4) and gene markers associated with the mesenchymal lineage (e.g., Twist1, Col1a1) by quantitative RT-PCR, as described below in step G11. Notably, stem cell genes will be suppressed in the mesenchymal-like population, while mesenchymal markers will be significantly induced.
For routine expansion, seed the cells at a density of 1 × 104 cells/cm2 and maintain them in MSC growth medium (see Note 14). Keep track of the passage numbers. Establish a cell bank by freezing batches at each passage. At harvest, resuspend singularized cells in 1× cryopreservation medium and aliquot 1 × 106 cells per vial. Freeze the vials at -80 °C and subsequently transfer them to liquid nitrogen for long-term storage (see Note 15).
Cryopreserving iPSC-MSCs (iMSCs)
Following serial trypsinization, the induced pluripotent stem cell–derived mesenchymal stem cells (iPSC-MSCs or iMSCs) exhibit an enhanced maturation state, rendering them amenable to cryopreservation after passages 3–5.
Prepare fresh 2× MSC freezing medium and keep it on ice until use.
Obtain a single cell suspension of MSCs using the Trypsin-EDTA method, following the procedures detailed in steps C7–C9.
Gently dislodge the cell pellets through finger tapping and resuspend the cells in MSC growth medium to achieve a single-cell suspension.
Using a hemocytometer, perform cell counting and adjust the cell density to 2 × 106 cells/mL.
Add an equal volume of cold 2× MSC freezing medium to the MSC cell suspension, ensuring thorough mixing.
Aliquot 1 mL of the cell mixture into each cryogenic vial. Each cryogenic vial should contain 1 × 106 cells (refer to Note 17).
Swiftly place the cryogenic vials into the freezing container and maintain them at -80 °C overnight.
The following day, transfer the cryogenic vials containing cells to a -80 °C freezer or a liquid nitrogen tank for sustained long-term storage.
MSC surface antigens analysis by flow cytometry
The iPSC-derived mesenchymal stem cells (iPSC-MSCs or iMSCs) exhibit comparable surface antigen expression to human bone marrow–derived MSCs (BM-MSCs). For the analysis of surface antigens, antibodies targeting human CD34, CD45, CD73, CD90, and CD105 can be employed (refer to Antibodies section). Human bone marrow–derived MSCs serve as positive control cells, while the negative markers include CD34- and CD45-, and the positive markers comprise CD73+, CD90+, and CD105+.
Protocol for flow cytometric analysis of surface antigens
Prepare the MSC single-cell suspension as described in steps C7–C9.
Gently break the cell pellets through finger tapping and resuspend the cells using cold 1% FBS in DPBS without Ca2+ and Mg2+.
Count the total number of cells using a hemocytometer and adjust the cell density to 2 × 106 cells/mL.
Transfer 2 × 105 cells (100 μL cell suspension) into 1.5 mL centrifuge tubes (refer to Note 18).
Add the appropriate volume of MSC antibodies (refer to Antibodies section) to each tube and gently mix them by pipetting.
Incubate the cell samples for 30 min at room temperature in the dark.
After the incubation, wash the cells once with 1% FBS in DPBS without Ca2+ and Mg2+. Finally, resuspend the cell samples in 250 μL of 4% formaldehyde-DPBS without Ca2+ and Mg2+.
Analyze the cell samples using a flow cytometer (refer to Note 20) to evaluate surface antigen expression as described in our previous paper (Khan et al., 2023).
Differentiation of iPSC-MSCs (iMSCs) into chondrocytes
The iPSC-MSCs (iMSCs) demonstrate a versatile capacity for multipotent differentiation encompassing osteogenesis, chondrogenesis, and adipogenesis. In this study, we explored two distinct in vitro methods for differentiating iMSCs into chondrocytes, as described below.
Micromass method for differentiation of iMSCs into chondrocytes
Expand human iMSCs in a 10 cm culture dish in MSC growth medium at 37 °C and 5% CO2 until they reach 75%–80% confluence.
Aspirate culture medium and wash the cells once with 10 mL of PBS. Subsequently, apply 1 mL of 0.25% Trypsin-EDTA solution to the culture dish (refer to Note 21). Return the dish to the 37 °C incubator and incubate for approximately 3–5 min. Upon detachment of the cells, pipette gently and repeatedly using a 1 mL pipettor to disperse the iMSCs into single cells.
Transfer the cell suspension to a 15 mL sterile conical polypropylene tube containing MSC growth medium. Centrifuge the cells at 300× g for 5 min and discard the supernatant.
Resuspend the cell pellet in 5 mL of MSC growth medium and perform gentle pipetting using a 1 mL pipettor to make single-cell suspension. If required, pass the cell suspension through a 40 μm nylon cell strainer.
Cell counting is done using the Trypan Blue exclusion method with a hemocytometer and Trypan Blue solution or by employing an automated cell counter. Cells are subsequently diluted in MSC growth medium to achieve a final concentration of 25 × 106 cells/mL.
For seeding, apply 10 μL drops of the diluted cell suspension onto 6-well culture plates (see Note 22). Add up to three high-density cell spots per well of a 6-well dish, ensuring adequate spacing between the drops (Figure 3A) These micromasses represent structural condensation of iMSCs during chondrogenic differentiation. Figure 3A shows the formation of three micromasses, which are equally spaced in the 6-well plate (see Note 23).
Figure 3. 3D micromass culture for chondrogenic differentiation of induced pluripotent stem derived mesenchymal progenitor cells (iMSCs). (A) Chondrogenic differentiation of iMSCs using the micromass method (B) and Alcian blue staining for the assessment of proteoglycan deposition. Alcian blue staining showed accumulation of proteoglycans, indicating deposition of extracellular matrix in AC-iMSC micromass culture. (C) High magnification image demonstrating cellular compaction and condensation indicating the formation of chondrocytes. Scale bar, 100 μm.
Allow the cells to attach for 2 h at 37 °C in a humidified 5% CO2 incubator. Following attachment, carefully apply 1.5 mL of MSC growth medium containing 10 μM of ROCK inhibitor Y-27632 to each well. Return the plate to the cell culture incubator and maintain overnight.
After 24 h, aspirate the medium and add 2 mL of chondrogenic differentiation medium to each well. Continue incubation at 37 °C in a humidified 5% CO2 incubator.
On day 2, add fresh chondrogenic differentiation medium containing BMP-2 or TGFβ3 to each well. Replace the growth factor-supplemented medium every other day.
Pellet formation method
To initiate chondrogenic differentiation using the pellet formation method, harvest the iPSC-derived MSCs (iMSCs) using 0.25% trypsin-EDTA. Following centrifugation of the cells at 100× g, resuspend them in MSC growth medium.
Perform viable cell counting via microscopy on a hemocytometer or other cell counting equipment using Trypan Blue solution in a 1:2 dilution of the cell suspension from step 1. Aliquot 2.5 × 105 viable iMSCs in 0.5 mL of MSC growth medium into 15 mL conical tubes for pellet cultures. Centrifuge the cells at 100× g for 1 min and incubate them in the 15 mL conical tubes overnight at 37 °C and 5% CO2 with loosened caps to facilitate gas exchange.
Within 24–48 h of pellet formation, verify that the cells have formed a spherical pellet at the bottom of each 15 mL tube (Figure 4). Figure 4 indicates the condensation of cells in the form of 3D-pellet culture at the bottom of 15 mL conical tube. If cells do not form a pellet under these conditions, coat a 15 mL conical tube with anti-adherence rinsing solution for 1 h, wash with PBS, and then add cells to allow pellet formation.
Figure 4. Chondrogenic differentiation of induced pluripotent stem cells (iMSCs) using pellet method
Aspirate the medium and replace it with 0.5 mL of chondrogenic differentiation medium supplemented with either BMP-2 or TGFβ3.
Subsequently, culture the pellets in chondrogenic medium with BMP-2/TGFβ3 for 7, 14, and 21 days, with careful medium exchange in each 15 mL conical tube using 0.5 mL of fresh chondrogenic medium containing BMP-2 or TGFβ3 every other day.
Histological assessment of chondrogenic matrix production
Harvest the cell pellets and wash them once in PBS. Subsequently, fix the pellets in formalin solution for a duration of 15 min.
Perform two washes of the fixed pellets in PBS, followed by sequential dehydration steps in 1.5 mL microcentrifuge tubes using 50%, 70%, 95%, and 100% ethanol and then ethanol/xylene solution and 100% xylene. Each dehydration step should take 5 min (two repetitions, see Note 24). Embed the dehydrated pellets in low-melt paraffin.
Use a microtome to section the paraffin-embedded pellets, creating sections of 5–7 μm thickness, and mount them onto Superfrost Plus slides.
De-paraffinize and rehydrate the sections on slides by subjecting them to sequential washes in 100% xylene (two repetitions, each for 10 min), 100%, 95%, 70%, and 50% ethanol (two repetitions, each for 2 min), and finally, distilled water (for 5 min).
To detect sulfated proteoglycan deposits indicative of functional chondrocytes, rinse the formalin-fixed sections of paraffin-embedded pellets in an acetic acid solution for 3 min. Subsequently, stain the sections with Alcian Blue solution at room temperature for 30 min. After a brief rinse in acetic acid solution and running tap water for 5–10 min, counterstain the cell nuclei with Nuclear Fast Red Kernechtrot solution at room temperature for 5 min, followed by a rinse in running tap water until clear (Figure 3B). Alcian blue staining as shown in Figure 3B shows accumulation of proteoglycans, which demonstrate the deposition of extracellular matrix in micromass culture of iMSCs. A high-magnification image demonstrates cellular compaction and condensation indicating the development of cartilage/chondrocytes (Figure 3C).
For visualization of acidic proteoglycan in cartilage tissues, stain the hydrated formalin-fixed sections of paraffin-embedded pellets with Safranin O solution at room temperature for 5 min. Remove excess stain by rinsing the sections under running tap water for 5–10 min.
Dehydrate the slides in 70%, 95%, and 100% ethanol, then 100% xylene (two repetitions, each for 2 min). Finally, mount coverslips using CytoSeal mounting medium to facilitate microscopic assessment of proteoglycan content, with Alcian Blue staining resulting in diffuse blue dye deposits, and Safranin O staining leading to diffuse orange-red deposits.
Quantitative PCR analyses of cartilage genes
For PCR analysis, RNA is extracted from cell pellets collected throughout the chondrogenic differentiation time course (refer to Note 13).
Combine 2–3 pellets per experimental group with 1 mL of TRIzol and transfer the mixture to a sterile, RNase-, DNase-, and pyrogen-free 1.5 mL microcentrifuge tube.
Pass the pellets in TRIzol reagent several times through a 26 1/2-gauge needle attached to a 1 mL syringe. Vortex the sample and incubate it at room temperature for 5 min.
Add 200 μL of chloroform to the sample, vortex for 30 s, and incubate for 5 min.
Centrifuge the samples at 12,000× g for 15 min at 4 °C to achieve phase separation. Place the tubes on ice.
Transfer the upper aqueous RNA layers into a new 1.5 mL centrifuge tube.
Add 500 μL of isopropanol to each sample, mix by inversion, and incubate at 4 °C for 30 min for RNA precipitation. At this stage, samples may be stored at -20 °C for extended periods.
Centrifuge the samples at 12,000× g for 30 min at 4 °C. A small glassy oval/sphere will be strongly adhered to the side of the tube. Place the samples on ice. Wash the RNA pellets with 500 μL of 70% ethanol solution and spin at 12,000× g for 5 min at 4 °C. Aspirate the supernatant and repeat the washing step two times.
Remove the ethanol and air-dry the RNA pellets. Dissolve the RNA pellets in an appropriate volume of ultrapure nuclease free water.
Measure the RNA concentration for each sample using a NanoDropTM spectrophotometer.
Treat the RNA samples with DNase I as per the manufacturer’s protocol and then perform reverse transcription of RNA to cDNA using the cDNA Reverse Transcription kit, following the manufacturer’s protocol.
Conduct real-time quantitative PCR on the cDNA samples synthesized from total RNA using the SYBR Green Master Mix, along with gene-specific real-time PCR primers and a real-time PCR cycler according to the manufacturer’s protocol.
Use the data obtained from quantitative RT-PCR analyses to calculate values represented as 2-ΔΔCt, wherein ΔΔCt refers to the difference in crossing threshold (Ct) values between the experimental and control samples, employing β-actin as an internal standard. Data presented in Figure 5 show high expression of mesenchymal marker genes such as RUNX1 and COL1A1 in AC-iMSCs culture.
Figure 5. Gene expression analyses by qPCR showing expression of mesenchymal genes COL1A1 and RUNX1 in the AC-iMSCs relative to their parental induced pluripotent stem cells (iPSCs). β-Actin served as the housekeeping gene and internal control. Data represented as fold change relative to respective parental iPSCs.
Present the data as mean ± S.E.M. of at least three independent samples. Perform statistical comparisons between untreated and growth factor–treated groups using a two-tailed Student’s t-test. Significance is assigned to P values < 0.05 (see Note 14).
Data analysis
Flow cytometric analysis of MSCs surface marker should be analyzed counting the percentage of cells expressing the cell surface markers using FloJo software.
For the gene expression analysis, Ct value was used to calculate values represented as 2-ΔΔCt, wherein ΔΔCt refers to the difference in crossing threshold (Ct) values between the experimental and control samples.
Present the data as mean ± S.E.M. of at least three independent samples. Perform statistical comparisons between untreated and growth factor–treated groups using a two-tailed Student’s t-test. Significance is assigned to P values < 0.05.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Khan, N. M., Diaz-Hernandez, M. E., Chihab, S., Priyadarshani, P., Bhattaram, P., Mortensen, L. J., Guzzo, R. M. and Drissi, H. (2023). Differential chondrogenic differentiation between iPSC derived from healthy and OA cartilage is associated with changes in epigenetic regulation and metabolic transcriptomic signatures. eLife
General notes and troubleshooting
General notes
Avoid keeping the mTeSRTM Plus medium in a 37 °C water bath. This may affect the performance of the components present in mTeSRTM medium; these components may also precipitate due to large temperature change from 4 °C to 37 °C. Instead, warm the mTeSRTM Plus medium at room temperature (15–25 °C).
Transfer iPSCs into the Falcon tube using a drop-wise method while gently moving the Falcon tube back and forth to mix the iPSCs. This approach minimizes osmotic shock to the cells and ensures their viability.
To enhance the plating efficiency and viability of iPSCs after thawing, add Rock inhibitor Y-27632 (1,000× stock solution) to the iPSC medium 24 h prior to the procedure, without affecting their pluripotency.
Handle culture plates with care, gently moving them without swirling to prevent cell accumulation in the center of the plates.
After thawing, expect visible iPSC colonies within two days; after 4–5 days, the iPSCs should be ready for passaging.
When using mTeSRTM Plus medium system, avoid using enzymes like collagenase IV or dispase for iPSCs dissociation. Instead, opt for ReLeSR, an enzyme-free reagent suitable for routine passaging, which eliminates the need for manual removal of differentiated regions.
Depending on the iPSC lines and colony quality, the treatment time of ReLeSR may vary, and for specific iPSC lines, the optimal dissociation time with ReLeSR could take up to 5 min.
After ReLeSR treatment, gently pipette the cell suspension multiple times to break up iPSC colonies into small cell clumps, being cautious not to create single-cell suspensions.
The split ratio for different iPSC lines can vary. Established iPSC cultures can be split with a ratio of 1:3 to 1:6, meaning cell clumps from one well can be plated into three to six wells.
Avoid disturbing the plate overnight to ensure maximum attachment of cells and prevent uneven distribution of cell clumps, which may lead to increased iPSC differentiation.
Some cell debris the day after passaging is normal. iPSCs can be passaged or frozen down with mTeSRTM Plus freezing medium when they reach 80% confluence. Avoid over-confluence to prevent spontaneous differentiation.
For MSC differentiation, iPSCs should be at least 40% confluent two days after seeding. If there are insufficient cells for differentiation initiation, incubate them for one or two additional days, but avoid over-confluence before MSC differentiation to prevent random differentiation.
During the initial days of iPSC differentiation into MSCs, wash the differentiated cells with DPBS w/o Ca2+ and Mg2+ when renewing the medium due to a high number of dead cells. Subsequently, if the spent medium is clear and there are fewer dead cells, washing the cells becomes unnecessary.
Culture cells on gelatin-coated plates up to passage 2; as passaging continues, cells acquire a homogeneous, fibroblast-like morphology on gelatin-coated tissue culture plates. Beyond passage 2–3, use tissue culture plates without gelatin coating.
This methodology has been successfully employed in our lab to generate mesenchymal-like progenitors from various sources of human iPS cells. The multilineage differentiation potential of human iMSCs, including osteogenesis, adipogenesis, and chondrogenesis, has been established in vitro (Khan et al., 2023).
A cell density of 1 × 104 cell/cm2 is recommended for the growth of MSCs derived from human iPSCs. At this stage, cells can be directly split into new T25 flasks at a ratio of 1:3–1:4.
When iPSC-MSCs (iMSCs) reach 80% confluence in a 10 cm culture dish, they can be frozen into 3–5 cryogenic vials.
Remember to include groups for unstained control and negative control (isotype-matched antibodies, such as IgG1-PE and IgG2b-FITC) during experimentation.
Ensure that MSCs are in a single-cell suspension to prevent clogging of the flow tubing and to avoid damage to the flow cell instrument.
For optimal results, it is recommended to analyze samples on the same day. If immediate analysis is not possible, store the samples in the dark at 4 °C for several days until analysis.
Use enzyme solutions (trypsin-EDTA, ReLeSR) and PBS at ambient temperature.
To promote optimal micromass attachment, it might be necessary to coat the plates with 0.1% gelatin. Ensure thorough drying of the plates before micromass formation to prevent cell spreading.
Prevent micromasses from drying during the 2 h incubation period by adding PBS to the reservoir between wells.
To facilitate visualization of the pellets during paraffin sectioning, a brief addition of Eosin or Alcian blue stain at 1:100 to the final 95% ethanol wash can provide color before complete dehydration in 100% ethanol and xylene.
During handling of the cryogenic vials from liquid N2 tank, always wear proper personal protective equipment (PPE) such as safety goggles and/or face shield and cryogen gloves; do not leave skin exposed. Do not wear metal jewelry or watches.
Acknowledgments
This work was supported by funds from Veteran Affairs and Emory University School to Medicine. This research was funded by Georgia CTSA/REM Pilot Project 00080502 to H.D., Veteran Affairs CaReAP Award (I01-BX004878) to H.D. This protocol is derived from the original work Khan et al. (2023)
Competing interests
The authors declare no conflicts of interest.
References
Dicks, A. R., Steward, N., Guilak, F. and Wu, C. L. (2023). Chondrogenic Differentiation of Human-Induced Pluripotent Stem Cells. Methods Mol. Biol. 2598: 87–114.
Guzzo, R. M. and Drissi, H. (2015). Differentiation of Human Induced Pluripotent Stem Cells to Chondrocytes. Methods Mol. Biol. 1340: 79–95.
Guzzo, R. M., Gibson, J., Xu, R. H., Lee, F. Y. and Drissi, H. (2013). Efficient differentiation of human iPSC-derived mesenchymal stem cells to chondroprogenitor cells. J. Cell. Biochem. 114(2): 480–490.
Khan, N. M., Diaz-Hernandez, M. E., Chihab, S., Priyadarshani, P., Bhattaram, P., Mortensen, L. J., Guzzo, R. M. and Drissi, H. (2023). Differential chondrogenic differentiation between iPSC derived from healthy and OA cartilage is associated with changes in epigenetic regulation and metabolic transcriptomic signatures. eLife 12: e83138.
Lamandé, S. R., Ng, E. S., Cameron, T. L., Kung, L. H. W., Sampurno, L., Rowley, L., Lilianty, J., Patria, Y. N., Stenta, T., Hanssen, E., et al. (2023). Modeling human skeletal development using human pluripotent stem cells. Proc. Natl. Acad. Sci. U. S. A. 120(19): e2211510120.
Mahboudi, H., Soleimani, M., Enderami, S. E., Kehtari, M., Ardeshirylajimi, A., Eftekhary, M. and Kazemi, B. (2018). Enhanced chondrogenesis differentiation of human induced pluripotent stem cells by MicroRNA-140 and transforming growth factor beta 3 (TGFβ3). Biologicals 52: 30–36.
Suchorska, W. M., Augustyniak, E., Richter, M. and Trzeciak, T. (2017). Gene expression profile in human induced pluripotent stem cells: Chondrogenic differentiation in vitro, part A. Mol. Med. Rep.15(5): 2387–2401.
Wu, C. L., Dicks, A., Steward, N., Tang, R., Katz, D. B., Choi, Y. R. and Guilak, F. (2021). Single cell transcriptomic analysis of human pluripotent stem cell chondrogenesis. Nat. Commun. 12(1): 362.
Zujur, D., Al-Akashi, Z., Nakamura, A., Zhao, C., Takahashi, K., Aritomi, S., Theoputra, W., Kamiya, D., Nakayama, K., Ikeya, M., et al. (2023). Enhanced chondrogenic differentiation of iPS cell-derived mesenchymal stem/stromal cells via neural crest cell induction for hyaline cartilage repair. Front. Cell Dev. Biol. 11: e1140717.
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© 2023 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Peer-reviewed
A New Behavioral Paradigm for Visual Classical Conditioning in Drosophila
MB Mercedes Bengochea
TP Thomas Preat
BH Bassem Hassan
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4875 Views: 614
Reviewed by: Nafisa M. Jadavji Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Cell Reports Jul 2023
Abstract
Visual learning in animals is a remarkable cognitive ability that plays a crucial role in their survival and adaptation. Therefore, the ability to learn is highly conserved among animals. Despite lacking a centralized nervous system like vertebrates, invertebrates have demonstrated remarkable learning abilities. Here, we describe a simple behavioral assay that allows the analysis of visual associative learning in individually traceable freely walking adult fruit flies. The setup is based on the simple and widely used behavioral assay to study orientation behavior in flies. A single wing-clipped fly that has been starved for 21 h is placed on a platform where two unreachable opposite visual sets are displayed. This visual learning protocol was initially developed to study the cognitive ability of fruit flies to process numerical information. Through the application of the protocol, flies are able to associate a specific visual set with an appetitive reward. This association is revealed 2 h later during the testing session where we observed a change in their preference upon learning (i.e., change in their spontaneous preference). Moreover, this protocol could potentially be used to associate any other visual object/property to the reward, expanding the opportunities of studying visual learning in freely walking fruit flies at individual level.
Graphical overview
Graphical overview of conditional learning protocol. Forty-eight hours before conditioning, the wings of the flies are clipped, and individual flies are left to recover in a fresh food vial. Twenty-one hours before the conditional learning starts, flies are transferred to a starvation vial containing wet paper. The training session consists in placing a drop of sugar next to the place with the lower number of objects (numerosity) and a drop of water next to the larger numerosity. The fly is placed in the arena and left to freely walk for three minutes. Once the session is finished, the fly is placed back in their empty vial for 2 h until the testing session starts.
Keywords: Drosophila melanogaster Appetitive learning Visual conditioning Short-term memory Cognitive ability
Background
Associative learning enables animals to prepare for behaviorally relevant events, thus being highly conserved. Learning to predict events in the environment and which stimuli tend to occur together help us interact effectively with our environment. This cognitive process has been shown to play a role in a wide diversity of behaviors, including interactions with predators, prey, rivals, and mates. Thus, learning is expected to be essential for survival and reproduction in many species. The study of such associative learning in simple model systems, like the fruit fly Drosophila melanogaster, facilitates its understanding especially at multiple levels of analysis. This model allowed researchers to uncover principles and mechanisms of learning and memory as it offers a wide range of powerful genetic tools to dissect intrinsic molecular mechanisms (McGuire et al., 2005; Pitman et al., 2009).
During Pavlovian classical conditioning, animals learn to associate a salient unconditioned stimulus with a neutral stimulus. Olfactory classical conditioning is a well-studied form of associative learning in the fruit fly D. melanogaster (see Busto et al., 2010). Subsequently, other methods to measure various types of learning and memory have been proposed, including courtship conditioning (Koemans et al., 2017), aversive phototaxic suppression assay (Ali et al., 2011), and wasp-exposure conditioning (Kacsoh et al., 2019).
Regarding visual learning, it has been extensively studied using the so-called flight simulator (Wolf and Heisenberg, 1991). Here, a tethered fly is flying stationarily in the center of a cylindrical virtual arena panorama where different visual stimuli are displayed. Combined with heat punishment, the setup changes into an operant conditioning paradigm in which the fly learns to control the appearance of the punisher. Like this, animals have to learn from the consequences of their own and voluntary actions in order to avoid the punishment. Other studies show that populations of flies are able to associate the color of the illumination alone (Vogt et al., 2015) or in combination with odors (Vogt et al., 2014; Thiagarajan et al., 2022; Okray et al., 2023) with an appetitive or aversive stimulus.
Despite the relative abundance of aversive learning assays in which flies operantly avoid heat by choosing certain orientations relative to landmarks, only a few behavioral paradigms for appetitive visual learning in adult Drosophila have been reported to date (Heisenberg, 1989; Schnaitmann et al., 2010; Vogt et al., 2014). Here, we established a new behavioral paradigm for visual classical conditioning in freely-walking adult Drosophila. In their natural environment, fruit flies primarily move by walking rather than flying. Studying them in a walking state allows us to better understand their visual learning abilities in a context that aligns with their ecological niche. This assay allows analysis of the behavior of individual flies while they make a choice, learn, and memorize (short-term memory) visually rewarded associations. The paradigm described here is based on a simple behavioral assay to study orientation behavior in flies (Buridan paradigm), which can be easily assembled with a few available supplies (Figure 1). Particularly, this visual learning is based on the spontaneous numerical preference of flies, which manifests as a larger occupancy next to the higher numerosity in a two-choice test (Bengochea et al., 2023). The setup consists of presenting two visual sets of different numbers of objects that are located opposite to each other, in order to be processed individually and not treated as a whole. Since flies have a spontaneous preference for larger sets of objects, the appetitive reward is placed next to the non-preferred numerical set (lower numerosity), while the innately preferred one is water-paired (Figure 1B). A single fly is placed on the platform (Figure 1C). During the 3-min training trial, the fly explores the arena and eventually reaches the sugar and water. Two hours after training, the learned preference of the fly is evaluated during a 5-min testing phase (Graphical overview). By analyzing the walking occupancy in the arena, we are able to observe a decrease or change in their natural tendency to prefer the larger quantities due to the association of the sugar with the visual set. The representative results of this assay reveal that the application of the protocol impacts on the fly’s numerical preference (Figure 2, Bengochea et al., 2023). A single training trial of 3 min is sufficient to teach flies to associate an appetitive reward with a visual set that lasts at least two hours and to partially or totally reverse a spontaneous tendency.
Materials and reagents
Fly stocks
Wildtype CantonS flies (BL#64349, 7 days old at the moment of the training session)
Materials
Eight circular fluorescent tubes (Philips, L 40 w, 640C circular cool white)
Lamp holder and connectors G10Q (Vossloh, catalog number: 101528)
Two electrical transformers (Osram Quicktronic QT-M 1×26–42)
Eight power cables with switch (Tibelec, catalog number: 163910)
Diffuser paper (Canson, Translucent paper 180 g/m2)
Ten squares of black paper (31 mm × 31 mm)
Filter paper (Whatman, catalog number: 1001-110)
Paint brush (Size 1; Boesner, model: Da Vinci Nova Serie 1570, catalog number: D15701))
Empty vials (Polystyrene; Dutscher, catalog number: 789001B)
Invisible tape (Scotch Magic 3M, catalog number: 7100027389)
Seamstress ruler
Pencil
Reagents
Distilled water
Mineral water (Evian)
Glucose 1.5 M (Sigma-Aldrich, catalog number: G8270), prepared with Evian water
70% ethanol (VWR, catalog number: 83801)
Agar (Genesee Scientific, catalog number: 66-103)
Cornmeal (Genesee Scientific, catalog number: 62-100)
Yeast (Genesee Scientific, catalog number: 62-106)
Glucose (Sigma-Aldrich, catalog number: 68270)
Molasses (Genesee Scientific, catalog number: 62-117)
Ethanol (VWR, catalog number: 83801)
Nipagin (Thermo Fisher Scientific, catalog number: A14289)
Propionic acid (Sigma-Aldrich, catalog number: P5561)
Drosophila standard cornmeal/agar food (see Recipes)
Recipes
Drosophila standard cornmeal/agar food
8 g of agar
60 g of cornmeal
50 g of yeast
20 g of glucose
50 g of molasses
19 mL of ethanol 70%
1.9 g of Nipagin
10 mL of propionic acid
Bring up to 1 L of final volume with distilled water
Equipment
Scissor (Vannas Spring Scissors, 3 mm Blades; Fine Science Tool, catalog number: 15000-00)
CO2 pads (Dutscher, catalog number: 789183)
CO2 pistol (Dutscher, catalog number: 789096)
Two Buridan’s Paradigm: include baseplate with platform and chamber, acrylic cylinder, and posts to hold the lamps (Peira Scientific Instruments, https://www.peira.be/)
Two webcams (Logitech HD Pro c920 webcam full HD)
Two PCs
Fly incubator (Panasonic, MIR-554-PE; fluorescent lamp: Panasonic FL15D 15W)
Microscope (Leica, M80)
Software
Buritrack (http://buridansourceforge.net)
R Core Team (https://www.R-project.org/)
RStudio (https://support--rstudio-com.netlify.app/products/rstudio/)
Procedure
Prepare flies for behavioral recording
Raise flies at 12:12 h light/dark cycle, 60% humidity and 25 °C.
Hatched flies are disposed of in case their age is not controlled (including those that transiently stick to the food or walls).
Collect all newly hatched flies into a new experimental vial (no more than 20 flies per vial). Flies should be 0–1 day old.
Collect 5-day-old flies (at least 60 flies: 30 for control group and 30 for trained group) by tapping and transferring them to a new vial with fresh food.
Anesthetize the flies by holding the vial upside down and inserting the CO2 pistol. Fill in the CO2 for a few seconds until flies are asleep (flies do not move anymore and fall into the vial cap).
Transfer the flies to a CO2-dispensing porous pad.
Clip both wings using scissors under the microscope. The wings need to be shortened to at least 1/3 of the original length and must be straight to the crossing line of both wings. The flies must not be anesthetized for more than 5 min; therefore, it is useful to portion the flies.
Save each fly in individual vials with food (1 cm full) at 25 °C. Split and store them in two different racks—one for the control group (CT) and one for the trained group (TR).
Seven-day-old flies are used for the experiment. Twenty-one hours before running the experiment, place each fly in a starving vial containing a piece of filter paper watered with Evian water. Store them at 25 °C until running the experiment.
Prepare behavioral arena for learning experiment
Two setups will be prepared in parallel to run the control (CT) and trained (TR) group at the same time. It is important to switch the setups, so that the two groups are run in both assays. For half of the animals in the experiment, the acrylic cylinder must be rotated 180° to exclude any uncontrolled and systematic influence of other stimuli of the surroundings.
Place the visual diffuser paper on the inside of the acrylic cylinder.
Tape the black squares with scotch tape: on one side of the cylinder, place the three squares at 40 mm from the bottom with a distance between them of 22 mm. Tape the remaining two squares in the opposite side of the cylinder (180° apart) using the same distances. It is important that the sets are well aligned. The center of the sets must be 180° apart (Figure 1).
Open the Buritrack software and place the visual sets in the top and bottom part of the image captured by the webcam (Figure 1B). The contrast and luminosity of your camera may need to be adjusted, such that the arena is visible.
Add distilled water to the Buridan chamber.
Darken the room: close blinds and curtains and switch off the room lights.
Switch on the Buridan fluorescent lights.
Clean the platform with ethanol 70%.
Place the round filter paper on the platform.
In the Buridan planned to run the trained group, add two drops (~100 μL) of glucose solution next to the visual stimulus to be conditioned (e.g., 2 squares set) and two drops (~100 μL) of Evian water to the other visual stimulus (e.g., 3 squares set). For the control group, in the other Buridan, place two drops of water next to each stimulus (Figure 1B).
Figure 1. Behavioral apparatus and walking recording. A. Outside view of the Buridan paradigm. B. Top view of the assay, where a filter paper (gray dash line) is placed in the platform to add water or sugar next to the visual stimulus (red dashed lines). C. Picture of the fly performing during the testing session. D, E. Image of the Buritrack software to set the parameters of the experiment.
Run the training session
Set the temperature of the platform to 25 °C.
Press the Button New Tracking and adjust all the parameters in the Tracking dialog (Figure 1D). Generate a folder named “Training” to save your recording. Open Buritrack and set the parameters of the recording. Browse the Training folder, select the name of the file (e.g., TR_f01 for the case of TR group female #1; CT_m01 for the case of CT group male #1) and the fly group (e.g., TR_f, for female TR flies; CT_m for CT male flies), and the recording time (in seconds).
Set the size of the arena. While setting the platform position, left-click on three points on the platform edge. Check that the drawn circle fits with the platform; if not, you can right-click to erase one point (one per click) (yellow line in Figure 1D).
Gently place the fly on the platform. You may want to adjust the luminosity of your camera again.
Activate tracking in Buritrack to record the walking behavior (Figure 1D).
If the fly jumps off of the platform, the tracker stops and plays a sound alarming the experimenter. Put the fly back on the platform using a brush and click Activate Tracking again to continue with the experiment.
After 3 min the session is over. Gently remove the fly from the platform and place it back in their starving vial.
Clean the platform with 70% ethanol, prepare a new filter paper with sucrose and water for the next fly, and set the new file name in Buritrack. Once it is done, place the following fly and start a new recording by clicking the New Tracking button.
Continue running each fly individually by cleaning the platform with 70% ethanol between flies and using a new clean filter paper each time. Half of the animals will be trained and tested with one stimulus orientation (e.g., three squares on the top part of the image in the Buritrack software), and the other half will be trained and tested with the opposite orientation of the visual sets (e.g., three squares on the bottom part of the image).
Run the testing session
Right after finishing the training session, clean the platform with abundant water and 70% ethanol to remove any trace of glucose.
Two hours after finishing the training session, place a clean filter paper on the platform.
Place the fly to record the testing session.
Generate a folder named “Testing” to save your recording. Open Buritrak and set the parameters of the recording. Browse the Testing folder and select the name of the file, fly group, and the recording time (300 s). It is important to use the same names as in the training session (e.g., TR_f01, CT_m01) to then analyze the behavior of the very same fly during the training and testing sessions.
Set the size of the arena.
Activate tracking in Buritrack to record walking behavior.
After running the recording, remove the filter paper, clean the platform with 70% ethanol, and place a new round filter paper for testing the next fly.
Data analysis
Once the experiment is finished (at least 30 animals per group), store the data of the training and testing sessions in different folders by pooling the data collected in each setup.
Open RStudio.
Invert the x-y position of half of the animals (that were run with inverted configuration of the visual stimuli) by using the change_orientation_xy.R file (https://github.com/HassanLab/Bengochea_etal_2023/blob/main/change_orientation_xy.R).
Analyze the training and testing sessions by running the Analysis.R code (https://github.com/HassanLab/Bengochea_etal_2023/blob/main/Analysis_Learning.R).
Check if all flies (from trained and control groups) reached the sugar and water. Flies that did not visit either the sucrose or the water must be discarded.
After running the Analysis.R code, you will find the following output files:
Individual trajectories: In a single PDF you will find the trajectory of each fly. The color and the size of the path changes depending on the amount of time spent in each position in the platform.
Individual transition plots: Transition plots were done as described before (Colomb et al., 2012). Briefly, the platform was divided in 60 × 60 hexagons and the fly’s position raised the count of each hexagon by one in the arena. The scale starts at 0 (blue) and goes up until a value calculated by the 95% quantile of the count-distribution (red).
YPosition density plot: Population Y position density plot by group (Figure 2).
Transition plots by group/group and sex: Population transition plots depending on the group of interest.
PI Comparison: Final boxplot to compare the performance of each group. Each dot indicates the PI for each fly tested. To calculate the preference index, the arena was divided into three zones. We sum the density of passage of the hexagons within zones close to the visual stimuli while the center part of the arena was not analyzed. Values indicate mean ± SD. The preference index was calculated as:
For the statistical analyses, you should check for normal data distribution using the Shapiro-Wilk normality test. Then, we chose the appropriate parametric or non-parametric test. We statistically compared groups with the non-parametric Wilcoxon rank sum test (Figure 2).
Figure 2. Flies trained in a 2 vs. 3 squares contrast showed a preference for the two squares during the testing session. Kernel density plots on the left denote the position permanence of the fly along y-axis for each group and session. On the right, boxplots show median population preference index. Trained flies significantly preferred the smaller set of squares [n = 45, PI = -0.20 ± 0.34, t(44) = -3.93, p = 2.9e-04, One sample t-test], opposite to the control group that shows a preference for the three squares set [n = 42, PI = 0.39 ± 0.36, t(41) = 7.19, p = 8.87e-09, One sample t-test; comparison between groups: t(84) = 7.9, p = 7.94e-12, Welch two sample t-test]. Box limits: upper (75) and lower (25) quartiles, and whiskers, 1.5× inter quartile range. Each dot corresponds to a single fly performance. Gray dots indicate outliers.
Video 1. Trajectories of two different flies: one that belongs to the control group (red traces) and one to the trained group (blue traces) during the training and testing sessions.This visual representation shows that trained flies tend to spend significantly more time in the proximity of the set of two squares when compared to the control group of untrained flies.
Limitations
Since flies are placed in the platform to freely explore the arena, one important limitation of the protocol is that we cannot control the amount of time each fly spends in contact with the sugar or water. Furthermore, we cannot control which stimulus will first reach the fly.
Discussion and conclusions
Studying the cognitive abilities and learning processes of animals is crucial for understanding their behavior and adaptation in their respective ecological niches. Visual learning, in particular, plays a vital role in an animal's survival and decision-making abilities.
Here, we established a new behavioral paradigm for visual classical conditioning in freely walking individual adult Drosophila. Our protocol, with a single training trial of 3 min, generates a visual memory that lasts at least 2 h, allowing the study of short-term visual memories. However, it remains to be analyzed whether flies can form long-term visual memories by applying longer training protocols. This protocol for studying visual learning in fruit flies has a significant impact on the field of research. Due to the flexibility of visual objects that can be presented, the protocol can be adapted to study other properties of visual learning and will allow to delve deeper into the intricate mechanisms underlying visual learning in insects.
This protocol adapts the well-standardized methodology for studying orientation behavior in freely walking flies, enabling other researchers to easily replicate the experiments. This contributes to the cumulative knowledge in the field and facilitates the validation of findings across different laboratories.
Studying invertebrate learning not only provides insights into the cognitive capacities of these animals but also contributes to our understanding of the evolution and mechanisms of learning across different species. By unraveling the intricacies of invertebrate learning, we can gain valuable knowledge about the fundamental principles of learning and cognition in general.
General notes and troubleshooting
Under optimal conditions, the results of experiments conducted on different days, seasons, and by different experimenters are reproducible. However, fluctuations in room temperature and humidity can impact behavioral performance. Therefore, it is crucial to maintain stable room conditions. When clipping the wings, it is important to handle the flies carefully to avoid causing harm. It is advisable to clip more flies than necessary for the experiment in case of any losses due to clipping or starvation. The remaining wing length should not exceed 1/3 of the original length to prevent excessive jumping from the platform. Strict timing is essential for both the training and testing sessions, and it can be challenging to run parallel groups. Efficiently placing and removing the flies from the platform is crucial to ensure consistent session durations for all flies. Prior training and practice are recommended before starting the experiment.
Acknowledgments
This protocol was originally published in a research manuscript by Bengochea et al. (2023). This work was supported by the Investissements d’Avenir program (ANR-10-IAIHU-06), Paris Brain Institute-ICM core funding, Allen Distinguished Investigator award (12202), the Roger De Spoelberch Prize, an NIH Brain Initiative RO1 grant (1R01NS121874-01) and ANR grant QuantSocInd (ANR-19-CE16-000) (to B.A.H.) and The Big Brain Theory Program from the Paris Brain Institute (BBT.3400.COUNTINGFLIES, to B.H.). We thank the ICM RnD unit and its staff for help with constructing the Buridan arenas.
Competing interests
The authors have no conflict of interest to declare.
References
Ali, Y. O., Escala, W., Ruan, K. and Zhai, R. G. (2011). Assaying Locomotor, Learning, and Memory Deficits in Drosophila Models of Neurodegeneration. J. Visualized Exp.: e3791/2504. doi: 10.3791/2504
Bengochea, M., Sitt, J. D., Izard, V., Preat, T., Cohen, L. and Hassan, B. A. (2023). Numerical discrimination in Drosophila melanogaster. Cell Rep. 42(7): 112772. doi: 10.1016/j.celrep.2023.112772
Busto, G. U., Cervantes-Sandoval, I. and Davis, R. L. (2010). Olfactory Learning in Drosophila. Physiology 25(6): 338–346. doi: 10.1152/physiol.00026.2010
Colomb, J., Reiter, L., Blaszkiewicz, J., Wessnitzer, J. and Brembs, B. (2012). Open Source Tracking and Analysis of Adult Drosophila Locomotion in Buridan's Paradigm with and without Visual Targets. PLoS One 7(8): e42247. doi: 10.1371/journal.pone.0042247
Heisenberg, M. (1989). Genetic approach to learning and memory (mnemogenetics) in Drosophila melanogaster. In Fundamentals of memory formation: Neural plasticity and brain function. Gustav Fisch. Stuttg. Ger. 3–45.
Kacsoh, B. Z., Bozler, J., Hodge, S. and Bosco, G. (2019). Neural circuitry of social learning in Drosophila requires multiple inputs to facilitate inter-species communication. Commun. Biol. 2(1): e1038/s42003–019–0557–5. doi: 10.1038/s42003-019-0557-5
Koemans, T. S., Oppitz, C., Donders, R. A. T., van Bokhoven, H., Schenck, A., Keleman, K. and Kramer, J. M. (2017). Drosophila Courtship Conditioning As a Measure of Learning and Memory. J. Visualized Exp.: e3791/55808. doi: 10.3791/55808
McGuire, S. E., Deshazer, M. and Davis, R. L. (2005). Thirty years of olfactory learning and memory research in Drosophila melanogaster. Prog. Neurobiol. 76(5): 328–347. doi: 10.1016/j.pneurobio.2005.09.003
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Schnaitmann, C. (2010). Appetitive and aversive visual learning in freely moving Drosophila. Front. Behav. Neurosci. 4: e00010. doi: 10.3389/fnbeh.2010.00010
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Article Information
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Peer-reviewed
Measuring Action Potential Propagation Velocity in Murine Cortical Axons
OK Oron Kotler
YK Yana Khrapunsky
IF Ilya Fleidervish
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4876 Views: 355
Reviewed by: Olga KopachMarco CanepariOlga tyurikova
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Cited by
Original Research Article:
The authors used this protocol in eLIFE Feb 2023
Abstract
Measuring the action potential (AP) propagation velocity in axons is critical for understanding neuronal computation. This protocol describes the measurement of propagation velocity using a combination of somatic whole cell and axonal loose patch recordings in brain slice preparations. The axons of neurons filled with fluorescent dye via somatic whole-cell pipette can be targeted under direct optical control using the fluorophore-filled pipette. The propagation delays between the soma and 5–7 axonal locations can be obtained by analyzing the ensemble averages of 500–600 sweeps of somatic APs aligned at times of maximal rate-of-rise (dV/dtmax) and axonal action currents from these locations. By plotting the propagation delays against the distance, the location of the AP initiation zone becomes evident as the site exhibiting the greatest delay relative to the soma. Performing linear fitting of the delays obtained from sites both proximal and distal from the trigger zone allows the determination of the velocities of AP backward and forward propagation, respectively.
Key features
• Ultra-thin axons in cortical slices are targeted under direct optical control using the SBFI-filled pipette.
• Dual somatic whole cell and axonal loose patch recordings from 5–7 axonal locations.
• Ensemble averaging of 500–600 sweeps of somatic APs and axonal action currents.
• Plotting the propagation delays against the distance enables the determination of the trigger zone's position and velocities of AP backward and forward propagation.
Keywords: Action potential Action current Pyramidal neuron Neocortex Propagation velocity Trigger zone Backpropagation Loose patch Whole-cell recording
Background
Neurons generate action potentials (APs), all-or-none electrical events propagating along their processes by eliciting transient changes in membrane permeability (Hodgkin and Huxley, 1952). One of the most critical AP parameters, propagation velocity, was first measured in peripheral axons in the second half of the 19th century (Helmholtz, 1850; Bernstein, 1868). Measuring the propagation velocity in ultra-thin central axons became possible only recently, as it requires previously unachievable temporal and spatial resolution. Several studies have attempted to measure propagation delays using simultaneous whole-cell recordings from soma and giant axonal blebs (Shu et al., 2007; Kole et al., 2007) or dual whole-cell, cell-attached recordings from the soma and axonal trunk (Sasaki et al., 2012). Optical recordings from neurons stained with voltage-sensitive dyes were used to determine the AP initiation site and describe the AP propagation pattern and velocity (Palmer and Stuart, 2006; Popovic et al., 2011). We describe how to measure AP propagation velocity in fluorescently labeled cortical axons in acute brain slice preparation by recording whole-cell somatic voltage and action currents from multiple axonal locations using a loose patch configuration. We previously employed this method in cortical brain slices (Baranauskas et al., 2013; Kotler et al., 2023). We believe that, with minor adjustments, this method can be applied to determine the AP propagation speed in various brain regions and experimental setups.
Accurate measurement of both distance and time delays is crucial to calculate propagation velocity. Measuring the distance poses fewer technical challenges once the location of the AP trigger zone is known. Thus, not establishing the trigger zone position before attempting to determine the propagation velocity could lead to inaccurate conclusions, given that the propagation delay between the soma and the axonal segment located 100 µm away is almost zero. Accurately measuring the time delays presents a greater challenge since it demands sufficient temporal and amplitude resolution in action current recording. For adequate temporal resolution, we suggest widening the bandwidth of the axonal voltage clamp recording to 20 kHz. Spike-triggered averaging of 500–600 sweeps should be sufficient to resolve action currents with a signal-to-noise ratio above 3.
Materials and reagents
Biological materials
Experimental animals: ICR mice, P6 or older (Envigo, Israel), are suitable for brain slice experiments. Any other mouse strain could be used. All animal experiments should be performed in accordance with institutional and national guidelines for the care and use of laboratory animals.
Materials
Whatman qualitative filter paper, Grade 1, circles, diam. 90 mm (Sigma-Aldrich, catalog number: WHA1001090)
Petri dishes, polystyrene, size 100 mm × 15 mm (Sigma-Aldrich, catalog number: P5856)
Feather double-edge carbon steel blades (Ted Pella, catalog number: 121-9)
GEM single edge, carbon steel blades, uncoated (Ted Pella, catalog number: 121-1)
PELCO Pro CA44 instant tissue adhesive (Ted Pella, catalog number: 10033)
Reagents
Sodium-binding benzofuran isophthalate (SBFI, Tetraammonium Salt, cell impermeant) (Invitrogen, catalog number: S1262)
Sodium chloride (NaCl) (Biolab, CAS number: 7647-14-5)
Potassium chloride (KCl) (Sigma-Aldrich, CAS number: 7447-40-7)
Calcium chloride dihydrate (CaCl2) (Sigma-Aldrich, CAS number: 10035-04-8)
Magnesium sulfate heptahydrate (MgSO4) (Sigma-Aldrich, CAS number: 10034-99-8)
Sodium phosphate monobasic monohydrate (NaH2PO4) (Acros Organics, CAS number: 10049-21-51)
Sodium bicarbonate (NaHCO3) (Biolab, CAS number: 144-55-8)
D-glucose (Sigma-Aldrich, CAS number: 50-99-7)
Potassium D-gluconate (Sigma-Aldrich, CAS number: 299-27-4)
Magnesium chloride anhydrous (MgCl2) (Sigma-Aldrich, CAS number: 7786-30-3)
HEPES (Sigma-Aldrich, CAS number: 7365-45-9)
Distilled water (H2O)
Isoflurane (Piramal Critical Care, Bethlehem, PA, USA)
Solutions
Artificial cerebrospinal fluid ×10 stock solution (aCSFx10) (see Recipes)
Artificial cerebrospinal fluid solution (aCSF)-working solution (see Recipes)
MgSO4 1 M stock (see Recipes)
CaCl2 1 M stock (see Recipes)
K+-based intracellular solution (see Recipes)
Recipes
Artificial cerebrospinal fluid ×10 stock solution (aCSFx10)
Reagent Final concentration (mM) m.w. Amount (g) for 1 L
NaCl 1,240 58.44 72.466
KCl 30 74.56 2.237
NaHCO3 260 84.01 21.843
NaH2PO4 12.5 137.99 1.725
Artificial cerebrospinal fluid solution (aCSF)-working solution
Reagent Final concentration Amount for 1 L
aCSFx10 n/a 100 mL
D-glucose 1 M 10 mM 10 mL
MgSO4 1 M 2 mM 2 mL
H2O n/a ~850 mL
Bubble with 95% O2-5% CO2 gas mixture for 15 min
CaCl2 1 M 2 mM 2 mL
Add H2O up to 1 L exactly
Remark: Bubbling with 95% O2-5% CO2 gas mixture is necessary to avoid precipitation of the divalent ions.
MgSO4 1 M stock
Reagent Final concentration Amount (g) for 50 mL
MgSO4 1 M 10.165
Add H2O up to 50 mL exactly
CaCl2 1 M stock
Reagent Final concentration Amount (g) for 50 mL
CaCl2 1 M 7.352
Add H2O up to 50 mL exactly
K+-based intracellular solution
Reagent Final concentration Amount for 100 mL
Potassium D-gluconate (m.w. 234.25) 130 3.045 g
KCl 1 M 6 0.6 mL
MgCl2 1 M 2 0.2 mL
HEPES 1 M 10 1 mL
NaCl 1 M 4 0.4 mL
H2O n/a to 100 mL
Titrate to pH 7.25 with KOH 1 M
Due to the large size and spatial complexity of L5 pyramidal neurons in slices, ATP and GTP supplements were not added to the intracellular solution. The freshly prepared K+-based intracellular solution was tested in whole-cell recording from several neurons before supplementing with SBFI. The SBFI-containing solution was separated in aliquots of 100 µL into small Eppendorf microtubes and stored at -20 °C.
Equipment
Upright microscope equipped with IR-DIC optics (Olympus, model: BX51WI)
Water-immersion objective lens (magnification: 60×, numerical aperture: 1.0, working distance: 2 mm) (Olympus, LUMPLFLN60XW, product number: N2667800)
Shifting table (Luigs & Neumann, model: V380FM), control box (Luigs & Neumann, model: SM7)
Two LN mini manipulators (Luigs & Neumann, model: Mini 25)
Slice mini chamber with temperature controller (Luigs & Neumann, 200-100 500 0150-S and 200-100 500 0145)
U-shaped platinum grid to weigh down a slice in a recording chamber
C2400 CCD Camera (Hamamatsu)
NeuroCCD-SMQ camera, 0.38× coupler (to achieve the final magnification of 1 pixel = 1 µm with 60× objective), and NeuroPlex software (Redshirt Imaging)
High Power Collimated LED Light Source, 385 nm (Prizmatix, model: Mic-LED-385)
Modified filter set, dichroic mirror 400 nm, long pass emission filter 420 nm (Olympus, model: U-MNU2)
MultiClamp-700B amplifier, equipped with two CV-7B headstages (Molecular Devices)
Semiautomatic microtome with a vibrating blade (Leica Biosystems, model: VT-1200) with Vibrocheck unit for adjustment of vertical deflection of the blade
Analytical balance (MRC-Laboratory Equipment, model: ASB-220-C2)
Bench pH meter (Hanna Instruments, model: HI2211)
Small scissors, tweezers, spatula, Pasteur pipette
Micropipette puller (Sutter Instrument, model: P-97)
Ultima IV 2P microscope (Bruker) equipped with Mai Tai DeepSee laser (Spectra Physics)
Semi-Automatic Vibrating Blade Microtome (Leica Biosystems, model: VT-1200)
Software and datasets
pCLAMP 9 Electrophysiology Data Acquisition & Analysis Software (Molecular Devices, Version 9.2.1.9, 2007) for data acquisition; pCLAMP 11 Electrophysiology Data Acquisition & Analysis Software (Molecular Devices, Version 11.2.2.17, 2022) for data analysis
IDL (Exelis Visual Information Solutions, Version 8.3, 2013) and NeuroPlex (RedShirt Imaging, Version 10.2.2, 2011)
ImageJ (Version 1.54f 29, 2023)
Excel (Microsoft Office 365, 2022)
Procedure
Preparation of brain slices
Coronal or sagittal slices are suitable, as these plains of section preserve the axonal and apical dendritic tree of L5 pyramidal neurons.
Preparing the solution for brain exposure and slicing:
Transfer 250 mL of freshly prepared aCSFx1 solution into a 300 mL chemical glass beaker.
Place the glass beaker into the ice box.
Bubble the aCSFx1 solution with 95% O2-5% CO2 gas mixture for at least 15 min to attain equilibrium in gas partial pressures.
Cover the glass beaker with Parafilm.
Cool the solution to 4 °C.
Add 0.7 mL of MgSO4 1 M.
Add 0.25 mL of CaCl2 1 M (prior bubbling with 95% O2-5% CO2 gas mixture is necessary to prevent Ca2+ ions precipitation).
Cover the chemical glass with Parafilm.
Preparing the incubation solution for brain slices:
Transfer 500 mL of freshly prepared aCSFx1 solution into the slice incubator.
Bubble with 95% O2-5% CO2 for at least 15 min.
Cover the slice incubator with Parafilm.
Warm the aCSFx1 solution to 30 °C.
Add 0.7 mL of CaCl2 1 M.
Preparing the vibratome:
Put the black buffer tray and specimen holder into the freezer (-20 °C).
Take the double-edge feather blade, wash it with 70% ethanol, rinse with water, and wipe.
Insert the blade into the blade holder.
Insert the Vibrocheck unit into its slot and connect to the vibratome by the USB cable.
Wait for the appearance of a green light on the Down button.
Insert the Allen screwdriver into the blade holder slot and rotate it 90° counterclockwise until a white strip appears on the left side.
Push the Down button.
Push the Run button.
If the display shows a value different from 0 (e.g., -0.1), push the Stop button.
Release the blade.
Insert the screwdriver in the adjustment hole opening and turn in the "-" direction.
Push the Run button.
Repeat if the display shows a value different from 0.
After reaching 0, push the "Down" button to remember the parameters.
Move the blade to the top position and remove the Vibrocheck block.
Slicing procedure (done in the chemical hood):
Put the Whatman filter paper 90 mm into the 100 mm Petri dish and place the dish on ice.
Prepare the single edge, carbon steel blade.
Prepare surgical instruments: scissors, small mini-scissors, tweezers, spatula, and Pasteur cut pipette.
Bring the mouse from the animal facility.
Check the temperature of the solutions.
Take the black buffer tray and specimen holder from the freezer and place the black buffer tray on ice.
Wipe the specimen holder.
Note: The holder should be completely dry.
Place a small amount of instant tissue adhesive on the center of the specimen holder.
Fill the Petri dish with cold slicing solution (step A1).
Put two weighing dishes near the surgical instruments.
Fill two weighing dishes with cold slicing solution.
The next steps have to be performed in accordance with the Institutional and National Animal Care regulations.
Measure 0.5 mL of isoflurane and distribute it over the towel attached to the cover of the exicator.
Cover the exicator.
Put the animal into the exicator and quickly cover it.
Wait a minute for the isoflurane to take effect.
Take a mouse and cut the skin from its head to expose the skull.
Cut the head close to occipital bone with scissors.
Wash the head in the first weighing dish with cold slicing solution for a few seconds.
Cut the bone with surgical scissors as follows:
i. Insert the surgical scissors through the foramen magnum in the occipital bone.
ii. Continue along the occipital bone until the end of the lacrimal bone.
iii. Cut down from the lacrimal's end to the eye cavity.
Gently remove the cut bone with tweezers and expose the brain.
Put the brain into the second weighing dish with ice-cold slicing solution.
Transfer the brain into a Petri dish.
Cut the block of the brain (coronal or sagittal plane).
Paste it in the specimen holder.
Transfer the holder to a buffer tray.
Insert the buffer tray into the insert in the vibratome.
Fill the buffer tray with an ice-cold slicing solution.
Cut the first thick slice.
Discard the slice.
Adjust to 300 µm slice thickness and cut the slice.
Remove the slice with a paintbrush.
Collect the slice with a cut Pasteur pipette.
Transfer the slice to a warm incubation solution (step A2).
Repeat until the desired number of slices is cut.
Incubate the slices for 60 min.
Caution: It is advisable to perform steps 4r–4ee with maximum speed, ideally completing them in under 1 min. More extended periods of ischemia/hypoxia could cause irreversible damage to the brain tissue.
Whole-cell recording and filling neurons with fluorescent dye (always document the experiment in a lab book)
Turn on all equipment and perfuse aCSFx1 through the slice mini chamber.
Set the temperature under the objective to 32 °C using the temperature controller.
Transfer a 300 µm thick slice from the incubating chamber to the slice mini chamber and put a platinum grid to pin down the slice.
Focus on Layer 5 cell body layer (approximately 500–700 µm below the pial surface).
Identify an L5 Pyramidal neuron to patch using the 60× objective. Do not move the X and Y position of the microscope from this point on.
Pull two pipettes by using the P-97 micropipette puller. Note that the pipette resistance should be 10–12 MΩ when filled with K+-based intracellular solution or 7–9 MΩ when filled with aCSF. Note that optimal pipette size may vary depending on the specific preparations.
Cut a 4–5 mm wide, 10 mm long strip of Parafilm. Wrap the pipette with one or two layers of Parafilm, ~1 mm from the tip, to reduce the pipette stray capacitance.
Fill the first pipette with K+-based intracellular solution supplemented with SBFI (2 mM).
Fill the other pipette with aCSFx1 supplemented with SBFI (2 mM).
Place both pipettes in the micromanipulator pipette holders.
Apply gentle positive pressure, place the pipettes in the bath, focus on their tips, and move them down, one by one, toward the surface of the slice.
Stop moving the pipettes before approaching the cell; ensure that the cell body is below the whole-cell pipette.
Set the Multiclamp-700B Commander Channels 1 and 2 to voltage-clamp mode and correct the pipette offset.
Apply a seal test, a -10 mV voltage step, to both pipettes.
Approach the cell body with the first pipette by moving it along the z-axis until a dimple appears on the membrane.
Release the positive pressure and apply gentle suction while monitoring the current response to the voltage step until the gigaohmic seal forms.
Move the holding voltage to -70 mV and correct fast capacitance using automatic correction of the Commander.
Apply gentle and brief suction pulse with a syringe to break into whole-cell configuration (or by mouth). The successful break-in is evident by an abrupt increase in the amplitude of capacitative transients in response to -10 mV voltage steps.
Switch the Commander to the Current Clamp mode.
Correct the Bridge.
A Current Clamp whole-cell recording is established for pipette 1.
Measuring action currents along the axon
After establishing the Current Clamp whole-cell recording with pipette 1, wait 15 min to allow SBFI diffusion to the cell.
Focus the NeuroCCD-SMQ camera on the axon. L5 cell axons are the only fine processes emerging from the soma opposite the apical dendrite and exhibiting distinctive Na+ transients (Fleidervish et al., 2010; Baranauskas et al., 2013). Some neurons were examined live using a two-photon microscope after the physiological experiment. In these cells, axons were easily discerned from basal dendrites due to their lack of spines, confirming earlier observations during recording.
Acquire an 8 ms long single frame image with a NeuroCCD-SMQ camera using the NeuroPlex software. Use 385 nm high-intensity LED device as a light source and collect emission using the U-MNU2 filter set. The 8 ms long exposure time is sufficient to obtain a high-quality image while minimizing light intensity that might induce photo-dynamic damage to the cell.
Find a point of interest along the axon. The first point of interest could be at ~10 µm from the edge of the soma if the backpropagation velocity has to be measured; in experiments designed to measure the forward propagation, the first point of interest might be more distal than the presumed trigger zone, ~30–40 µm from the edge of the soma.
Bring pipette 2 to the point of interest (Figure 1b).
Touch the axon with the pipette 2. Ensure the touch by taking one or more single frames with the NeuroCCD-SMQ camera. The proper touch could be established by fine-tuning the pipette 2 position, with no suction applied to the pipette.
Deliver 2–7 ms long current steps via pipette 1. Increase the current step amplitude until it will be sufficient to elicit an AP.
Deliver 600 steps of 1.5× threshold amplitude at 2–5 Hz frequency. Record the voltage with pipette 1 and current with pipette 2.
Store the recording file for further analysis.
Find another point of interest (Figure 1c). It should be 10–20 µm further distally from the previous point of interest to ensure a significant time delay difference.
Repeat steps 5–9 for 6–10 points of interest within 10–150 µm of the axonal length.
Figure 1. Measuring the distances and propagation delays between the cell body (soma) and distinct points along the axon. a. Maximal intensity Z-stack projection of 39 two-photon optical sections through part of a Layer 5 pyramidal neuron filled with SBFI (red) obtained at a wavelength of 760 nm. The arrows indicate sites along the axon where loose patch recordings were performed. The sites are color coded as follows: 1 (green): 35 μm, 4 (yellow): 96 µm from the edge of the soma. b. Measurement of the propagation delay for site 1. Left, the same neuron as seen during the imaging experiment with the NeuroCCD–SMQ camera. The green arrows indicate the region corresponding to the edge of the soma and the site from where action currents were obtained. Right, top, 600 somatic dV/dt sweeps and corresponding axon currents from site 1, unaligned and following alignment using the time of dV/dtmax as the reference point. Right, bottom, ensemble average of the somatic dV/dt (bottom trace) and axonal current (top trace). Note that the peak of the axonal action current precedes the peak of the somatic dV/dt by 130 µs. c. Measurement of the propagation delay for site 4. For details, see b. Note that, at the distance of 96 µm, the somatic dV/dt peak precedes the peak of the action current by 20 µs.
Neuronal morphology (optional)
Turn on the Ultima IV 2P microscope and perfuse aCSF through the slice mini chamber.
Set the temperature under the objective to 32 °C using the temperature controller.
Transfer the slice containing SBFI-labeled cell from the slice mini chamber of the Olympus microscope setup to the slice mini chamber of the Ultima 2P microscope and put a platinum grid to pin down the slice.
Using a low-resolution Live Scan at 740 nm excitation wavelength, identify the SBFI-labeled neuron and place it in the center of the field of a 40× or 60× Olympus water-immersion lens.
Create a Z series of 20–40 images at 0.5–1.0 μm depth intervals (Figure 1a).
Open the resulting Z-stack in ImageJ to obtain morphometric data.
Data analysis
Measuring the axonal length between the soma and pipette 2:
Open the fluorescent image containing the cell body and the proximal axon, taken using the NeuroCCD-SMQ camera, in ImageJ (Figure 1b, 1c).
Measure the total SBFI fluorescence along the soma-axon axis using the Analyze-Plot Profile command. The half-difference of the somatic and axonal fluorescence should be assigned as zero point for the axonal length measurements (Baranauskas et al., 2013).
Draw the straight line or lines connecting the zero point and the location of pipette 2 over the axon.
Use the Pythagorean theorem to find the distance between the two points.
Measuring the time latency between pipettes 1 and 2:
Open the data file containing 600 consecutive sweeps of somatic voltage and currents collected from an axonal location in Clampfit 10 (Figure 1b, c).
Go to the Analysis Window Toolbar and push the Arithmetic button. In the Arithmetic window dialogue, select the Active Window, Somatic Vm signal, and All visible traces. Differentiate the somatic voltage sweeps using the expression T{VISIBLE} = diff(T{VISIBLE}), to obtain the dV/dt sweeps.
Align all sweeps and signals using the dV/dt positive peaks as a reference point using the Analyze-Time Shift-Align peaks command (replace wrapped samples with zeros). The alignment of sweeps will compensate for unavoidable AP jitter.
Note: Current is proportional to differentiated voltage. Thus, we can measure the time lag between the peak dV/dt and action current using cursors 1 and 2.
Average the dV/dt and action current sweeps and measure the time delay between their peaks.
Repeat steps 1–2 for other files collected.
Plot the distance of the pipette 2 from the soma as a function of time delay (Figure 2).
Figure 2. Measuring the propagation velocity. Distance from the edge of the soma plotted against the delay of action current generation. Each dot corresponds to the mean delay to the onset of the somatic action potential (AP) at a given location in the same neuron shown in Figure 1. The red line is a linear fit of the data. Notice that the AP initiates in a region at ~35 μm from the soma and propagates forward with a velocity of 0.31 m/s.
Determine the pipette 2 location at which the time delay is maximal. Designate this location as the AP initiation zone. Note that the spatial precision of this measurement depends on the relative distances between the locations at which the action current has been measured.
All data points from sites proximal to the AP initiation zone correspond to time delays of the backpropagating AP. All points distal to the AP initiation zone are latencies of the forward propagating AP.
The slope of the linear fit of these datasets gives a velocity of back- and forward-propagating AP.
Acknowledgments
This research was supported by The Israel Science Foundation (grant No. 1384/19). This protocol was described and validated in the following papers: Baranauskas et al. (2013), Lezmy et al. (2017), and Kotler et al. (2023).
Competing interests
The authors declare no competing interests.
Ethics
This study was carried out at the Ben-Gurion University of the Negev in accordance with the recommendations of guidelines for the welfare of experimental animals. The Institutional Animal Care and Use Committee of Ben-Gurion University approved animal experiments.
References
Baranauskas, G., David, Y. and Fleidervish, I. A. (2013). Spatial mismatch between the Na+ flux and spike initiation in axon initial segment. Proc. Natl. Acad. Sci. U.S.A. 110(10): 4051–4056.
Bernstein, J. (1868). Ueber den zeitlichen Verlauf der negativen Schwankung des Nervenstroms. Pflüger, Archiv für die Gesammte Physiologie des Menschen und der Thiere 1(1): 173–207.
Fleidervish, I. A., Lasser-Ross, N., Gutnick, M. J. and Ross, W. N. (2010). Na+ imaging reveals little difference in action potential–evoked Na+ influx between axon and soma. Nat. Neurosci. 13(7): 852–860.
Helmholtz, H. (1850). Vorläufiger Bericht über die Fortpflanzungsgeschwindigkeit der Nervenreizung. Archiv für Anatomie, Physiologie und Wissenschaftliche Medizin: 71–73.
Hodgkin, A. L. and Huxley, A. F. (1952). A quantitative description of membrane current and its application to conduction and excitation in nerve. J. Physiol. 117(4): 500–544.
Kole, M. H., Letzkus, J. J. and Stuart, G. J. (2007). Axon Initial Segment Kv1 Channels Control Axonal Action Potential Waveform and Synaptic Efficacy. Neuron 55(4): 633–647.
Kotler, O., Khrapunsky, Y., Shvartsman, A., Dai, H., Plant, L. D., Goldstein, S. A. and Fleidervish, I. (2023). SUMOylation of NaV1.2 channels regulates the velocity of backpropagating action potentials in cortical pyramidal neurons. eLife 12: e81463.
Lezmy, J., Lipinsky, M., Khrapunsky, Y., Patrich, E., Shalom, L., Peretz, A., Fleidervish, I. A. and Attali, B. (2017). M-current inhibition rapidly induces a unique CK2-dependent plasticity of the axon initial segment. Proc. Natl. Acad. Sci. U.S.A. 114(47): E10234–E10243.
Palmer, L. M. and Stuart, G. J. (2006). Site of Action Potential Initiation in Layer 5 Pyramidal Neurons. J. Neurosci. 26(6): 1854–1863.
Popovic, M. A., Foust, A. J., McCormick, D. A. and Zecevic, D. (2011). The spatio-temporal characteristics of action potential initiation in layer 5 pyramidal neurons: a voltage imaging study. J. Physiol. 589(17): 4167–4187.
Sasaki, T., Matsuki, N. and Ikegaya, Y. (2012). Targeted axon-attached recording with fluorescent patch-clamp pipettes in brain slices. Nat. Protoc. 7(6): 1228–1234.
Shu, Y., Duque, A., Yu, Y., Haider, B. and McCormick, D. A. (2007). Properties of Action-Potential Initiation in Neocortical Pyramidal Cells: Evidence From Whole Cell Axon Recordings. J. Neurophysiol. 97(1): 746–760.
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Neuroscience > Neuroanatomy and circuitry
Biophysics > Electrophysiology > Patch-clamp technique
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This is a correction notice. See the corrected protocol.
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Correction Notice: A Simple Microplate Assay for Reactive Oxygen Species Generation and Rapid Cellular Protein Normalization
NN Neville S. Ng
LO Lezanne Ooi
Published: Oct 5, 2023
DOI: 10.21769/BioProtoc.4877 Views: 278
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After publication in Bio-protocol (https://bio-protocol.org/e3877), we amended the following corrections:
•"Empty plate into an appropriate corrosive waste container and wash with 200 μL of 1% acetic acid. A second wash may be required": the font text in the generated PDF was too large; it has now been changed.
•"The period of time to allow for cell adherence post-seeding can be confirmed by microscopy to avoid 0.5 h" should read "The period of time to allow for cell adherence post-seeding can be confirmed by microscopy to avoid the full 0.5 h room temperature incubation time."
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Generation of Mouse Primitive Endoderm Stem Cells
YO Yasuhide Ohinata
AS Atsunori Saraya
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Reviewed by: Komuraiah MyakalaWei Fanabhijnya kanugovi
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The authors used this protocol in Science Feb 2022
Abstract
The blastocysts consist of dozens of cells of three distinct lineages: epiblast (Epi), trophoblast (TB), and primitive endoderm (PrE). All embryonic and extraembryonic tissues are derived from Epi, TB, and PrE. Stem cell lines representing preimplantation Epi and TB have been established and are known as embryonic stem cells (ESCs) and trophoblast stem cells (TSCs). Extraembryonic endoderm cells (XENCs) constitute a cell line that has been established from PrE. Although in vivo, PrE gives rise to visceral endoderm (VE), parietal endoderm (PE), and marginal zone endoderm (MZE); XENCs, on blastocyst injection into chimeras, primarily contribute to the distal region of PE. Here, we provide a comprehensive protocol for the establishment of fully potent primitive endoderm stem cell (PrESC) lines. PrESCs are established and maintained on mouse embryonic fibroblast (MEF) feeder cells in a serum-free medium supplemented with fibroblast growth factor 4 (FGF4), heparin, CHIR99021, and platelet-derived growth factor-AA (PDGF-AA). PrESCs co-express markers indicative of pluripotency and endoderm lineage commitment, exhibiting characteristics akin to those of PrE. On transplantation of PrESCs into blastocysts, they demonstrate a high efficiency in contributing to VE, PE, and MZE. PrESCs serve as a valuable model for studying PrE, sharing similarities in gene expression profiles and differentiation potential. PrESCs constitute a pivotal cornerstone for in vitro analysis of early developmental mechanisms and for studies of embryo reconstitution in vitro, particularly in conjunction with ESCs and TSCs.
Key features
• Establishment and maintenance of primitive endoderm stem cell (PrESCs) capable of recapitulating the developmental prowess inherent to PrE.
• Offering a source of PrE lineage for embryo-like organoid reconstitution studies.
Graphical overview
Keywords: Primitive endoderm Primitive endoderm stem cells Blastocyst Extraembryonic endoderm Yolk sac Blastocyst complementation
Background
Epiblast (Epi), trophoblast (TB), and primitive endoderm (PrE) are differentiated from the zygote by the late blastocyst stage of preimplantation development. Epi lineage generates all embryonic tissues, extraembryonic mesodermal tissues (i.e., chorion, allantois, yolk sac mesoderm, amniotic mesoderm, and capillaries in the labyrinthine layer of the placenta), and amniotic ectoderm. TB lineage generates major parts of placenta and trophectoderm of parietal yolk sac (PYS). PrE lineage is vital for normal embryonic development and is the origin of the visceral endoderm (VE) layer of the visceral yolk sac (VYS), the parietal endoderm (PE) layer of the PYS, and the marginal zone endoderm (MZE) lining along the edge of the placenta/chorionic plate of the placental disk (Gardner, 1982; Igarashi et al., 2018). The extraembryonic endodermal tissues play a pivotal role in nourishing the embryo by facilitating nutrient provision at the maternal–fetal interface (Jollie, 1990; Lloyd, 1990), especially significant prior to the onset of placental circulation (~E10.5 in mice), anterior–posterior patterning (Brennan et al., 2001), and yolk sac hematopoiesis (Ueno and Weissman, 2010). In mice, the gestation period is 19–21 days, and the initial yolk sac is formed on the fourth day, so embryonic development is uniquely dependent on the yolk sac during the first quarter of development.
Stem cell lines representing preimplantation epiblasts and trophoblasts have been established and are known as embryonic stem cells (ESCs) (Evans and Kaufman, 1981; Nichols et al., 2009) and trophoblast stem cells (TSCs) (Tanaka et al., 1998; Ohinata and Tsukiyama, 2014), respectively. To gain further insight into how the PrE functions during pre- and early post-implantation development, establishment of stem cell lines that fully retain PrE developmental potential has long been awaited. Extraembryonic endoderm cells (XENCs) turned out to be derivatives of the PrE but could only contribute to the distal region of the PE in chimeras (Kunath et al., 2005). Efforts to improve XEN cell culture conditions enabled the generation of XEN-P (extraembryonic endoderm precursor) cells in rats (Debeb et al., 2009; Galat et al., 2009) and pXEN (primitive XEN) cells in mice (Zhong et al., 2018), which express pluripotent makers and endoderm markers simultaneously. XEN-P cells have a slightly improved ability to contribute to cells other than in the distal PE at gastrula stages in chimeras. Further improvement of XENCs was performed by treating them with Nodal or Cripto, a ligand and coreceptor in the nodal signaling, respectively, but the impact of this treatment was again limited (Julio et al., 2011). Therefore, a robust protocol to capture early PrE-derived cells that can generate all derivatives of the PrE in the chimeric conceptus awaits discovery.
Recently, we succeeded in establishing primitive endoderm stem cells (PrESCs) that can fully recapitulate the developmental potential of PrE in mice. PrESCs are established and maintained on MEF feeder cells in a serum-free medium supplemented with FGF4, heparin, CHIR99021, and PDGF-AA. PrESCs co-express markers indicative of pluripotency (i.e., OCT4 and E-Cadherin) and endoderm lineage commitment (i.e., GATA4, GATA6, and SOX17), exhibiting characteristics akin to those of PrE. On transplantation of PrESCs into blastocysts, they demonstrate a high efficiency in contributing to all PrE derivatives, VE, PE, and MZE (Ohinata et al., 2022). In this protocol, we describe the detailed procedure for efficient generation of PrESC lines in mice. PrESCs therefore represent not only a regenerative resource to complement the PrE, but also an intervention tool to elucidate the mechanisms of PrE specification and subsequent pre- and post-implantation development. Importantly, PrESCs can be the final component of stem cell line assemblage required to generate development-competent blastocysts in vitro, in collaboration with ESCs and TSCs. PrESCs will contribute to the opening of a new research field linking pre- and early post-implantation development.
Materials and reagents
Biological materials
Pregnant mouse, 3.5 days post coitum (dpc) (8–20 weeks old) (e.g., CLEA Japan, CD-1). We have confirmed this protocol in CD-1, BALB/c, and B6PWKF1 genetic backgrounds
Mouse embryonic fibroblasts (MEFs) (ATCC, SCRC-1045TM). MEFs can also be derived from DR4 mice [Jackson, Dnmt1tm3Jae Hprt1b-m3 Tg (pPWL512hyg) 1Ems/J (DR4)] that are dissected 14.5 dpc; cells are passaged five times before inactivation by mitomycin C treatment. We routinely use DR4 MEFs, but the widely used CD-1 MEFs can also be used
Human fibroblast growth factor 4 (FGF4) (Sigma, catalog number: F8424)
Human platelet-derived growth factor-AA (PDGF-AA) (R&D, catalog number: 221-AA)
Rabbit anti-OCT4 antibody (MBL, catalog number: PM048)
Goat anti-SOX17 antibody, stock solution: 10 mg/mL in PBS (R&D, catalog number: AF1924)
Donkey anti-rabbit IgG antibody Alexa 488 conjugated (Invitrogen, catalog number: A21206)
Donkey anti-goat IgG antibody Alexa 568 conjugated (Invitrogen, catalog number: A11057)
Reagents
Mitomycin C (MMC) (Sigma, catalog number: M4287)
Fetal bovine serum (FBS) (biosera, catalog number: FB-1380/500)
Dulbecco’s modified Eagle’s medium (DMEM) (Sigma, catalog number: D5796)
β-mercaptoethanol (100×) (Millipore, catalog number: ES-007-E)
Penicillin-streptomycin (100×) (Gibco, catalog number: 15140-122)
Gelatin from porcine skin (Sigma, catalog number: G1890)
TrypLETM Select Enzyme (1×), no phenol red (Gibco, catalog number: 12563-029)
StemFit® AK02N (or StemFit® Basic02, StemFit® Basic03) (Ajinomoto)
Heparin (Sigma, catalog number: H3149)
CHIR99021, GSK3 inhibitor (REPROCELL, catalog number: 04-0004)
CELLBANKER® 1 plus (ZENOGEN PHARMA)
Ethanol (e.g., Nacalai Tesque, catalog number: 14712-34)
Paraformaldehyde (PFA) (TAAB, catalog number: P001)
Tween 20 (Chem Cruz, catalog number: sc-29113)
Normal donkey serum (Sigma, catalog number: D9663)
Hoechst 33342 10 mg/mL solution (Invitrogen, catalog number: H3570)
Solutions
MEF medium (see Recipes)
AK02N medium (see Recipes)
PrESC medium (see Recipes)
100× Mitomycin C stock solution (see Recipes)
0.2% Gelatin (see Recipes)
70% Ethanol (see Recipes)
PBST (see Recipes)
10 N NaOH (see Recipes)
4% PFA/PBST (see Recipes)
Recipes
MEF medium
Reagent Final concentration Quantity
DMEM 500 mL
FBS 10% 55 mL
β-mercaptoethanol 1× 5.6 mL
Penicillin-streptomycin 1× 5.6 mL
Total n/a 566.2 mL
AK02N medium
Reagent Quantity
Liquid A 400 mL
Liquid B 100 mL
Penicillin-streptomycin 5 mL
Total 505 mL
PrESC medium
Reagent Final concentration Quantity
AK02N medium (liquid A+B) 50 mL
CHIR99021 10 μM 50 μL (10 mM stock solution)
Human FGF4 50 ng/mL 50 μL (50 μg/mL stock solution)
Heparin 10 μg/mL 50 μL (50 mg/mL stock solution)
Human PDGF-AA 25 ng/mL 50 μL (25 μg/mL stock solution)
Total 50 mL
100× Mitomycin C stock solution
Reagent Final concentration Quantity
Mitomycin C 0.5 mg/mL 5 mg
ddH2O n/a 10 mL
Total n/a 10 mL
0.2% Gelatin
Reagent Final concentration Quantity
Gelatin 0.2% 1 g
ddH2O n/a 500 mL
Total n/a 500 mL
Sterilize 0.2% gelatin solution by autoclaving.
70% Ethanol
Reagent Final concentration Quantity
Ethanol 70% 350 mL
ddH2O n/a 150 mL
Total n/a 500 mL
PBST
Reagent Final concentration Quantity
Tween 20 0.2% 1 mL
PBS n/a 500 mL
Total n/a 501 mL
10 N NaOH
Reagent Final concentration Quantity
NaOH 0.4 g/mL 40 g
ddH2O n/a 100 mL
Total n/a 100 mL
NaOH liberates thermal energy upon dissolution. It is prudent to refrain from dissolving it in its entirety in a single instance. Instead, endeavor to dissolve it incrementally in multiple aliquots before culminating in a final volume of 100 mL. Employ safety spectacles to shield your eyes.
4% PFA/PBST
Reagent Final concentration Quantity
PFA 4% 2 g
PBST n/a 50 mL
10 N NAOH n/a 10 μL
Total n/a 50 mL
Incubate at 65 °C and mix several times until the PFA melts.
Equipment
Conical tube (15 mL) (Corning, catalog number: 352196)
Conical tube (50 mL) (Corning, catalog number: 352098)
Tissue culture plate (24 well) (Corning, catalog number: 351147)
Tissue culture plate (12 well) (Corning, catalog number: 351143)
Tissue culture plate (6 well) (Corning, catalog number: 351146)
Culture dish (35 mm) (Thermo Scientific, catalog number: 150460)
Culture dish (150 mm) (Corning, catalog number: 353025)
Petri dish (60 mm) (Corning, catalog number: 351007)
Glass bottom culture dish (35 mm) (IWAKI, catalog number: 3910-035)
Cryotube vial (Thermo Scientific, catalog number: 377224)
P2 micro-pipette (e.g., Gilson, catalog number: F144054M)
P20 micro-pipette (e.g., Gilson, catalog number: F144056M)
P1000 micro-pipette (e.g., Gilson, catalog number: 144059M)
Dissection microscope (e.g., Evident, model: SZX7)
Inverted microscope (e.g., Evident, model: IX71)
Water bath (e.g., TAITEC, model: SJ-07N)
Centrifuge (e.g., TOMY, model: LCX-100)
Hemocytometer (e.g., Biomedical Science, model: BMS-OCC01)
CO2 incubator (e.g., Panasonic, model: MCO-170AICV-PJVH)
Scissors (e.g., Natsume, model: NAPOX B-68T)
Forceps (e.g., Inox-Med, L5, catalog number: 11253-27)
Syringe (5 mL) (e.g., Terumo, model: SS-05LZ)
21-gauge needle (e.g., Terumo, model: NN-2138R)
Freezing container (e.g., Nihon Freezer, model: BICELL)
Ultra-low temperature freezer (-80 °C) (e.g., Panasonic, model: MDF-DU502VHS1-PJ)
Ultra-low temperature freezer (-150 °C) (e.g., Panasonic, model: MDF-1156ATA-PJ)
Confocal microscope (e.g., Evident, model: FV1200)
Procedure
Mitotic inactivation of MEF: mitomycin C (MMC) treatment
Notes:
The size and number of culture dishes used depends on the scale of the experiment. We routinely use 10–20 150 mm dishes. This protocol describes the use of 150 mm dishes.
MMC is toxic, so be sure to wear gloves when performing the experiment.
Culture MEFs in 25 mL of MEF medium per dish (see Recipe 1). Beginning with a confluent layer of MEFs, remove MEF medium from dishes and replace with 5 μg/mL 1× Mitomycin C solution in MEF medium.
Incubate the dishes for 2 h at 37 °C with 5% CO2.
Remove the medium and wash the cells three times with 25 mL of PBS.
Add 10 mL of 0.05% trypsin-EDTA and incubate until the cells come off the dish (3–5 min).
Collect cells in 50 mL conical tubes. Conical tubes are pre-filled with 10 mL of MEF medium to inhibit trypsin.
Centrifuge the tubes at 300× g for 5 min at room temperature and aspirate the supernatant.
Resuspend the cell pellet in 25 mL of MEF medium. Centrifuge the tubes at 300× g for 5 min at room temperature and aspirate the supernatant. Repeat this step once.
Resuspend the cell pellet in CELLBANKER® 1 plus. Count the cells using a hemocytometer, dilute the cells with CELLBANKER® 1 plus to a concentration of 5 × 106 cells/mL, and aliquot 1 mL into each cryotube vial.
Freeze the cells in a -80 °C ultra-low temperature freezer by using a BICELL. By using BICELL, the temperature of cryotubes can be gradually lowered.
Mouse mating to obtain blastocysts
Both male and female mice are sexually mature at eight weeks, being able to mate and give birth. One male should be kept in each cage for mating. In the evening, select females in estrus with large vulvae and keep them with one male and one or two females per cage.
The next morning, check female mice. If they have mated, their vagina is filled with the copulatory plug. The noon of the plug confirmation day is defined as 0.5 dpc.
Separate plugged females from the male and keep them in an individual cage until being used for dissection.
Preparation of MEF feeders
Day -1. One day before starting the derivation, coat 24-well plate with 0.2% (weight/vol) gelatin (see Recipe 5) (200 μL/well) for 15 min at room temperature. Aspirate the gelatin solution before seeding the feeder cells (Figure 2A).
Rapidly melt a frozen vial of mitotically inactivated MEFs in a 37 °C water bath until thawed and transfer the cells into a 15 mL centrifuge tube containing 5 mL of MEF medium. Centrifuge the tube at 300× g for 5 min at room temperature and aspirate the supernatant.
Resuspend the cell pellet in MEF medium. Plate at a density of 5 × 106 cells per plate of a 24-well plate, 12-well plate, 6-well plate, or six 35 mm dishes in MEF medium, and then incubate the plates overnight at 37 °C with 5% CO2.
Recovery of blastocyst-stage embryos from uterus
Notes:
Use scissors and forceps that have been sterilized in an autoclave.
StemFit® AK02N comprises liquids A, B, and C. Liquid C remains unutilized in the establishment of PrESCs. Liquids A and B are entirely identical to StemFit® Basic02. StemFit® Basic03 serves as a viable substitute for AK02N. StemFit® AK02N, Basic02, and AK03N are exclusively accessible within Japan, whereas StemFit® Basic03 boasts global availability.
Day 0. Thoroughly clean an experiment bench and a dissection microscope with 70% ethanol.
Euthanize a pregnant mouse at 3.5 dpc, dissect out the uterus, and place it on a clean paper towel or filter paper. Follow the facility’s rules for euthanasia. Remove fat tissues and cut away the ovary and oviducts (Figure 1A). Place the uterus in a fresh 2 mL drop of AK02N medium (see Recipe 2) in a 60 mm Petri dish.
Under a dissection microscope, with a 5 mL syringe, take 5 mL of the AK02N medium and attach a needle (21 gauge). While securing the uterine horn with forceps, stick the needle and flush the uterine horn with ~0.5 mL of AK02N medium in the direction of the ovary to the vagina and remove the uterine horn from the dish. Repeat the same process for the other uterine horn (Figure 1B).
Under higher magnification, find the flushed blastocysts in the drop and transfer them to a new drop of AK02N medium by using a P20 micro-pipette.
Figure 1. Recovery of blastocyst-stage embryos from uterus
PrESC derivation
Notes:
Steps E1–E5 can be executed under ambient conditions; however, it is imperative to manipulate the culture plate lid with utmost care and minimal frequency to avert potential contamination by microorganisms or particulate matter. Avoid prolonged exposure of the plate lid. Consider the advantageous inclusion of a dissection microscope within a sterile workbench.
StemFit® Basic03 medium by Ajinomoto serves as a viable substitute for AK02N. StemFit® AK02N, Basic02, and AK03N are exclusively accessible within Japan, whereas StemFit® Basic03 boasts global availability.
The survival rate of MMC-treated MEFs following freeze-thaw exhibits variability; it is imperative to ensure comprehensive coverage of the well’s bottom by MEFs. Employing MEFs with an insufficient density may precipitate differentiation into PrESCs.
The establishment of PrESCs should be precluded by the utilization of N2B27/Neurobasal/DMEMF12 (1:1) medium, a conventional choice in stem cell culture, as a replacement for the specialized AK02N.
The utilization of serum-containing media in lieu of AK02N will result in failure to establish PrESCs, accompanied by the loss of pluripotency marker expression and the emergence of XENC-like cells.
Day 0. Thoroughly clean an experiment bench and a dissection microscope with 70% ethanol.
The day before starting the derivation, aspirate the medium on the MEF feeders and replace it with 500 μL of PrESC medium (see Recipe 3) in the prepared 24-well plate (Figure 2A).
Under a dissection microscope, using P2 micro-pipette, place one blastocyst per each MEF-covered well in the plate and then incubate the plates at 37 °C with 5% CO2.
Day 5. Observe the plate under an inverted microscope. The blastocysts should have started to form an outgrowth (Figure 2B). Carefully aspirate the medium, change it with 500 μL of fresh PrESC medium, and return the plate to the incubator.
Day 7, passage 1. Observe the plate under an inverted microscope. The blastocysts should have formed an outgrowth. Carefully aspirate the medium and add 200 μL of TrypLETM Select to each well. Incubate the plate at 37 °C for 5 min.
Dissociate the outgrowth by pipetting into single cells using a P1000 pipette and transfer the cells into a 15 mL centrifuge tube containing 2 mL of AK02N medium. Centrifuge the tube at 300× g for 5 min at room temperature and carefully aspirate the supernatant as much as possible. Excessive carryover of TrypLETM Select adversely affects MEF after passage. Since the pellet after centrifugation is very small and fragile, gently aspirate and leave approximately 50 μL of medium together with the pellet.
Resuspend the cell pellet in 500 μL of PrESC medium, passage into one MEF-covered well in the prepared 24-well plate, and then incubate the plates overnight at 37 °C with 5% CO2.
Day 12. Observe the plate under an inverted microscope. Small colonies should be forming (Figure 2B and 2C). Carefully aspirate the medium and change it with 500 μL of fresh PrESC medium. When using StemFit® Basic03 medium, the morphology of the colony after the first passaging may be indistinct and difficult to see, but after the second passaging a well-defined colony becomes visible.
Figure 2. Primitive endoderm stem cell (PrESC) derivation from mouse blastocyst. A. Schedule for PrESC derivation. B. Outgrowth or colony morphology at each time point. C. Derivation of PrESCs in Basic03 medium. Scale bar, 100 μm.
Day 14, passage 2. Carefully aspirate the medium and add 200 μL of TrypLETM Select to each well. Incubate the plate at 37 °C for 5 min.
Dissociate the cells by pipetting into single cells by using a P200 pipette and transfer the cells into a 15 mL centrifuge tube containing 2 mL of AK02N medium. Centrifuge the tube at 300× g for 5 min at room temperature and aspirate the supernatant.
Resuspend the cell pellet in 1 mL of PrESC medium, passage into one MEF-covered well in the prepared 12-well plate, and then incubate the plate at 37 °C with 5% CO2.
Day 19. Observe the plate under an inverted microscope. PrESC colonies should be forming. Carefully aspirate the medium and change it with 1 mL of fresh PrESC medium.
Day 21, passage 3. Carefully aspirate the medium and add 200 μL of TrypLETM Select to each well. Incubate the plate at 37 °C for 5 min.
Dissociate the cells by pipetting into single cells by using a P1000 pipette and transfer the cells into a 15 mL centrifuge tube containing 2 mL of AK02N medium. Centrifuge the tube at 300× g for 5 min at room temperature and aspirate the supernatant.
Resuspend the cell pellet in 2 mL of PrESC medium, passage into MEF-covered wells in the prepared 6-well plate, and then incubate the plate at 37 °C with 5% CO2.
After PrESCs reach approximately 70% confluency, passage into MEF-covered wells at 1:5 to 1:10 dilution. Change medium every other day.
Cryopreservation
The viable rate of PrESCs after freeze-thaw is low, and cryopreservation must be at a higher density than for ESCs or TSCs. PrESCs have grown to 70% confluency in a 35 mm dish; aspirate the medium and add 1 mL of TrypLETM Select to each well. Incubate the plate at 37 °C for 5 min.
Dissociate the cells by pipetting into single cells by using a P1000 pipette and transfer the cells into a 15 mL centrifuge tube containing 5 mL of AK02N medium. Centrifuge the tube at 300× g for 5 min at room temperature and aspirate the supernatant.
Resuspend the cell pellet in 1 mL of CELLBANKER® 1 plus cryoprotective reagent and transfer into two cryotubes (500 μL/tube). Use of cryoprotective reagents that do not contain serum may significantly further reduce cell viability after thawing.
Freeze the cells in a -80 °C ultra-low temperature freezer by using a BICELL. By using BICELL, the temperature of cryotubes can be gradually lowered.
For long-term storage, store the tubes after freezing in a -150 °C ultra-low temperature freezer.
Checking gene expression of PrESCs markers
PrESCs co-express pluripotency and endodermal markers (Ohinata, Endo et al., 2022). Check the gene expression by immunofluorescence staining for OCT4 (a pluripotency marker) and SOX17 (an endoderm marker) (Figure 3).
Figure 3. Co-expression of a pluripotent marker (OCT4) and an endoderm marker (SOX17) in primitive endoderm stem cell (PrESCs). Scale bar, 100 μm.
Culture PrESCs in a MEF-covered 35 mm glass-bottom cell culture dish.
When PrESCs reach desired cell density (usually approximately 70% confluency recommended), aspirate medium, wash twice with 2 mL of PBS, and fix with 2 mL of 4% PFA/PBST at room temperature for 30 min.
Wash cells three times in 2 mL of PBST (see Recipe 7) at room temperature for 15 min.
Block cells in 2 mL of 0.5% normal donkey serum (NDS)/PBST at room temperature for 30 min.
React cells with 1 mL of primary antibody solution [rabbit anti-OCT4 (1:1,000), goat anti-SOX17 (1:1,000, 10 mg/mL stock solution) in 0.5% NDS/PBST] at room temperature for 30 min.
Wash cells three times in 2 mL of PBST at room temperature for 15 min.
React cells with 1 mL of secondary antibody solution [donkey anti-rabbit IgG Alexa 488 (1:1,000), donkey anti-goat IgG Alexa 568 (1:1,000), Hoechst 33342 (5 μg/mL) in PBST] at room temperature for 30 min.
Wash cells three times in 2 mL of PBST at room temperature for 15 min.
Observe cells with an inverted confocal microscope.
Validation of protocol
This protocol was used in the following paper:
Ohinata, Y. et al. (2022). Establishment of mouse stem cells that can recapitulate the developmental potential of primitive endoderm. Science 375(6580): 574-578.
Acknowledgments
This work was supported by JSPS KAKENHI under Grant Number 18H05366 and 19H05757 to Y.O.
This protocol was derived from the original work of Ohinata et al. (2022).
Competing interests
RIKEN has a patent pending (JP2019-118733) on the method for derivation and maintenance of PrESCs. Y.O. and H.K. are the inventors of the patent. There is no conflict of interest relevant to this study.
Ethical considerations
All animal experiments conformed to our guidelines for the care and use of laboratory animals and were approved by the Institutional Committee for Laboratory Animal Experimentation (RIKEN Center for Integrative Medical Sciences AEY2023-019(2) and Chiba University A5-002).
References
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Identification of Topologically Associating Domains (TADs) and Long-distance Enhancer–gene/gene–gene Interactions with Hi-C and HiChIP
XH Xianhui Huang
LZ Longfu Zhu
Xianlong Zhang
MW Maojun Wang
Published: Nov 20, 2023
DOI: 10.21769/BioProtoc.4879 Views: 192
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Abstract
The hierarchy featured architecture of 3D genome influences the expression of genes to regulate development and defense processes. Hi-C is one of the most widespread technologies to explore hierarchy 3D genome organizations, including compartment, topologically associating domains (TADs), and loops. HiChIP can accurately identify chromatin loops between cis-regulatory elements in non-coding regions and genes. At the same time, growing innovative methods can be used to identify different hierarchy architectures of 3D genome, but most analyses are performed with a combination of a variety of software. Therefore, a clear and complete pipeline is essential. We summarize the detailed usage of various tools for identifying the 3D genome structure using Hi-C and HiChIP, as well as for visualizing the final result. Using these software tools, we identify the TADs and loops from the Hi-C and HiChIP libraries in human cell lines and cotton plants. In brief, this pipeline will help researchers choose a suitable tool with less time cost.
Keywords: 3D genome Hi-C HiChIP TAD Loop Human Cotton
Background
The genetic diversity of organisms is the fundamental basis for forming complex and diverse global ecosystems. Adequate and comprehensive knowledge of genetic information will help us understand the nature of evolution more deeply and thus improve our living environment without affecting the survival of other species. The study of the interpretation of genetic information from multiple perspectives has been booming (Misteli, 2020; Purugganan and Jackson, 2021; Song et al., 2021; Zhang et al., 2021). Among them, the 3D genome has become a fascinating study field. It has been extensively used to comprehensively study how the hierarchy of 3D genome regulates biological processes such as evolution, development, and resistance to pathogens by combining 1D genomics, transcriptomics, and epigenomics (Fullwood et al., 2009; Lieberman-Aiden et al., 2009; Mumbach et al., 2017; Sun et al., 2018; Wang et al., 2018; Norrie et al., 2019; Zheng and Xie, 2019; Yang et al., 2022). So far, there is a growing body of literature on topologically associating domains (TADs) and loops, since they are finely architected to regulate changes in genetic information (Rao et al., 2014; Wang et al., 2018; Zheng et al., 2019; Espinola et al., 2021; Hoencamp, 2021). Hence, accurate identification of TAD and loop structures is essential for understanding how the hierarchy of 3D genome architecture regulates biological processes.
With the innovation in experimental methods of 3D genomic study and the advancement of sequencing technologies, a large number of software tools were developed to identify 3D genome structures (Ay et al., 2014; Wang et al., 2015; Forcato et al., 2017; Bhattacharyya et al., 2019; Yardımcı et al., 2019; Fernandez et al., 2020; Wolff et al., 2020; Mourad, 2022). They have contributed to a boom in the field of 3D genomic research. This also increases the difficulties for researchers to realize them (Forcato et al., 2017). Therefore, a complete order pipeline is very useful for scientific researchers. To achieve this, we summarized the processing of several software tools, including identifying TADs by TADLib and Juicer, inferring loops of Hi-C data by Fit-Hi-C and Juicer, and inferring loops of HiChIP data by FitHiChIP and hichipper. The integration of these software into a whole pipeline by combining HiC-Pro and Juicebox can provide final visualization results from raw reads (Figure 1). The pipeline provides a comprehensive 3D genome analysis process that is applicable not only to Hi-C datasets but also compatible with HiChIP datasets. Moreover, the pipeline eliminates the need for complex intermediate steps in multiple software, providing users a simple and efficient experience while preserving the software's customizable function. In summary, the pipeline provides user-friendly features that are especially beneficial for researchers who are new to 3D genome analysis.
Figure 1. Flowchart for the identification of topologically associating domains (TAD) and loops from the datasets of Hi-C and HiChIP by different software
Software and datasets
Software
Python (Version 3.9.5/Version 2.7.18, https://www.python.org/downloads/) (2020/09)
R (Version 4.0.0, https://cran.r-project.org/bin/windows/base/old/) (2020/09)
Bowtie2 (Version 2.4.4) (Langmead and Salzberg, 2012)
BWA (Version 0.7.17) (Li and Durbin, 2009)
SAMtools (Version 1.9) (Li et al., 2009)
BEDTools (Version 2.27) (Quinlan and Hall, 2010)
MACS2 (Version 2.1.1) (Zhang et al., 2008)
HiC-Pro (Version 2.11.4) (Servant et al., 2015)
Juicer (Version 1.6) (Durand et al., 2016)
Juicer tools jar (Version 1.22.01, https://github.com/aidenlab/juicer/wiki/Download) (2020/11)
Juicebox (Version 1.11.08, https://github.com/aidenlab/Juicebox/wiki/Download) (2020/11)
TADLib (Version 0.4.1) (Wang et al., 2015)
HiCPeaks (Version 0.3.4, https://github.com/XiaoTaoWang/HiCPeaks) (2021/04)
Fit-Hi-C (Version 2.0.8) (Ay et al., 2014)
FitHiChIP (Version 9.1) (Bhattacharyya et al., 2019)
hichipper (Version 0.7.7) (Lareau and Aryee, 2018)
Data
Library of Hi-C
Hi-C data of human GM12878 B-lymphoblastoid cells (Rao et al., 2014)
Hi-C data of cotton fiber of Gossypium barbadense 3–79 at 20 days post anthesis (DPA) (Pei et al., 2022)
Library of HiChIP
HiChIP data (H3K27ac) of human GM12878 B-lymphoblastoid cells (Mumbach et al., 2017)
Procedure
HiC-Pro (Servant et al., 2015)
The HiC-Pro is a pipeline for Hi-C data processing, from a raw Hi-C dataset to a normalized contact matrix by reads aligning, reads pairing, pairs dumping, and contact map building. The bowtie2 index, the restriction fragments after digestion, and chromosome size information are necessary files for processing HiC-Pro. Raw interaction matrix is used for subsequent analysis.
Construct bowtie2 index that will be used to reads alignment by bowtie2.
$ bowtie2-build reference_genome.fa prefix_of_output_file
Calculate the chromosome size that is needed to build Hi-C matrix.
$ samtools faidx reference_genome.fa
$ awk -v OFS = “\t” ‘{print $1, $2}’ reference_genome.fa.fai > chromosome_size.bed
Create enzyme fragment file to get Hi-C fragment information.
$ python HiC-Pro/bin/utils/digest_genome.py reference_genome.fa -r mboi/dpnii/bglii/hindiii -o output_file_name (python 2.7.18)
Rebuild the config file of HiC-Pro that adds the path of bowtie2 index, chromosome size, and fragment of enzymatic digestion.
$ BOWTIE2_IDX_PATH = “The path of bowtie2 index produced in step 1”
$ GENOME_SIZE = “The file of single chromosome size produced in step 2”
$ GENOME_FRAGMENT = “The file of fragments produced in step 3”
Run HiC-Pro on a personal computer in a standalone model.
$ HiC-Pro -i path_input_file (raw reads) -o path_output_file -c you_config_file
Run HiC-Pro on a cluster.
$ HiC-Pro -i path_input_file (raw reads) -o path_output_file_path -c you_config_file -p
$ bsub < HiCPro_step1.sh # Parallel workflow
$ bsub < HiCPro_step2.sh # Merge all outputs in single thread
Juicer (Durand et al., 2016)
Juicer is a platform that integrates producing Hi-C matrix from Hi-C library, identifying compartment (Eigenvector), presuming TAD (Arrowhead), and inferring loops (HiCCUPS). Juicer aligns reads to the reference genome using BWA and requires an enzyme cut site file.
Construct BWA index files.
$ bwa index reference_genome.fa -p prefix_of_output_file
Calculate the size of a single chromosome.
$ samtools faidx reference_genome.fa
$ awk -v OFS = “\t” ‘{print $1, $2}’ reference_genome.fa.fai > chromosome_size.bed
Generate an enzyme cut sites file. The enzymes available include HindIII, DpnII, MobI, Sau3AI, and Arima. Also, you can add other enzyme cut fragments in the python file.
$ python juicer/misc/generate_site_positions.py DpnII prefix_of_output_file reference_genome.fa (python 3.9.5)
The “rawdata,” “references,” “restriction_sites,” and “scripts” folders should preferably exist in the working directory. The rawdata folder includes the “fastq” folder where the Hi-C library is stored. The references folder includes bwa index, genome, and chromosome size. The enzyme cut sites file is stored in the restriction_sites folder. The scripts folder includes all scripts that need to be run. Copy the juicer/cpu/common folder into the scripts folder.
$ juicer/cpu/juicer.sh -d full_path_of_fastq_file -z reference_genome.fa -g genome_name -D workdir -p chromosome_size -y full_path_of_enzyme_cut_site -t number_of_threads
Identify TAD by Juicer with arrowhead algorithm. Users should download juicer_tools.jar and put it in the scripts folder.
$ java -jar scripts/juicer_tools.jar arrowhead -m size_of_the_sliding_window (must be an even number) -r resolution -k normalization_method (NONE/VC/VC_SQRT/KR) --threads number_of_threads *.hic_file directory_of_output
Medium resolution maps: -m 2,000 -r 10,000 -k KR. High resolution maps: -m 2,000 -r 5,000 -k KR.
Infer loops from a .hic file data by using HiCCUPS algorithm.
$ java -jar scripts/juicer_tools.jar hiccups --cpu -r resolution -f values_of_FDR -p width_of_peak -i width_of_window -d distances_used_for_merging_nearby_pixels -t thresholds_for_merging_loop -k normalizations_methods (NONE/VC/VC_SQRT/KR) --threads number_of_threads *.hic_file directory_of_output
Convert the result of HiC-Pro to *.hic file.
$ java -Xmx2g -jar juicebox_tools.jar pre -r resolution *.VaildPairs *.hic genome_name
The memory can be expanded by adjusting the parameter -Xmx2g to a desired value, such as -Xmx500g,whicht can allocate 500 gigabytes of memory.
TADLib (Wang et al., 2015)
TADLib uses a machine-learning approach to predict TAD structure after training the model. The contact matrix produced by HiC-Pro is used as an input file of TADLib after conversion to a cool file format.
Convert result files of HiC-Pro to the input file of TADLib. The intrachromosomal interactions are used to identify TAD structures by TADLib, so they must be selected.
Choose single chromosome interaction matrix.
$ awk -v OFS=“\t” ‘{if(($1>=chr_bin_start)&&($1<=chr_bin_end)&&($2>=chr_bin_start)&&($2<=chr_bin_end)) print $1-chr_bin_start+1,$2-chr_bin_start+1,$3}’ matrix_file > result_file (The format of file name: ChromosomeNumber_ChromosomeNumber.txt)
Merge all single chromosome interaction matrices.
$ toCooler -O output_filename.cool -d meta_file --chromsizes-file reference_ChrSize.bed --no-banlance
The format of meta file.
res: The size of resolution
rep1: path_of_single_chromosome_interaction_matrix_of_repeat1
rep2: path_of_single_chromosome_interaction_matrix_of_repeat2
Identify TAD by TADLib.
$ hitad -O output_TAD_filename.bed -d meta_file --logFile hitad.log -p number_of_threads -W RAW --maxsize Largest_TAD_size
The format of meta file.
res: The size of resolution
rep1: cool_file::The size of resolution
rep2: cool_file::The size of resolution
Fit-Hi-C (Ay et al., 2014)
Fit-Hi-C can infer significant interaction loops by balancing distance, strength, and probability of two interaction loci. The input file of Fit-Hi-C from the interaction matrix is produced by HiC-Pro.
Transform of HiC-Pro result file to Fit-Hi-C input file.
$ python Fithic/hicpro2fithic.py -i filename.matrix -b filename_abs.bed -r resolution_of -o output_file_directory -n prefix_of_output_file (python 3.9.5)
Infer loops by Fit-Hi-C.
$ fithic -f fragment -i interaction -r resolution -L lowerbound -U upperbound -p number_of_spline_passes -o output_file_directory -l prefix_of_output_file
FitHiChIP (Bhattacharyya et al., 2019)
FitHiChIP infers significant cis interactions from a given HiChIP/PLAC-seq experiment. In addition to calling peaks of histone modification sites from the HiChIP library, it can also combine existing peak data. For calling peaks from the HiChIP library, *.ValidPairs, *.DEPairs, *.REPairs, and *.SCPairs files generated by HiC-Pro are required. At the same time, the available pairwise interaction file generated by HiC-Pro is required as the input file of FitHiChIP. In addition, chromosome size information is also needed.
Run HiC-Pro to produce the available pairwise interaction file.
Call peaks in the HiChIP library by FitHiChIP.
$ sh FitHiChIP/Imp_Scripts/PeakInferHiChIP.sh -H Path_of_HiC-Pro -D Path_of_output -R Chromosome_size -M Parameters_of_macs2
Path of HiC-Pro include *.ValidPairs, *.DEPairs, *.REPairs, *.SCPairs and *.ValidPairs files.
Modify the config file of FitHiChIP.
$ ValidPairs = The valid pairs result file produced by HiC-Pro
$ PeakFile = File containing reference ChIP-seq/HiChIP peaks
$ OutDir = Path of the result
$ ChrSizeFile = Chromosome size files for reference genomes
$ BINSIZE = size of the bins
$ QVALUE = FDR value
$ UseP2Pbackgrnd = Two possible values 0 or 1. 0 represent loose background, 1 represent stringent background.
$ BiasType = Can be 1 (coverage bias) or 2 (ICE bias).
Run FitHiChIP.
$ bash FitHiChIP_HiCPro.sh -C config_file
hichipper (Lareau and Aryee, 2018)
The hichipper package can be used to determine library quality, identify and characterize DNA loops, and interactively visualize loops. This package takes the output from a HiC-Pro run. Similar to FitHiChIP, hichipper can also be used to call peaks from the HiChIP library or select an existing peak file.
Run HiC-Pro to produce some input files of hichipper.
Modify the configuration file that call peaks of histone modification sites in the HiChIP library by hichipper. Configuration parameters are written in the .yaml file. hichipper aggregates reads density from either all samples (COMBINED) or each sample (EACH) individually. Additionally, users can specify whether all reads density (ALL) is used or whether only self-ligation reads are used (SELF). The result path of HiC-Pro includes *.DEPairs, *.DumpPairs, *.RSstat, *.SCPairs, *.SinglePairs, and *.ValidPairs files.
The format of config.
peaks:
- COMBINED/EACH, ALL/ SELF
resfrags:
- The file of the enzyme section
hicpro_output:
- The path of the HiC-Pro output file
Modify the config file to identify loops by hichipper.
Peaks:
- The file of peaks
resfrags:
- The file of the enzyme section
hicpro_output:
- The path of the HiC-Pro output file
Run hichipper to identify loops of HiChIP data.
$ hichipper –out path_output_file config_file
Data analysis
Result interpretation
By using the above parameters of Juicer, TADs and loops were identified from the GM12878 library. The result is generally consistent with previous studies (Figure 2A). TADLib and Fit-Hi-C were used to identify TADs and loops in cotton. The results show that the loop interaction primarily occurs in TAD interior (Figure 2B). A comparison of the loops inferred by FitHiChIP and hichipper shows that most of them are the same, and a few differences may be due to different algorithms (Figure 2C).
Figure 2. Topologically associating domains (TAD) and loops were identified from Hi-C and HiChIP library by different software. A. The TAD and loop were identified by Juicer in GM12878 Hi-C library. The purple line and the blue point on the heat map represent TADs and loops identified in a previous study (Durand et al., 2016). The yellow and cyan lines on the heat map represent TADs and loops identified by Juicer. B. The TAD and loops were identified by TADLib and Fit-Hi-C in Gossypium barbadense 3-79 library. C. The loops of GM12878 HiChIP (H3K27ac) library were inferred by FitHiChIP and hichipper. The data of ChIP-Seq with different antibodies and RNA-Seq were from previous studies (Rao et al., 2014; Davis et al., 2018; Pei et al., 2022).
Acknowledgments
This work was supported by the National Key Research and Development Program of China (2021YFF1000100) and the National Natural Science Foundation of China (32170645, 31922069). Appreciation to all research scholars who participated in the software of pipeline development and provided experimental data.
Competing interests
The authors declare no competing interests.
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Supplementary information
Data and code availability: All data and code have been deposited to GitHub: https://github.com/Bio-protocol/TAD-and-loop-identification-workflows.git.
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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A New, Rapid Method for the Quantification of Dolichyl Phosphates in Cell Cultures Using TMSD Methylation Combined with LC–MS Analysis
DK Dipali Kale
TS Timo Sachsenheimer
AS Albert Sickmann
BB Britta Brügger
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4880 Views: 439
Reviewed by: Manjula MummadisettiRohit JainBhanu Jagilinki
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Original Research Article:
The authors used this protocol in Analytical Chemistry Feb 2023
Abstract
Dolichyl phosphates (DolP) are ubiquitous lipids that are present in almost all eukaryotic membranes. They play a key role in several protein glycosylation pathways and the formation of glycosylphosphatidylinositol anchors. These lipids constitute only ~0.1% of total phospholipids, and their analysis by reverse phase (RP) liquid chromatography–high-resolution mass spectrometry (LC–HRMS) is challenging due to their high lipophilicity (log P > 20), poor ionization efficiency, and relatively low abundance. To overcome these challenges, we have introduced a new approach for DolP analysis by combining trimethylsilyldiazomethane (TMSD)-based phosphate methylation and HRMS analysis. The analytical method was validated for its reproducibility, sensitivity, and accuracy. The established workflow was successfully applied for the simultaneous characterization and quantification of DolP species with different isoprene units in lipid extracts of HeLa and Saccharomyces cerevisiae cells.
Keywords: LC–HRMS LC-MS DolP Dolichyl phosphate TMSD Methylation Lipid analysis
Background
Dolichyl phosphates (DolP) are long-chain polyisoprenoid phosphates that function as glycan carriers in cellular N- and O-glycosylation, C- and O-mannosylation, and glycosylphosphatidylinositol-anchor biosynthesis (Orlean, 1990; Rosenwald et al., 1990). Numerous studies have reported altered endogenous DolP levels in pathophysiological conditions such as Alzheimer’s disease, dementia, and Prion disease (Kranz et al., 2007; Park et al., 2014; Sabry et al., 2016; Jaeken et al., 2020). Given the essential role of DolP in cellular glycosylation, there is a need for a rapid method to accurately characterize and measure DolP in biological samples, including those from patients with disorders associated with glycosylation defects.
Generally, DolP are quantified after their fluorescence derivatization by combining normal-phase liquid chromatography (LC) with fluorescence detection (Haeuptle et al., 2010). This method, based on a multi-step sample preparation and derivatization procedure, is labor intensive and time consuming. Few mass spectrometry (MS)-based methods exist for the structural characterization of endogenous DolP species (Löw et al., 1991; Wolucka et al., 1996; Guan et al., 2010; Taguchi et al., 2016). To date, a method for the simultaneous profiling and quantification of DolP species of different isoprene chain lengths in biological membranes has been lacking. We have developed a method for the characterization and quantification of DolPs using trimethylsilyldiazomethane (TMSD) derivatization followed by reverse phase (RP) liquid chromatography–high-resolution mass spectrometry (LC–HRMS) analysis. TMSD is frequently used as a methylating agent for anionic phospholipids (Lee et al., 2017) and free fatty acids (Mok et al., 2016) to increase the electrospray ionization efficiency. The scheme of the methylation reaction of DolP is illustrated in Figure 1.
Figure 1. Chemical reaction of trimethylsilyldiazomethane (TMSD)-based methylation of dolichyl phosphates (DolPs). The simplified linear structure of DolP is displayed.
We found that TMSD methylation enabled chromatographic retention of DolPs in RPLC analyses. In the Tandem MS (MS/MS or MS2) analysis of all methylated DolP species by high-energy collisional dissociation, the most intense fragment ion at m/z 127.0155 represents a characteristic headgroup fragment, corresponding to a dimethylphosphate group. Transitions from all parent m/z (Table 1) to 127.0155 in the MS2 spectra were used to determine the identification of methylated DolPs and dodecaprenyl phosphate (PolP C60).
Table 1. Molecular formulas, exact masses, and retention times of [M+NH4]+ of DolP species and the internal standard (IS) PolP C60
Analyte Molecular formula Exact mass [M+NH4]+ Retention time (min)
PolP C60(IS) C62H103O4P1 960.7932 19.1
DolP C65 C67H113O4P1 1030.8715 20.0
DolP C70 C72H121O4P1 1098.9341 20.6
DolP C75 C77H129O4P1 1166.9967 21.2
DolP C80 C82H137O4P1 1235.0593 21.8
DolP C85 C87H145O4P1 1303.1219 22.3
DolP C90 C92H153O4P1 1371.1845 22.9
DolP C95 C97H161O4P1 1439.2471 23.5
DolP C100 C102H169O4P1 1507.3097 24.1
DolP C105 C107H177O4P1 1575.372 24.7
The established workflow uses a fast and efficient single-step derivatization procedure. Furthermore, the method was used to quantify DolP species from lipid extracts of Saccharomyces cerevisiae and HeLa cells. In the future, this method can be used to study the role of DolP homeostasis in health and disease and how DolP availability may be regulated by microenvironmental factors such as nutrient availability. This method has been thoroughly validated for its quantification utility in Kale et al. (2023).
Materials and reagents
Pipette tips, 50–1,000 μL (Eppendorf, catalog number: 613-3505)
Gastight syringe, 5 mL (Hamilton, catalog number: 81517)
Pipette tips Combitips advanced 25 mL (Eppendorf, catalog number: EP 0030089472)
Round bottom threaded glass tubes (PYREX, catalog number: 9447161)
Pasteur pipettes, soda-lime glass (Brand, Wertheim, Germany, catalog number: 747715)
Micro inserts, Neochrom (Neolab, catalog number: 7-0635)
LC vials, Neochrom (Neolab, catalog number: EC-1184)
Threaded screw caps with Teflon Liner (Corning, catalog number: 9998-15)
Green nitrile gloves (TouchNtuff, catalog number: 92600)
Ammonium acetate (AmAc), eluent additive for LC-MS, LiChropur (Sigma-Aldrich, Merck KGaA, catalog number: 73594)
Formic acid (FA), eluent additive for LC-MS (Honeywell, catalog number: 56302)
Ammonium bicarbonate (Sigma-Aldrich, Merck KGaA, catalog number: 09830)
Acetonitrile, LC-MS grade (Fisher Chemical, catalog number: 10616653)
Methanol, LC-MS grade (Fisher Chemical, catalog number: 10532213)
Isopropanol, LC-MS grade, Optima (Fisher Chemical, catalog number: A461212)
Water, LC-MS grade, Optima (Fisher Chemical, catalog number: 10505904)
Dichloromethane (Sigma-Aldrich, Merck KGaA, catalog number: 270997)
Dolichol 13–21 phosphate mixture (DolP standard hereafter) (Sigma-Aldrich, Merck KGaA, catalog number: 900201X)
Dodecaprenyl phosphate (PolP C60) (CymitQuimica, catalog number: 48-62-1060)
Mini-BeadBeater q mill beads (0.5 mm) (BioSpec Products, catalog number: 11079105)
Potassium hydroxide pellets (KOH) (Sigma-Aldrich, Merck KGaA, catalog number: 21473)
Trimethylsilyldiazomethane (TMSD) (Sigma-Aldrich, Merck KGaA, catalog number: 362832)
Acetic acid ACS grade (Sigma-Aldrich, Merck KGaA, catalog number: 33209)
HeLa/Fibroblast cells (~1 M cells)
S. cerevisiae cells (~0.8 OD)
Acquity UPLC CSH C18 Column, 1.7 μm, 1 mm × 150 mm (Waters, catalog number: 186005294)
Acquity UPLC CSH C18 VanGuard Pre-column, 1.7 μm, 2.1 mm × 5 mm (Waters, catalog number: 186005303)
For the preparation of liquid chromatography, mobile phase A (Solvent A) and mobile phase B (Solvent B), and wash solvents, please see Recipes.
Solutions
15 M KOH (see Recipes)
10 M AmAc (see Recipes)
Solvent A: acetonitrile/water 60:40 (v/v) with 0.1 % FA and 10 mM AmAc (see Recipes)
Solvent B: isopropanol/water 90:10 (v/v) with 0.1 % FA and 10 mM AmAc (see Recipes)
Wash solvent: dichloromethane:methanol:water (3:48:47, v/v/v) (see Recipes)
Dichloromethane:methanol mixture (6.5:5.2, v:v) (see Recipes)
155 mM ammonium bicarbonate buffer (see Recipes)
Recipes
15 M KOH
Accurately weigh 8.4 g of KOH pellets into a 100 mL volumetric flask. Add LC-grade water dropwise with intermittent slow vortex to dissolve KOH. Be careful, as an exothermic reaction will occur, and the reaction flask will be hot. After complete dissolution, dilute the solution to the mark. Use a glass bottle with a PTFE lid for storage of the KOH solution.
10 M AmAc
Take the 10 mL volumetric flask along the glass stopper and rinse both twice with MS-grade methanol. Dry them carefully under Argon. Transfer accurately weighed 7.708 g of LC-MS grade AmAc to the volumetric flask. To dissolve the solid, add distilled water in steps of approximately 100 µL, followed by sonication. Once all solids are dissolved, fill the volumetric flask to the etched line. This stock can be stored at 4 °C for up to six months.
Solvent A: acetonitrile/water 60:40 (v/v) with 0.1% FA and 10 mM AmAc
Prepare 1,000 mL of solvent mixture of acetonitrile/water 60:40 (v/v) and add 1 mL of formic acid and 1 mL of 10 mM AmAc stock. Mix solvent mixture thoroughly and degas for 5 min in an ultrasonic water bath sonicator.
Solvent B: isopropanol/water 90:10 (v/v) with 0.1% FA and 10 mM AmAc
Prepare 1,000 mL of solvent mixture of isopropanol/water 90:10 (v/v) and add 1 mL of FA and 1 mL of 10 mM AmAc stock. Mix solvent mixture thoroughly and degas for 5 min in an ultrasonic water bath sonicator.
Wash solvent: dichloromethane:methanol:water (3:48:47, v/v/v)
Dichloromethane:methanol mixture (6.5:5.2, v:v)
155 mM ammonium biocarbonate buffer
Transfer accurately weighed 1.2 g of ammonium bicarbonate into a 100 mL volumetric flask. Dissolve in distilled water and make up to the mark with distilled water. This stock can be stored at 4 °C for up to six months.
Equipment
Eppendorf Multipette E3 (Eppendorf, catalog number: VB-1748)
Eppendorf Research plus pipette 100–1,000 μL (Eppendorf, catalog number: 3123000063)
Ultimate 3000 LC system (Dionex, Thermo Fisher Scientific) system coupled to a Q-Exactive HRMS (Thermo Scientific)
Vortexer, SI Vortex-Genie 2 (Scientific Industries, catalog number: 15547335)
Nitrogen evaporator (TurboVap, catalog number: 416200)
Syringe, gastight HamiltonTM1700 Series 100 μL (Sigma-Aldrich, MerckKGaA, catalog number: 111HAM201050)
Refrigerated centrifuge, Heraeus Megafuge 16R (Thermo Scientific, catalog number: 75004271)
Ultrasonic water bath sonicator (Bandelin sonorex)
Water bath, LSB Aqua Pro (Grant Instruments, catalog number: LSB12)
Software
XCalibur software 3.2 (Thermo Fisher Scientific)
Online EnviPat R package Tool (available at https://www.envipat.eawag.ch/, Accessed: 21 August 2019)
Procedure
A schematic of the workflow for the analysis of DolPs is shown in Figure 2. The different procedures involved in this workflow are mentioned below.
Figure 2. Overview of the developed workflow, including sample preparation, methylation, and reverse phase liquid chromatography–mass spectrometry (RPLC-MS) analysis of dolichyl phosphates (DolPs)
Note: Below mentioned steps should take place in a cold room or on ice where possible.
Thaw the biological samples, such as HeLa/fibroblast cells (~1 M cells) or S. cerevisiae cells (~0.8 OD), on ice for 20 min.
If performing Hela cells extraction, skip this step. Suspend S. cerevisiae cell pellets in 200 μL of 155 mM ammonium bicarbonate buffer. Add 0.5 mm glass beads to samples and vortex for 1 min. Then, centrifuge samples at 3,500× g for 1 min at room temperature (RT). Transfer the aqueous (top) layer to a clean microcentrifuge tube.
Add 20 pmol of internal standard (IS) PolP C60 to cell pellets/lysates.
Add 1 mL of methanol and vortex for 1 min at normal speed, followed by sonication for 1 min in an ultrasonic water bath sonicator.
Add 1 mL of water and vortex for 1 min at normal speed, followed by sonication for 1 min with an ultrasonic water bath sonicator.
Centrifuge at 3,500× g for 5 min at RT.
Note: If you wish to normalize DolP amounts to total lipid/total phosphatidylcholine (PC) (as bulk membrane lipid), measure the total lipid/PC levels of samples at this point. See Özbalci et al. (2013) for the procedure to determine the total lipid/PC levels of samples (Özbalci et al., 2013).
Procedure for alkaline hydrolysis and extraction
Note: Use round bottom screw-top glass tubes with Teflon-lined caps for DolP sample preparation. Dichloromethane (which is added in subsequent steps) can dissolve plastic materials. Therefore, always use glass bottles, pipettes, and syringes while working with dichloromethane.
Fill the water bath with distilled water to the upper level of 2.5 mL of solvent in a glass tube immersed in the water bath. Heat the water bath to 85 °C.
Add 0.5 mL of 15 M KOH to samples to initialize alkaline hydrolysis.
Vortex for 1 min, followed by centrifugation at 3,500× g for 5 min at RT.
Then, immerse glass tubes in a preheated linear shaking water bath at 85 °C for 1 h, under constant shaking at 40 rpm.
Note: This strong alkaline treatment releases DolP from DolP-linked monosaccharides, dolichyl diphosphate (DolPP)-linked mono- and oligosaccharides, and DolPP (Keller et al., 1985; Lee Adair and Kennedy Keller, 1985; Fernandez et al., 2001; Schenk et al., 2001).
Cool samples to RT. Vortex for 1 min, followed by centrifugation at ~3,500× g for 5 min at RT.
Increase the level of water in the water bath with distilled water to the upper level of 7.5 mL of solvent in a glass tube immersed in the water bath. Heat the water bath to 40 °C.
To the cooled samples, carefully add 1 mL of methanol and 4 mL of dichloromethane to induce phase partitioning.
Then, immerse glass tubes in a preheated linear shaking water bath at 40 °C for 1 h, under constant shaking at 40 rpm.
Cool the samples at RT; then, vortex and centrifuge at 3,500× g for 5 min at RT.
Following centrifugation, remove the upper aqueous phase using a Pasteur pipette and wash the lower organic phase four times with wash solvent (see Recipes).
Carefully transfer the bottom layer to a new glass tube using a Pasteur pipette.
Dry samples under a gentle stream of nitrogen at RT in a Turbovap evaporator.
Methylation of DolPs
Note: Inhaling TMSD may cause lung injury or central nervous system depression. TMSD must be handled with caution and following proper safety standards. Before starting the reaction, ensure that the hood is working properly. When working with TMSD, personal safety equipment such as lab aprons, latex gloves, goggles, and respirators should be worn at all times. Screw cap glass tubes immediately after the addition of the reagents.
QC standard: methylate DolP standard and IS mixture along with samples. In a screw-top glass tube, take 20 pmol of DolP standard and 20 pmol of IS and dry under a gentle stream of nitrogen.
Add 200 μL of dichloromethane:methanol mixture (6.5:5.2, v:v, see Recipes) to dried samples.
Vortex for 1 min followed by centrifugation at 3,500× g for 5 min at RT.
To these tubes, carefully add 10 μL of TMSD using a gas-tight Hamilton syringe.
Vortex for 1 min, followed by centrifugation at 3,500× g for 5 min at RT.
Incubate for 40 min at RT.
Note: The chemical reaction of TMSD (yellow) with the acetic acid produces (colorless) methyl acetate. After adding TMSD to the lipid extracts, the solution should have a faint yellow color. If the yellow color associated with TMSD disappears in the solution, add TMSD in 5 μL increments until a yellow color appears. It is recommended to dry the lipid extracts completely before methylation to avoid neutralization of TMSD.
When the reaction is complete, neutralize excess TMSD by slowly adding 10 μL of acetic acid.
Note: The color of the solution should change from yellow to colorless after the addition of acetic acid. Add acetic acid in 5 μL increments until the yellow color disappears.
Dry samples under a gentle stream of nitrogen at RT in a Turbovap evaporator.
For LC–HRMS analysis, reconstitute dried samples in 120 μL of methanol. Vortex for 1 min and transfer lipid films to micro inserts placed in 1.5 mL Eppendorf tubes.
Centrifuge the samples in Eppendorf tubes in a microcentrifuge at 1,500× g for 5 min at 4 °C. Transfer micro inserts to LC vials for analysis.
LC system preparation
Prepare Solvents A and B according to Recipes.
Mix each solvent thoroughly by shaking and degas both solvents in the water bath sonicator with loose caps for at least 10 min.
Put up both solvents on the LC system and purge all lines for 5 min.
Set the flow rate to 0.1 mL/min with both solvents in the ratio 60:40 (solvent A:B).
Maintain the column at 55 °C.
Equilibrate the column for at least 15 min after the column pressure is stable.
Set the LC gradient as follows: 40% B at 0 min, 40%–50% B (0–3 min), 50%–54% B (3–9 min), 54%–70% B (9–9.1 min), 70%–90% B (9.1–17 min), 90% B (17–27.5 min), 90%–40% B (27.5–27.6 min), and 40% B (27.6–30 min).
MS system preparation
Note: Perform mass calibration for the mass spectrometer with XCalibur software.
Set the tune MS parameters to the ESI positive mode, as shown in Table 2.
Table 2. MS parameters used for Q-Exactive Plus high-resolution mass spectrometer
MS parameter value/unit MS parameter value/unit
Spray voltage 3.5 kV Scan range 960 to 1,600 m/z
Sheath gas 30 L/min Spectrum data type Profile
Auxiliary gas 10 L/min Isolation window 4.0 m/z
Spare gas 1 unit Collision energy NCE 50%
S-Lens 50 eV Maximum IT 100 ms
Resolution
AGC Target
17,500 at m/z 200
2 × 105 for MS1
Capillary temperature 250 °C
70,500 at m/z 200
3 × 106 for MS2
Inject 100 μL of each sample in the following order:
Solvent blank: inject solvent as blank at least three times for gradient stabilization.
Then inject QC sample to check the performance of methylation and the LC-MS system.
Randomize the sequence of samples.
Data analysis
Evaluation of raw data
Raw MS1 and MS2 spectral data were processed, analyzed, and visualized using the XCalibur Qualbrowser as shown in Figure 3.
Figure 3. Default layout for the dolichyl phosphates (DolP) C95 example data file using XCalibur Qualbrowser
Qualbrowser in XCalibur was used for MS data visualization. The XICs for [M+NH4]+ ions of commercially available DolP standards and IS were created by extracting theoretical m/z values (shown in Table 1) with a 10 ppm accuracy window.
Chromatographic retentions for DolPs and IS were obtained from the most intense peak in XIC traces.
Further, structural information of DolP species and IS was derived from MS1, and MS2 obtained at their retention time. A representative XIC, MS1, and MS2 analysis of DolP C95 from DolP standard mixture and HeLa cell extracts is shown in Figure 4.
Figure 4. Identification of dolichyl phosphates (DolP) C95 in DolP standard mixture (left panel) and in HeLa cell extracts (right panel) based on A) extracted ion chromatograms (XIC), B) MS spectrum, and C) MS2 spectrum of [M+NH4]+ ions of DolP C95. For more MS spectral characterization and protocols, please refer to Kale et al. (2023).
All methylated DolP species and IS PolP C60 exhibited a characteristic fragment of 127.0155, which is used as a qualifier ion (assigned to the dimethylphosphate group), independent of dolichol or isoprenyl chain length.
Endogenous DolP species were identified using matching MS1, retention time, and MS2 spectrum to standards.
Quantitative analysis of DolP in samples
MS data quantification was established in Quanbrowser by creating a processing method using XCalibur Quanbrowser (Thermo Fisher Scientific XCalibur 3.1 Quanbrowser User Guide, 2014)
Get the peak area from the XICs of DolPs and its internal standards based on its accurate m/z value with an accuracy of 10 ppm. The measured peak areas of the methylated DolPs and internal standards must be corrected by multiplying the isotope correction factors. This correction will take into account the differences in the relative 13C natural isotope abundance of different DolP species.
Theoretical isotopic distributions were obtained using the online EnviPat R package Tool (Loos et al., 2015). Finally, the concentration of DolPs was estimated by comparing the peak areas of internal standards with known concentrations to the peak areas of DolPs.
Validation of protocol
The method mentioned in the present protocol has been thoroughly validated according to ICH guidelines [see supplementary information mentioned in Kale et al. (2023)]. The following method validation parameters were tested: lower limit of quantification (LOD) and the linearity range, intra- and inter-day accuracy and precision, extraction recovery, and post-preparative autosampler stability. As described in Kale et al. (2023), the analytical procedure provided acceptable values of validation parameters tested, and the experimental protocol was robust and reproducible.
Acknowledgments
This protocol was adapted from our previous work (Kale et al., 2023). All experiments described in the protocol were conducted at BZH, Heidelberg, and part of Writing – Review & Editing was performed at ISAS, Dortmund. This project was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) – Project-ID 289991887 – Forschungsgruppe FOR 2509, project 02, – Project-ID 112927078 – Transregio TRR83, project 01, and – Project-ID 278001972 – TRR 186, project Z04. The authors gratefully acknowledge the data storage service SDS@hd supported by the Ministry of Science, Research, and the Arts Baden-Württemberg (MWK) and the DFG through grant INST 35/1314-1 FUGG and INST 35/1503-1 FUGG.
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The authors used this protocol in Science Immunology Jul 2023
Abstract
Cancer cells evade the immune system by downregulating antigen presentation. Although immune checkpoint inhibitors (ICI) and adoptive T-cell therapies revolutionized cancer treatment, their efficacy relies on the intrinsic immunogenicity of tumor cells and antigen presentation by dendritic cells. Here, we describe a protocol to directly reprogram murine and human cancer cells into tumor-antigen-presenting cells (tumor-APCs), using the type 1 conventional dendritic cell (cDC1) transcription factors PU.1, IRF8, and BATF3 delivered by a lentiviral vector. Tumor-APCs acquire a cDC1 cell-like phenotype, transcriptional and epigenetic programs, and function within nine days (Zimmermannova et al., 2023). Tumor-APCs express the hematopoietic marker CD45 and acquire the antigen presentation complexes MHC class I and II as well as co-stimulatory molecules required for antigen presentation to T cells, but do not express high levels of negative immune checkpoint regulators. Enriched tumor-APCs present antigens to Naïve CD8+ and CD4+ T cells, are targeted by activated cytotoxic T lymphocytes, and elicit anti-tumor responses in vivo. The tumor-APC reprogramming protocol described here provides a simple and robust method to revert tumor evasion mechanisms by increasing antigen presentation in cancer cells. This platform has the potential to prime antigen-specific T-cell expansion, which can be leveraged for developing new cancer vaccines, neoantigen discovery, and expansion of tumor-infiltrating lymphocytes.
Key features
• This protocol describes the generation of antigen-presenting cells from cancer cells by direct reprogramming using lineage-instructive transcription factors of conventional dendritic cells type I.
• Verification of reprogramming efficiency by flow cytometry and functional assessment of tumor-APCs by antigen presentation assays.
Keywords: Cell reprogramming Tumor-APCs Conventional type 1 dendritic cells Magnetic-activated cell sorting Antigen presentation Immunotherapy Immunogenicity Cancer vaccine
Background
Immune evasion, an important hallmark of cancer, is characterized by the exclusion of effector immune cells and immunosuppression, the inherent heterogeneity of cancer cells and reduced antigen presentation, which all contribute to tumor immunogenicity (Sharma et al., 2017; Jhunjhunwala et al., 2021). Immune checkpoint inhibition (ICI) revolutionized cancer treatment by enhancing the patient’s immune system, resulting in long-term responses in melanoma and other types of cancer. Nevertheless, some cancers, such as prostate cancer and glioblastoma, are more resistant to immunotherapeutic approaches. Additionally, among indications eligible for immunotherapy, responses vary greatly across patients (Sharma et al., 2017). Antigen presentation and tumor’s intrinsic immunogenicity play crucial roles in promoting efficient CD8+ and CD4+ T-cell responses during ICI treatment (Manguso et al., 2017; Patel et al., 2017; Oh et al., 2020; Blomberg et al., 2023). Immune evasion mechanisms contribute to tumor progression by reducing immunogenicity through upregulation of immune checkpoints (Hashimoto et al., 2018), tumor antigen editing, and downregulation of antigen presentation pathways (Jhunjhunwala et al., 2021). These mechanisms arise as a consequence of transcriptional and epigenetic alterations in antigen processing and presentation pathways (Guo et al., 2021) and deregulation of interferon (IFN) signaling (Kalbasi and Ribas, 2020). Despite attempts at increasing tumor immunogenicity by manipulating the IFN-γ pathway (Griffin et al., 2021; Guo et al., 2021), successful strategies to engineer tumor immunogenicity are still lacking. For the past decade, conventional dendritic cells type I (cDC1) have been established as key players in anti-tumor immunity. cDC1 cells are specialized in cross-presenting tumor antigens to CD8+ T cells, which is fundamental for successful anti-tumor immunity (Poulin et al., 2012; Barry et al., 2018; Kvedaraite and Ginhoux, 2022). Additionally, cDC1 cells recruit and orchestrate immune effectors through the secretion of pro-inflammatory chemokines and cytokines (Spranger et al., 2017). Importantly, the presence of cDC1 cells within the tumor microenvironment correlates positively with immunotherapy’s success and better survival in humans (Spranger et al., 2017; Hubert et al., 2020; Mayoux et al., 2020).
Recently, a cDC1 cells–based vaccine promoted tumor rejection and abscopal tumor control in murine models, independently of host’s cDC1 cell compartment (Ferris et al., 2022). Current methods to generate cDC1 cells rely on the culture of peripheral blood mononuclear cells (PBMCs), bone marrow (BM) progenitors, or monocytes with cytokine cocktails that promote cDC1 cell differentiation (Poulin et al., 2010; Balan and Dalod, 2016). Efforts have also been placed to differentiate leukemic blasts from acute myeloid leukemia into leukemia-derived dendritic cells with cytokine cocktails (Amberger and Schmetzer, 2020). However, ex vivo production of blood-derived dendritic cells is costly, has limited yields, and the differentiation approach is inefficient and limited to blood cancers (Amberger and Schmetzer, 2020; Makino et al., 2022). On the other hand, direct reprogramming approaches have the advantage to generate the desired cell fate from any cell type, have great potential for in vivo application, and are more suited for clinical applications as transdifferentiation processes often bypass progenitor cell states (Wang et al., 2021; Zimmermannova et al., 2021). Our group has defined the minimum network of transcription factors to induce cDC1 cell’s identity through overexpression of PU.1, IRF8, and BATF3 in fibroblasts (Rosa et al., 2018, 2020 and 2022). While this approach could be potentially used to generate induced cDC1 cells, we have hypothesized that the same combination of transcription factors could restore immunogenicity in cancer cells. Cancer cells retain cellular plasticity that allows the modification of cancer cell fate with direct cellular reprogramming (Hochedlinger et al., 2004; Suva et al., 2013; Ishay-Ronen et al., 2019; Zimmermannova et al., 2021), and we have recently demonstrated that ectopic expression of PU.1, IRF8, and BATF3 induces a cDC1 cell–like state in cancer cells (Zimmermannova et al., 2023). Reprogramming of mouse and human cancer cells to antigen-presenting cells occurs by imposing a cDC1 cells’ transcriptional and epigenetic program. Tumor-antigen-presenting cells (tumor-APCs) acquire a cDC1 cell–like immunophenotype, respond to toll-like receptor (TLR) stimuli to secrete pro-inflammatory cytokines, as well as engulf, process, and cross-present antigens as early as day 3 of reprogramming. Tumor-APCs also show endogenous antigens more promptly, becoming targets for T cell–mediated killing, and upon intratumorally injection reduce tumor growth and increase survival in mice (Zimmermannova et al., 2023).
Here, we describe a protocol to generate tumor-APCs within nine days and follow the process at the phenotypic and transcriptional levels. We also describe a magnetic-activated cell sorting (MACS) protocol to purify a high number of tumor-APCs that can be used for in vitro and in vivo assays. While fluorescence-activated cell sorting (FACS) is used to purify rare populations with higher purity, MACS-based purification recovers more viable cells in 4–6-fold less time than FACS (Sutermaster and Darling, 2019; Pan and Wan, 2020). Additionally, MACS requires less expensive instrumentation than FACS, and is easily scalable without adding significantly more processing time. MACS purification of tumor-APCs recovers an average of 2 × 106 of tumor-APCs per 1 × 106 of original cells seeded at day -1 (Figure 1). We also illustrate the functionality of MACS-purified tumor-APCs by performing antigen presentation assays to Naïve CD4+ T cells. As tumor-APCs can be generated from different cancer types, including mouse and human cell lines and primary tumor cells (Zimmermannova et al., 2023), this technology supports a vast array of in vitro approaches that require large numbers of tumor-APCs, e.g., in vitro tumor antigen-specific T-cell expansion or neoantigen identification.
Figure 1. Protocol overview to reprogram mouse and human cancer cells to antigen-presenting dendritic cell-like cells. Tumor cells from murine or human origin are plated one day before transduction with lentiviral particles encoding PU.1, IRF8, and BATF3 in a polycistronic cassette under the control of the spleen focus-forming virus promoter (SFFV). An empty vector with a multiple cloning site (MCS) is used as a control. Both constructs encode eGFP after an internal ribosome entry site (IRES) to trace transduced cells by flow cytometry and fluorescence microscopy. At day 3, confluent plates are split and evaluated for transduction efficiency by flow cytometry. Cells are maintained until day 9 at 60%–80% confluency. Reprogramming efficiency is followed by flow cytometry at days 5, 8, and 9. Tumor-antigen-presenting cells (tumor-APCs) can be purified by fluorescent-activated cell sorting (FACS) for applications that require high purity, including RNA-sequencing (RNA-seq). For in vitro and in vivo functional assessments that require large numbers of reprogrammed cells, tumor-APCs can be enriched by magnetic-activated cell sorting (MACS) through positive selection. To assess enhanced immunogenicity in vitro, tumor-APCs can be co-cultured with activated CD8+ T cell (killing assays). These evaluate whether reprogrammed cancer cells can be targeted by cytotoxic lymphocytes. To evaluate antigen presentation capacity, tumor-APCs are co-cultured with antigen-specific Naïve CD8+ or CD4+ T cells, followed by assessment of T-cell activation and proliferation. In this protocol, we describe antigen presentation to CD4+ T cells to validate tumor-APC function. Moreover, tumor-APCs can be injected intratumorally to evaluate anti-tumor efficacy using tumor growth delay and survival as read-outs.
Materials and reagents
Biological materials
B16F10 cells (B16) (ATCC, catalog number: CRL-6475)
888-mel cells (88MEL) (gifted by Göran B. Jönsson from Lund University)
B6.Cg-Tg (TcraTcrb)425Cbn/J (OT-II mouse) (The Jackson Laboratory, catalog number: 004194) as a source for spleens
C57BL/6j (The Jackson Laboratory, catalog number: 000664) as a source for spleens and bone marrow
Human leucocytes concentrate (Skåne University Hospital, Lund) as a source for PBMCs
Reagents
Plasmid pMD2.G (Addgene, catalog number: 12259)
Plasmid psPAX2 (Addgene, catalog number: 12260)
Plasmid SFFV-mPIB-eGFP containing a tricistronic cassette encoding mouse PU.1, IRF8, and BATF3 (mPIB) (Cell Reprogramming in Hematopoiesis and Immunity Lab, Division of Molecular Medicine and Gene Therapy, Lund University, Sweden)
Plasmid SFFV-hPIB-eGFP containing a tricistronic cassette encoding human PU.1, IRF8, and BATF3 (hPIB) (Cell Reprogramming in Hematopoiesis and Immunity Lab, Division of Molecular Medicine and Gene Therapy, Lund University, Sweden)
Plasmid SFFV-MCS-eGFP empty vector with a multiple cloning site and expressing eGFP (Cell Reprogramming in Hematopoiesis and Immunity Lab, Division of Molecular Medicine and Gene Therapy, Lund University, Sweden)
Dulbecco’s modified Eagle’s medium (DMEM) with high glucose, L-glutamine, and sodium pyruvate (Cytiva, Hyclone, catalog number: SH30243.01)
RPMI 1640 with L-glutamine (Cytiva, Hyclone, catalog number: SH30027.01)
Phosphate buffered saline (PBS) without calcium and magnesium (Cytiva, Hyclone, catalog number: SH30256.01)
Fetal bovine serum (FBS) (Cytiva, Hyclone, catalog number: SV30150.03)
Lenti-XTM qRT-PCR Titration kit (Takara, catalog number: 631235)
Rat serum (GeneTex, catalog number: GTX73226)
Mouse serum (Merck, catalog number: M5905)
GlutaMAXTM Supplement (Thermo Fisher Scientific, Gibco, catalog number: 35050061)
Penicillin-streptomycin solution (Cytiva, Hyclone, catalog number: SV30010)
Sodium pyruvate solution (Thermo Fisher Scientific, Gibco, catalog number: 11360070)
2-Mercaptoethanol (Thermo Fisher Scientific, Gibco, catalog number: 31350010)
Hexadimethrine bromide (Merck, catalog number: H9268-5G)
TrypLE Express (Thermo Fisher Scientific, Gibco, catalog number: 12605010)
Anti-Biotin microbeads (Miltenyi Biotec, catalog number: 130-090-485)
Dead cell removal kit (Miltenyi Biotec, catalog number: 130-090-101)
BD Pharm Lyse lysing buffer (BD Biosciences, catalog number: 555899)
Ammonium chloride solution (Stem Cell Technologies, catalog number: 07800)
Pan-DC Enrichment kit, human (Miltenyi Biotec, catalog number: 130-100-777)
Naïve CD4+ T cell isolation kit, mouse (Miltenyi Biotec, catalog number: 130-104-453)
CellTrace Violet Proliferation kit for flow cytometry (Thermo Fisher Scientific, Invitrogen, catalog number: C34557)
Polyinosinic:polycytidylic acid [poly(I:C)] HMW (InvivoGen, catalog number: tlr-pic-5)
Ovalbumin (OVA) 323–339 (InvivoGen, catalog number: vac-isq)
4′,6′-diamidino-2-phenylindole (DAPI) (Thermo Fisher Scientific, Invitrogen, catalog number: D1306)
Fixable Viability Dye eFluor eFluor520 (Thermo Fisher Scientific, Invitrogen, catalog number: 65-0867-14)
Anti-mouse CD45 Monoclonal Antibody (clone: 30-F11), Biotin (Thermo Fisher Scientific, eBioscience, catalog number: 13-0451-82)
Anti-mouse CD45 Monoclonal Antibody (clone:30-F11), Allophycocyanin-Cyanine7 (Thermo Fisher Scientific, eBioscience, catalog number: 25-0451-82)
Anti-mouse MHC class II (I-A/I-E) Monoclonal Antibody (clone: M5/114.15.2), Biotin (Thermo Fisher Scientific, eBioscience, catalog number: 13-5321-82)
Anti-mouse MHC class II (I-A/I-E) Monoclonal Antibody (clone: M5/114.15.2), PE-Cyanine7 (Thermo Fisher Scientific, eBioscience, catalog number: 25-5321-82)
Anti-mouse CD274 (PD-L1) Monoclonal Antibody (clone: M1H5), PE, (Thermo Fischer Scientific, eBioscience, catalog number: 12-5982-82)
Anti-mouse CD275 (ICOSL) Monoclonal Antibody (clone: HK5.3), PE (Thermo Fisher Scientific, eBioscience, catalog number: 12-5985-82)
Anti-mouse CD366 (Tim-3) Monoclonal Antibody (clone: B8.2.C12), Allophycocyanin (BioLegend, catalog number: 134008)
Anti-mouse CD3 Monoclonal Antibody (clone: 17A2), FITC (BioLegend, catalog number: 100204)
Anti-mouse/human CD45RA (B220) Monoclonal Antibody (clone: RA3-6B2), FITC (Thermo Fisher Scientific, eBioscience, catalog number: 11-0452-85)
Anti-mouse Ly-6G/Ly-6C (Gr-1) Monoclonal Antibody (clone: RB6-8C5), FITC (BioLegend, catalog number: 108406)
Anti-mouse NK1.1 Monoclonal Antibody (clone: PK136), FITC (Thermo Fisher Scientific, eBioscience, catalog number: 11-5941-82)
Anti-mouse CD8a Monoclonal Antibody (clone: 53-6.7), Allophycocyanin-Cyanine7 (BioLegend, catalog number: 100722)
Anti-mouse CD11c Monoclonal Antibody (clone: N418), Brilliant Violet 650 (BioLegend, catalog number: 117339)
Anti-mouse CD4 Monoclonal Antibody (clone: RM4-5), PE/Cyanine7 (Thermo Fisher Scientific, eBioscience, catalog number: 25-0042-82)
Anti-mouse TCR b chain Monoclonal Antibody (clone: H57-597), Allophycocyanin (BioLegend, catalog number: 109212)
Anti-mouse/human CD44 Monoclonal Antibody (clone: IM7), Allophycocyanin-Cyanine7 (BioLegend, catalog number: 103028)
Anti-human HLA-DR Monoclonal Antibody (clone: L243), PE-Cyanine7 (BioLegend, catalog number: 307616)
Anti-human HLA-DR Monoclonal Antibody (clone: L243), Brilliant Violet 711 (BioLegend, catalog number: 307644)
Anti-human CD45 Monoclonal Antibody (clone: H130), Allophycocyanin-Cyanine7 (BioLegend, catalog number: 304014)
Anti-human CD11c Monoclonal Antibody (clone: B-ly6), Violet 450 (BD Biosciences, catalog number: 560369)
Anti-human CD1c Monoclonal Antibody (clone: L161), Allophycocyanin-Cyanine7 (BioLegend, catalog number: 331520)
Anti-human CD141 (thrombomodulin) Monoclonal Antibody (clone: M80), PE-Cyanine7 (BioLegend, catalog number: 344110)
Anti-human CD3 Monoclonal Antibody (clone: UCHT1), FITC (BioLegend, catalog number: 300452)
Anti-human CD19 Monoclonal Antibody (clone: H1B19), FITC (BioLegend, catalog number: 302206)
Anti-human CD56 (NCAM) Monoclonal Antibody (clone: 5.1H11), FITC (BioLegend, catalog number: 362546)
Anti-human VISTA Monoclonal Antibody (clone: B7H5DS8), Allophycocyanin (Thermo Fisher Scientific, eBioscience, catalog number: 17-1088-42)
Anti-human CD274 (PD-L1) Monoclonal Antibody (clone: M1H1), PE (Thermo Fisher Scientific, eBioscience, catalog number: 12-5983-42)
Anti-human CD366 (TIM3) Monoclonal Antibody (clone: F38-2E2), Allophycocyanin (Thermo Fisher Scientific, eBioscience, catalog number: 17-3109-42)
Lentiviral vectors for expression of PU.1, IRF8, and BATF3 individually in inducible vectors (Addgene, catalog numbers: 139839, 139838, and 139837)
Solutions
OVA 323–339 1 mg/mL, prepared according to manufacturer’s protocol
CellTrace Violet (CTV) solution, prepared according to manufacturer’s protocol
DMEM complete media (see Recipes)
RPMI complete media for 88MEL culture (see Recipes)
RPMI complete media for bone-marrow dendritic cells (BM-DCs) (see Recipes)
MACS buffer (see Recipes)
Polybrene 8 mg/mL (see Recipes)
Poly(I:C) 1 mg/mL, prepared according to manufacturer’s protocol (see Recipes)
Recipes
DMEM complete media
Reagent Final concentration Quantity
DMEM high glucose, with glutamine and sodium pyruvate 440 mL
Heat-inactivated FBS 10% (v/v) 50 mL
GlutaMAX 1% (v/v) 5 mL
Penicillin-Streptomycin solution 1% (v/v) 5 mL
Total 500 mL
RPMI complete media for 88MEL culture
Reagent Final concentration Quantity
RPMI 1640 440 mL
Heat-inactivated FBS 10% (v/v) 50 mL
GlutaMAX 1% (v/v) 5 mL
Penicillin-Streptomycin 1% (v/v) 5 mL
Total 500 mL
RPMI complete media for BM-DCs culture
Reagent Final concentration Quantity
RPMI 1640 435 mL
Heat-inactivated FBS 10% (v/v) 50 mL
GlutaMAX 1% (v/v) 5 mL
Penicillin-Streptomycin 1% (v/v) 5 mL
Sodium Pyruvate 1% (v/v) 5 mL
2-Mercaptoethanol 0.05 mM 0.5 mL
Total 500.5 mL
MACS buffer
Reagent Final concentration Quantity
PBS without calcium and magnesium 480 mL
Heat-inactivated FBS 2% (v/v) 10 mL
Penicillin-Streptomycin solution 2% (v/v) 10 mL
Total 500 mL
Polybrene solution
Reagent Final concentration Quantity
Hexadimethrine bromide 8 mg/mL 5 g
Milli-Q H2O 625 mL
Total 625 mL
Poly(I:C) 1 mg/mL solution
Reagent Final concentration Quantity
Poly(I:C) 1 mg/mL 10 mg
Endotoxin free water (provided by the manufacturer) 10 mL
Total 10 mL
Laboratory supplies
Serological pipette, sterile, non-pyrogenic/endotoxin free 2–50 mL (Sarstedt, catalog numbers: 86.1253.001, 86.1256.001, 86.1252.001, 86.1254.001, 86.1685.001)
Pasteur pipette, sterile (Sarstedt, catalog number: 86.1171.001)
1 L vacuum filter system, 0.22 mm PE (Corning, catalog number: 431098)
100 mm tissue culture plates (Corning, catalog number: 430167)
Petri dish, 92 mm × 16 mm (Sarstedt, catalog number: 82.1473)
150 mm tissue culture plates (Corning, catalog number: 430599)
6-well tissue culture plates (Corning, catalog number: 353046)
6-well non-tissue-culture-treated plates (Corning, catalog number: 351146)
96-well tissue culture U-shaped bottom plates (Corning, catalog number: 353077)
96-well non-tissue-culture-treated U-shaped bottom plates (Corning, catalog number: 351177)
15 mL centrifuge tubes (Sarstedt, catalog number: 62.547.205)
50 mL centrifuge tubes (Sarstedt, catalog number: 62.547)
MidiMACSTM LS columns (Miltenyi Biotec, catalog number: 130-042-501)
Hemocytometer chamber for cell counting (Corning, CytoSmart, catalog number: 480202)
Microcentrifuge tubes (Sarstedt, catalog number: 72.690.001)
Cell strainer 45 mm nylon mesh (Corning, catalog number: 11587522)
Syringe 5 mL (Avantor, VWR, catalog number: 613-2042)
Round bottom polystyrene test tube 5 mL (Corning, Falcon, catalog number: 352008)
Tweezers curved ends (Avantor, VWR, catalog number: 232-0106)
Forceps (Avantor, VWR, catalog number: 232-0085)
Scissors (Avantor, VWR, catalog number: 233-0225)
Equipment
Scanlaf Mars class 2 laminar flow hood (LaboGene, model: Mars 1200 mm)
Forma Steri-cycle i160 CO2 incubator (Thermo Scientific, model: i160)
Integra Pipetboy acu 2 (Integra-Biosciences, model: acu 2)
BD FACSymphony A1 or LSRFortessa flow cytometer analyzers (16-color, violet/blue/red/yellow-green) (BD Biosciences, model: A1 Cell Analyzer; LSRFortessa Analyzer)
MACS MultiStand (Miltenyi Biotec, catalog number: 130-042-303)
MidiMACSTM separator (Miltenyi Biotec, catalog number: 130-042-301)
4–16K refrigerated centrifuge (Sigma, catalog number: 10474)
Automatic cell counter (Corning, CytoSmart, catalog number: 6749)
Milli-Q Type 1 Ultrapure Water systems (Merck, model: IQ7000)
Thermo Scientific Finnpipette F2 pipettes (Thermo Fisher Scientific, catalog number: 4642010)
Thermo Scientific Finnpipette F2 Multichannel pipettes (Thermo Fisher Scientific, catalog number: 4662030)
Vortex (IKA, model: MS3 basic)
Water bath (Grant Instruments, model: JB series)
Software and datasets
FACSDIVA (v 9.0) (Access date, 03 May 2023)
FlowJo (v 10.8.1) (Access date, 03 May 2023)
GraphPad Prism (v 9) (Access date, 05 May 2023)
CytoSmart (Cell Count Algorithm V3) (Access date, 01 May 2023)
Mouse Tumor-APC RNA-seq data GSE184527 (Access date, 07 July 2023)
Mouse cDC1 cells RNA-seq data GSE103618 (Access date, 07 December 2018)
Procedure
Preparing cancer cell cultures for reprogramming
Note: This protocol is performed in sterile conditions under a class 2 laminar flow hood, using sterile pipettes, materials, and reagents.
Procedures A–C from this protocol were performed using B16 and 88MEL murine and human melanoma cell lines but can be extended to other types of cancer cell lines and primary samples, including adherent and non-adherent cells (Zimmermannova et al., 2023). Cancer cell lines and primary samples can be bought from ATCC, Riken, AMSBio, BioIVT, or VitroBiopharma, among other commercial sources. Alternatively, cancer biopsies can be isolated, processed, and expanded for reprogramming experiments.
Thaw and culture cancer cells in the recommended culture media for the cell type under standard conditions (37 °C, 5% CO2). It is recommended that cells are passaged at least once before plating for reprogramming. For better reproducibility when using the same cell line, ensure that the passage is similar between reprogramming experiments.
Before reprogramming, ensure the cells are dividing normally and display usual morphology by using an inverted light microscope regularly.
Using a hemocytometer, count cells and expand them as follows:
For adherent cells, plate 1 × 106 live cells per 100 mm tissue culture plate.
Note: This step can be adjusted according to the purpose of the experiment. To assess reprogramming efficiency or immunophenotyping during reprogramming, 6-well plates are recommended. For functional assays or in vivo studies, it is recommended to use 100 mm or 150 mm plates. Adjust cell numbers according to the surface area.
For non-adherent cells, skip to step B2.
Allow cells to adhere by incubating at 37 °C, 5% CO2 overnight (approximately 16 h), and proceed with reprogramming (Procedure B).
Tumor-APC reprogramming
Before starting this protocol, prepare lentiviruses as previously described (Rosa et al., 2020 and 2022). The lentiviral vectors for expression of PU.1, IRF8, and BATF3 individually in inducible vectors can be found in Addgene and can be subcloned into any backbone as individual factors or in a polycistronic cassette by adding self-cleaving 2A sequences between sequences.
Titer viral concentrates by qRT-PCR to determine their concentration in viral particles/mL using, for example, Lenti-X qRT-PCR titration kit (see Reagents). This will require viral RNA isolation from supernatant before qRT-PCR reaction to quantify viral RNA content. Calculating viral particles per milliliter of viral supernatant does not provide functional titration, but it ensures reproducibility even when using virus lacking a fluorescent protein. Alternatively, functional titration by FACS can be performed based on eGFP expression and calculating transducing units per milliliter.
The reprogramming process can be done in multiple cell culture media. Use the appropriate media for culturing the cancer cell type of choice throughout reprogramming. B16 were cultured in complete DMEM and 88MEL in complete RPMI (see Recipes 1 and 2).
Pre-warm appropriate volume of cell culture media for 5 min at 37 °C.
Cancer cell transduction is performed as described below for adherent (2a) and non-adherent cells (2b):
To transduce adherent cells seeded in a 100 mm tissue culture plate (Procedure A, Figure 1):
i. Resuspend the volume necessary containing of 4 × 104 viral particles per cell in 10 mL.
Note: The range of 4 × 104 viral particles per cell is optimal for B16 and 88MEL cancer cell lines and can be used as a starting point for similar cell types. However, it is recommended to evaluate the optimal multiplicity of infection (MOI) for efficient reprogramming without affecting cell viability for each individual cell cancer cell type. This is performed by testing increasing viral titers or volumes with known number of particles (determined by qRT-PCR) and assessing transduction efficiency at day 3 and reprogramming efficiency at day 9 (Figure 1). Optimal transduction efficiency with eGFP lentivirus is approximately 70%–80% at day 3. If the lentivirus lacks a fluorescent reporter, reprogramming efficiency can be assessed at day 9 by flow cytometric analysis using CD45 and MHC-II as read-out.
ii. Add 10 mL of polybrene solution to 10 mL of transduction media prepared in the previous step (final concentration: 8 mg/mL, see Recipe 5).
iii. Scale volume of transduction media, lentivirus particles, and polybrene according to the number of plates and plated cells.
iv. Incubate at 37 °C, 5% CO2 for 24 h in transduction media. Proceed to step B3.
To transduce non-adherent cells:
i. Resuspend cells in an adequate volume of media to a final density of 1 × 106 cells/mL.
ii. Add optimized MOI of lentiviral particles for the cell type of choice followed by polybrene (final concentration: 8 mg/mL) to the cell suspension.
iii. Dispense 2 mL of cell suspension per well in a non-tissue-culture-treated 6-well plate.
Note: Non-adherent cells may require higher viral load for efficient reprogramming. It is recommended that viruses are titrated before this step (see note in previous step).
iv. Perform spin infection by centrifugation at 800× g for 1 h at room temperature.
v. After centrifugation, resuspend the cells and incubate at 37 °C, 5% CO2 for 24 h in transduction media.
Change media of transduced cells.
For adherent cultures, aspirate supernatant and carefully add media on top of the cells.
Note: Cautiously dispense media at low speed by adjusting settings in a pipette aid to avoid displacing transduced cells, especially semi-adherent cells.
For non-adherent cells, transfer cell suspension to a 50 mL centrifuge tube. Centrifuge cell suspension for 5 min at 350× g at room temperature, before resuspending cells in adequate media volume for a final density of 1 × 106 cells/mL.
At day 3, observe cells under an inverted light microscope.
Note: Reprogramming-associated cell death is expected after transduction and during the first days of reprogramming (approximately 10%), which can be variable among different cell lines.
If transduced cancer cells in a 100 mm tissue culture plate reach 80% confluency at day 3, aspirate media, wash with room-temperature PBS, and add 1.5 mL of TrypLE Express. Incubate for the minimum amount of time needed until all cells have lifted (minimum 3 min). Resuspend cells in 7 mL of their usual fresh culture media and centrifuge at 350× g for 5 min at room temperature. Aspirate media and resuspend cells in 60 mL to passage 1:6 (i.e., to plate in 6 × 100 mm tissue culture plates). For large experiments, an 80% confluent 100 mm tissue culture plate can be divided into 3 × 150 mm tissue culture plates.
Note: Cell dilution can be adjusted according to cancer cell line of choice and experimental setup. Fast replicating cells can be diluted in 1:6 or 1:8 ratios. Slow dividing cells should be diluted in 1:3 or 1:4 ratios to avoid over dilution.
If transduced cancer cells are not 80% confluent and flow cytometric analysis will not be performed, just change media as in step B3.
For non-adherent cells, transfer cell suspension to a 50 mL centrifuge tube and count cells. Centrifuge at 350× g for 5 min at room temperature followed by resuspension in adequate volume of fresh media for a final density of 1 × 106 cells/mL.
Transduction efficiency can be measured at day 3 of reprogramming by flow cytometric analysis of eGFP expression. Perform step B4a and collect 0.5–1 mL out of 7 mL of cell suspension for flow cytometric analysis in step B4e. Divide the remaining cells evenly on 100 mm or 150 mm plates according to dilution ratio.
Flow cytometric analysis at day 3: Divide the cell suspension collected in step B4d evenly in a sufficient number of round-bottom polystyrene tubes to include single color (SC) controls and fluorescence minus one (FMO) control to assess expression of other markers along with eGFP (i.e., CD45, MHC-II, and MHC-I) and establish the optimal fluorochrome panel and compensation settings. See Table 1 for an example of number of tubes and their contents for flow cytometric analysis of eGFP expression combined with surface staining for CD45, MHC-II, and MHC-I. Note that, by using a fluorescent tag protein like eGFP, all single-color staining will include eGFP as well and FMO eGFP cannot be done unless panel establishment is performed using vectors without eGFP. If measurements are done on the same machine, compensation settings are only required to be performed once during reprogramming, generally at day 5 post-transduction when marker expression is more intense (see step B6).
Table 1. Single-color and FMO tubes required for flow cytometry panel establishment
Sample Tube Fluorochromes
Untransduced cancer cells Unstained Not applicable
Viability dye Viability dye (i.e., DAPI)
eGFP-transduced cancer cells SC eGFP Only eGFP
eGFP all stained All antibodies and viability dye
Tumor-APCs SC PE-Cyanine7 + eGFP Only PE-Cyanine7 antibody (i.e., anti-MHC-II)
SC Allophycocyanin-Cyanine7 + eGFP Only Allophycocyanin -Cyanine7 antibody (i.e., anti-CD45)
SC-Allophycocyanin + eGFP Only Allophycocyanin antibody (i.e., anti-MHC-I)
FMO PE-Cyanine7 All antibodies except PE-Cyanine7
FMO Allophycocyanin All antibodies except Allophycocyanin
FMO Allophycocyanin-Cyanine7 All antibodies except Allophycocyanin -Cyanine7
FMO viability dye All antibodies except viability dye
Tumor-APCs all stained All antibodies and viability dye
Change media every 2–3 days, according to the color of the media and cell confluency until the endpoint of reprogramming.
Note: For cancer cells with high doubling rates, replace media every 24 h. Do not let cells reach >80% confluency, as high confluency levels result in low pH, altered metabolism, and mechanical stress that may affect the reprogramming process. Whenever 80% confluency is reached, split cells according to step 4a. For non-adherent cells, maintain cell density between 1 × 106 and 2 × 106 cells/mL during the reprogramming process.
The emergence of tumor-APCs can be assessed at days 5, 8, and 9 post-transduction by flow cytometric analysis.
For adherent cancer cells:
i. Aspirate media, wash with 3 mL of room-temperature PBS, and dissociate cells with 1.5 mL TrypLE Express per plate followed by incubation at 37 °C, 5% CO2 for 3 min (or until all cells have lifted from the plate).
ii. Harvest cells by resuspending them with 3 mL per 100 mm plate of cold MACS buffer (see Recipe 4, keep it at 4 °C) and transferring the resultant cell suspension to a conical 50 mL centrifuge tube. Centrifuge at 350× g for 5 min at 4 °C.
iii. Discard supernatant and resuspend a pellet equivalent to a 100 mm tissue culture plate in 3 mL of MACS buffer.
For non-adherent cells:
i. Resuspend the cells using serological pipettes and transfer culture suspension to a 50 mL centrifuge tube.
ii. Count cell number using a hemocytometer before centrifuging at 400× g for 5 min at 4 °C.
iii. Resuspend non-adherent cell cultures in an adequate volume of cold MACS buffer for a final concentration of 1 × 106 cells/mL.
Cells are now ready for downstream analysis by flow cytometry, FACS-based purification for fluorescence microscopy, RNA-sequencing (RNA-seq), assay for transposase-accessible chromatin with sequencing (ATAC-seq), or MACS-enrichment.
Divide cell suspension evenly into 3–11 round-bottom tubes to assess stained and unstained samples. Include SC and FMO controls if it is the first time doing flow cytometric analysis (see step B4e and Table 1 for examples). CD45, MHC-II, and MHC-I can be used for FACS purification.
Reprogramming of cancer cells can be evaluated by RNA-seq data by quantifying mRNA expression of endogenous PU.1, IRF8, and BATF3, as well as antigen presentation genes and TLR genes (see Data analysis).
Purifying tumor-APCs for functional assays by MACS-enrichment
The MACS protocol here described was optimized for adherent murine melanoma cells using Miltenyi Biotec LS columns and MidiMACS separators. Further optimizations for different cancer cell types might be needed.
For experiments requiring large numbers of reprogrammed cancer cells, it is recommended to perform MACS-enrichment ensuring feasibility of the experiment and viability of the reprogrammed population. For experiments requiring tumor-APCs with >85% purity, it is recommended to perform MACS-enrichment 1–2 days before FACS. This will enrich total reprogrammed cells 10-fold and yield an average of 2 × 106 reprogrammed cells at day 9 per 1 × 106 cancer cells seeded at day -1 (Figure 1), reducing time for FACS purification.
At day 9 of reprogramming, detach cells as described in step B6a.
Resuspend and dissociate cells gently by pipetting up and down 10 times with 3 mL of cold MACS buffer per pellet from 100 mm tissue culture plate. Count cells using a hemocytometer and calculate total cell number.
Note: Adjust starting cell numbers depending on reprogramming efficiency and final application. For cancer cells that have low reprogramming efficiency (<10% of double-positive cells for CD45+ MHC-II+), we recommend starting with a minimum of 1 × 107 cancer cells.
To perform quality control by flow cytometric analysis after the procedure is completed, it is recommended that 0.1–0.3 mL of cell suspension (pre-MACS) is stored on ice in a round-bottom tube.
Note: If a high level of cell debris and cell death is noticed, consider using a dead cell or debris removal kit in advance, as dying cells can bind unspecifically to antibodies and result in poor enrichment yields (see Table 6, Troubleshooting).
Centrifuge cell suspension from step 2 at 350× g for 5 min at 4 °C.
Note: The volumes included in the next steps are for 107 cells. Adjust volumes according to the total cell number obtained in step C2.
Aspirate supernatant and resuspend 107 cells in 0.1 mL of MACS buffer.
Pipette 2 µL of rat serum per 107 total cells of rat serum into the cell suspension.
(Critical) Place cells on ice and incubate for 15 min.
Note: It is crucial that incubation steps are always performed on ice to avoid capping of antibodies on the cell surface and non-specific cell labeling.
Pipette 0.125 µg of biotin-labeled anti-CD45 and 0.06 µg of biotin-labeled anti-MHC-II per 107 total cells. Resuspend cells by lightly vortexing the sample.
Note: the volume of antibodies may require further optimization (see Table 6, Troubleshooting). If the cell line of choice does not express a marker abundantly (CD45 or MHC-II), it is possible to perform MACS with one antibody only targeting the most expressed marker.
Place cell suspension on ice and incubate for 5 min. See General note 1 for an alternative to biotin beads.
Wash cells with 2 mL per 107 total cells of cold MACS buffer followed by centrifugation at 350× g for 5 min at 4 °C.
Aspirate supernatant and resuspend cells in 80 µL of cold MACS buffer per 107 total cells.
Add 20 µL of anti-biotin microbeads per 107 total cells and resuspend cell suspension by lightly vortexing the sample.
Place cells on ice and incubate for 15 min.
In the meantime, prepare magnetic MACS MultiStand and columns. Place the MACS MultiStand inside the laminar flow hood and attach magnetic MidiMACS separators.
Insert LS MACS columns into MidiMACS separators.
Equilibrate columns by washing with 3 mL of cold MACS buffer. Discard the liquid and place a 50 mL centrifuge tube under the column.
After incubation, wash cells by adding 2 mL of MACS buffer per 107 total cells followed by centrifugation at 350× g for 5 min at 4 °C.
Aspirate supernatant and resuspend cells in 1 mL per 107 total cells.
(Critical) For optimal enrichment, it is crucial to obtain a single-cell suspension at this step. Pipette up and down to remove cell clumps before introducing magnetic-labeled cells in columns. If necessary, filter cells through a 50 mm nylon mesh.
Note: When working with columns, always wait until the column reservoir is empty before proceeding to the next step.
(Critical) Transfer 0.5 mL of cell suspension onto the column. It is recommended to only apply 0.5–1 mL of filtered magnetic-labeled, single-cell suspension per column to avoid column clogging.
Wash column with 3 mL of MACS buffer. Repeat twice.
In this step, cells in flowthrough are untransduced and unsuccessfully reprogrammed cancer cells. It is recommended that 0.1–0.2 mL of flowthrough [post-MACS negative fraction, (-)] is saved for evaluation by flow cytometry after the protocol is completed. Place remaining cells on ice (see Table 6, Troubleshooting).
Remove column from the separator and place it on top of a 15 mL centrifuge tube. Wait for 1 min.
Pipette 5 mL of MACS buffer into the column reservoir and immediately but carefully plunge the liquid out of the column (see Table 6, Troubleshooting). In this step, the eluted cell suspension contains partial and complete reprogrammed tumor-APCs labeled by CD45 or MHC-II antibodies.
Centrifuge at 350× g for 5 min at 4 °C.
Resuspend cell pellet in 5 mL of media. Count cells using a hemocytometer.
(Critical) Save 0.1–0.2 mL of the cell suspension in step C23 [post-MACS positive fraction, (+)] for flow cytometric analysis.
Prepare enough round-bottom polystyrene tubes for the following samples: pre-MACS, post-MACS(-), post-MACS(+), unstained, viability stain (DAPI or fixable viability dye), and SC and FMO controls. Unstained, viability stain, SC, and FMO controls should be run with pre-MACS cell suspension saved in step 3 of this procedure. For example, for full quality control assessment, pre-MACS, post-MACS(-), and post-MACS(+) should be stained with fluorescently labeled anti-CD45 and anti-MHC-II and viability stain before flow cytometry evaluation (see Data analysis). Optionally, anti-MHC-I can also be used as a marker. See Table 2 for an example of number of tubes and specific contents for each before flow cytometric analysis.
Table 2. Example of number of tubes needed for flow cytometric analysis of MACS efficiency using pre-MACS and post-MACS samples
Sample Tube Fluorochromes
Untransduced cancer cells Unstained Not applicable
Viability dye Viability dye (i.e., DAPI)
Pre-MACS SC staining See Table 1
FMO staining See Table 1
Pre-MACS All fluorochromes (anti-CD45, anti-MHC-II, optional anti-MHC-I) and viability dye
Post-MACS(-) Post-MACS(-) All fluorochromes and viability dye
Post-MACS(+) Post-MACS(+) All fluorochromes and viability dye
eGFP-transduced cancer cells eGFP All fluorochromes and viability dye
After quality control of the enriched tumor-APCs population, post-MACS positive fraction should contain >70% of partial and complete reprogrammed cells. Total number of cells expected after MACS enrichment is 2 × 105–6 × 106 cells per 106 cells seeded.
Resuspended cells are now ready for downstream application in functional antigen presentation assays, killing assays by activated CD8+ T cells, and in vivo intratumoral injections.
Note: Tumor-APCs exit the cell cycle during the reprogramming process and cannot expand as the original cancer cell (Zimmermannova et al., 2023). We did not test whether tumor-APCs can be freeze-thawed before functional assays. We recommend using freshly reprogrammed tumor-APCs for further applications.
Evaluate exogenous antigen presentation to Naïve murine CD4+ T cells
This part of the protocol is performed using reprogrammed murine B16 melanoma cells as tumor-APCs. eGFP-transduced cancer cells and BM-DCs should be included as negative and positive controls, respectively. See General note 2 for BM-DCs preparation and freezing before experimental setup. A minimum of 3.3 × 105 BM-DCs are necessary. See General note 3 for alternative APCs to BM-DCs.
One day before MACS procedure, thaw one vial of BM-DCs (containing 3 × 106 cells) by gently resuspending the frozen cells in 10 mL of pre-warmed complete RPMI for BM-DCs (see Recipe 3).
Transfer BM-DCs cell suspension to a 15 mL conical tube. Centrifuge at 350× g for 5 min at room temperature to remove freezing solution.
Aspirate supernatant and resuspend BM-DCs in 10 mL of new RPMI for BM-DCs. Plate cell suspension in a non-tissue-culture-treated Petri dish. Incubate cells overnight at 37 °C, 5% CO2.
After MACS enrichment of tumor-APCs, count enriched cells and prepare a microcentrifuge tube with 1 × 106 resuspended tumor-APCs.
Note: 1 × 106 tumor-APCs is ideal to account for pipetting errors. The minimum number of tumor-APCs that can be used for this protocol is 3 × 105.
Centrifuge tumor-APCs at 350× g for 5 min at room temperature. Remove supernatant and resuspend tumor-APCs in 0.5 mL of complete RPMI for BM-DCs.
Detach eGFP-transduced cancer cells as described in step B6a. Resuspend cells in 10 mL of warm media and calculate cell concentration using a hemocytometer.
Prepare a microcentrifuge tube with 1 × 106 eGFP-transduced cancer cells and repeat step D5.
Prepare a tissue-culture-treated 96-well U-shaped bottom plate with media only as described below (see Table 3 for an example of the experimental setup):
Table 3. Experimental setup for antigen presentation using tumor-APCs and control eGFP-transduced cells in a tissue-culture-treated 96-well U-shaped bottom plate
Wells A B C D E F G
1 Tumor-APCs
NA Tumor-APCs
OVA 323–339 eGFP-transduced cancer cells
NA eGFP-transduced cancer cells
OVA 323–339
2
3
4
9 Tumor-APCs
NA
Poly(I:C) Tumor-APCs
OVA 323–339
Poly(I:C) eGFP-transduced cancer cells
NA
Poly(I:C) GFP-transduced cancer cells
OVA 323–339
Poly(I:C)
10
11
12
Prepare 1.5 mL of RPMI for BM-DCs, no antigen (NA) added. Pipette 0.1 mL of this media in eight wells.
Prepare 1.5 mL of RPMI for BM-DCs, 10 mg/mL poly(I:C), NA added. Pipette 0.1 mL of this media in eight wells.
Note: Alternatively, or in conjunction with poly(I:C), other TLR agonists can be used.
Prepare 1.5 mL of RPMI for BM-DCs, 10 mg/mL OVA 323–339. Pipette 0.1 mL of this media in eight wells.
Prepare 1.5 mL of RPMI for BM-DCs, 10 mg/mL poly(I:C), and 10 mg/mL OVA 323–339. Pipette 0.1 mL of this media in eight wells.
Note: Other TLR agonists can be used, replacing or in combination with poly(I:C).
(Critical) Resuspend and pipette 5 µL (dispensing 1 × 104 cells per well) of each APC suspension (tumor-APC or eGFP-transduced cancer cells) in the middle of their assigned U-shaped bottom wells (see Table 3) and observe each well under the microscope. It is important that seeded cells cluster together at the center of the well for optimal antigen presentation.
Note: Avoid dispersing cells in the walls of the plate. This will ensure maximum contact between APCs and T cells.
Prepare BM-DCs by transferring the supernatant from step D3 to a conical 15 mL centrifuge tube and resuspend well. Calculate cell density using a hemocytometer.
Transfer 1 × 106 BM-DCs to a microcentrifuge tube. Centrifuge cells at 350× g for 5 min at room temperature. Aspirate supernatant as much as possible and resuspend cells in 0.5 mL of complete RPMI for BM-DCs.
Prepare a non-tissue-culture-treated 96-well U-shaped bottom plate with media only as described below (see Table 4 for an example of the control samples setup):
Prepare 0.7 mL of RPMI for BM-DCs, NA added. Pipette 0.1 mL in wells A1–A4.
Prepare 0.7 mL of RPMI for BM-DCs, 10 mg/mL poly(I:C). Pipette 0.1 mL in wells C1–C4.
Prepare 1.8 mL of RPMI for BM-DCs, 10 mg/mL OVA 323–339. Pipette 0.1 mL of media in wells A9-A12. Add 0.1 mL of media to wells E1–E12, which will be later used as staining controls for flow cytometric analysis.
Prepare 0.7 mL of RPMI for BM-DCs, 10 mg/mL Poly(I:C), and 10 mg/mL OVA 323–339. Pipette 0.1 mL in wells C9–C12.
(Critical) Resuspend BM-DCs from step D11, pipette 5 µL (dispensing 1 × 104 cells per well) in the middle of each U-shaped bottom wells (see Table 4), and observe each well under the microscope. It is important that seeded cells cluster together at the center of the well for optimal antigen presentation.
Note: BM-DCs are used as a positive control for the assay. Alternatively, it is possible to seed up to 6 × 104 cells per well. Adjust final resuspension volume to pipette only 5 µL per well and keep seeded cells in the center of the well.
Table 4. Experimental setup for antigen presentation using BM-DCs in a non-tissue-culture-treated 96-well U-shaped bottom plate
Wells A B C D E
1 BM-DC
NA BM-DC
NA
Poly(I:C) BM-DC
OVA 323–339
2
3
4
5
6
7
8
9 BM-DC
OVA 323–339 BM-DC
OVA 323–339
Poly(I:C)
10
11
12
Incubate all 96-well plates at 37 °C, 5% CO2 overnight (12–16 h). Observe cells the day after. All cells should be gathered at the center of the well.
On the day after seeding APCs for antigen presentation assays, start by preparing sterile forceps, tweezers, a pair of scissors, and 20 mL of cold MACS buffer. This will be used for spleen isolation.
Euthanize OT-II mice according to protocols approved by local ethical committees and the regulations of the animal facility where the procedure is being conducted. For this protocol, mice were sacrificed in a CO2 chamber followed by cervical translocation.
Note: To calculate the number of mice needed, estimate how many Naïve T cells are necessary for the whole assay, considering that the number seeded per well is 1 × 105 cells. The number of isolated Naïve CD4+ T cells is expected to be between 1.5 × 106 and 3 × 106 cells per spleen. For the setup exemplified in this protocol, a total of 7 × 106 Naïve T cells will be needed.
Place mice on a surface where they can lie on the left side facing upwards (see Data analysis).
Spray the torso with 70% ethanol to reduce the chance of contamination.
Cut the skin and peritoneum right below the thorax and slightly off center to the left of the mouse’s body to reveal the spleen (see Data analysis).
Note: If the spleen looks enlarged (splenomegaly) or black, discard the carcass and the spleen.
Pull out and hold the spleen with the help of tweezers with fine tips and cut the connective tissue and fat attached to the spleen.
Note: Removing connective tissue and fat as much as possible at this stage will facilitate Naïve T-cell isolation by avoiding cell clumping.
Remove the spleen and store it in cold MACS buffer prepared in step D15. Repeat for the number of mice sacrificed.
Discard the carcasses according to local animal facility’s procedures and continue the protocol under sterile conditions.
Place a 45 mm cell strainer on top of a 50 mL centrifuge tube.
Place a 45 mm cell trainer inside a Petri dish. Pipette 3 mL of cold MACS buffer to prepare the filter.
Using forceps or a pipette, place one spleen inside the cell strainer. With the help of a 5 mL syringe pestle, crush the spleen against the cell strainer mesh.
Once most of the spleen is crushed, remove filter from plate. With a 10 mL serological pipette, pipette 5 mL of MACS buffer into the plate and wash the plate before transferring the cell suspension through the cell strainer from step D23. Serial filtration will avoid cell clumping with connective tissue and fat in later stages.
Repeat steps D24–D26 for every spleen.
Note: The filter from step D23 and the plate from step D24 can be reused for up to four spleens. Substitute filter in step 24 for every new spleen that needs to be crushed. The syringe pestle can be reused for all spleens. If needed, use several centrifuge tubes to collect the samples.
When splenocytes collection is finished, centrifuge cell suspension at 350× g for 5 min at room temperature.
Meanwhile, prepare red blood cell lysis solution. Pipette 0.1 mL of BD Pharm Lyse solution for every 0.9 mL of sterile room temperature Milli-Q water. Keep this solution in the dark. Use 1 mL of this solution per spleen, i.e., for 15 spleens, prepare 15 mL of BD Pharm Lyse solution as described.
Note: Alternatively, red blood cells can be lysed using ammonium chloride solution.
Aspirate supernatant of splenocytes solution. Resuspend solution in the appropriate volume of red blood cell lysis solution prepared in step D29 (1 mL per spleen) and incubate splenocytes for 8 min at room temperature in the dark.
Stop the reaction by adding 50 mL of cold MACS buffer. If you notice aggregates floating, filter through a 45 mm mesh.
Note: It is important that these aggregates are removed at this step to avoid loss of Naïve T cells in clumps.
Centrifuge filtered cell suspension at 350× g for 5 min at 4 °C.
Aspirate the supernatant and resuspend cells in 10–30 mL of cold MACS buffer. Count splenocytes using a hemocytometer.
Note: If splenocytes were separated into different centrifuge tubes, they can be combined at this point of the procedure. Resuspend all the pellets and combine them in a single cell suspension together in one tube. The cell suspension should look dense and cloudy. Dilute the sample before counting.
Centrifuge cell suspension at 350× g for 5 min at 4 °C. Aspirate supernatant and resuspend the pellet in 40 µL of MACS buffer per 1 × 107 total cells.
Proceed by following manufacturer’s protocol for Naïve CD4+ T-cell isolation (mouse Miltenyi kit). Naïve cells are unlabeled by the kit and are found in the negative fraction.
After isolating Naïve CD4+ T cells, count the number of cells using a hemocytometer.
Note: If the number of Naïve CD4+ T cells is higher than 5 × 106 per spleen, it is recommended to repeat the Naïve CD4+ T-cell isolation kit protocol to avoid contamination with activated CD4+ T cells and other cell lineages.
For each well containing APCs, prepare 1 × 105 T cells labeled with CellTrace Violet (CTV) as follows:
Transfer the volume of cell suspension necessary to contain 5.5 × 106 Naïve CD4+ T cells for 48 assay wells in addition to five more wells for control staining (see step D47 for flow cytometric analysis). Keep the leftover Naïve CD4+ T cells on ice for later use. Centrifuge cell suspension at 350× g for 5 min at 4 °C.
Prepare CTV according to manufacturer’s protocol by resuspending the powder in 20 mL of dimethyl sulfoxide (provided in the kit).
Aspirate supernatant and resuspend cells in 5.5 mL of cold PBS with no additives for a final density of 1 × 106 cells/mL. Pipette 5.5 µL of CTV reagent into the cell suspension (1 µL per 1 × 106 T cells).
Incubate T cells in a water bath at 37 °C for 20 min.
After incubation, add five times the initial staining volume of MACS buffer to block staining and incubate for 5 min in the water bath.
Centrifuge cells at 350× g for 5 min at 4 °C and aspirate supernatant. Resuspend T cells in RPMI for BM-DCs for a final cell density of 1 × 106 cells/mL.
Prepare the different conditions of T cells as follows:
Prepare 1.2 × 106 Naïve T cells in RPMI for BM-DCs, corresponding to 1.2 mL from the cell suspension prepared in step D37f, and dispense 0.1 mL (dispensing 1 × 105 T cells per well) on top of the wells for the following conditions: Tumor-APCs NA, eGFP-transduced cancer cells NA, and BM-DCs NA.
Prepare 1.2 × 106 Naïve T cells in RPMI for BM-DCs, corresponding to 1.2 mL from the cell suspension prepared in step D37f, add 10 mg/mL poly(I:C), and dispense 0.1 mL (dispensing 1 × 105 T cells per well) on top of the wells for the following conditions: Tumor-APCs NA poly(I:C), eGFP-transduced cancer cells NA poly(I:C), and BM-DCs NA poly(I:C).
Prepare 1.9 × 106 Naïve T cells in RPMI for BM-DCs, corresponding to 1.9 mL from the cell suspension prepared in step D37f, add 10 mg/mL OVA 323–339, and dispense 0.1 mL (dispensing 1 × 105 T cells per well) on top of the wells for the following conditions: Tumor-APCs OVA, eGFP-transduced cancer cells OVA, and BM-DCs OVA. Include five wells of the extra BM-DCs OVA condition seeded on the day prior (for example, on top of wells E1–E5).
Prepare 1.2 × 106 Naïve T cells in RPMI for BM-DCs, corresponding to 1.2 mL from the cell suspension prepared in step D37f, add 10 mg/mL OVA 323–339 and 10 mg/mL poly(I:C), and dispense 0.1 mL (dispensing 1 × 105 T cells per well) on top of the wells for the following conditions: Tumor-APCs OVA poly(I:C), eGFP-transduced cancer cells OVA poly(I:C), and BM-DCs OVA poly(I:C).
Note: B16 cancer cells do not express MHC-II before reprogramming (see Data analysis). Therefore, it is not needed to wash away the peptide from supernatant in 96-well plates and it is recommended to keep the peptides until the end of co-culture. However, if the cell line of choice expresses MHC-II before reprogramming, it is recommended to centrifuge the 96-well plates at 350× g for 5 min at room temperature, before plating Naïve T cells on top of the APCs. Remove supernatant by flipping the plate downwards in a fast movement, wash the wells with fresh RPMI media, centrifuge for 5 min at room temperature, and remove media again before proceeding with step D38.
For the leftover extra wells for BM-DCs OVA, prepare 8 × 105 unlabeled Naïve CD4+ T cells from step D37a by resuspending the cells in 0.8 mL of RPMI for BM-DCs and adding 10 mg/mL of OVA 323–339.
Pipette 0.1 mL of unlabeled T-cell resuspension on top of each of the remaining six wells (for example, wells E6–E11), dispensing 1 × 105 T cells per well. These samples will be used for flow cytometric analysis (see step D47).
Incubate plates with co-cultures at 37 °C, 5% CO2 for four full days. Analysis is done at day 5.
Antigen presentation is evaluated by OT-II CD4+ T cells proliferation on a flow cytometer. Prepare antibody staining solution by mixing 0.5 µL of rat serum, 0.5 µL of antibody anti-CD4 PE-Cyanine7, anti-TCRbeta allophycocyanin, and anti-CD44 allophycocyanin-Cyanine7 per 0.1 mL of MACS buffer per sample.
Add to the antibody staining solution fixable viability dye eFluor 520 (FVD520) at a 1:100 dilution (1 µL per 0.1 mL per sample)
Centrifuge 96-well U-shaped bottom plates with T cell tumor-APCs, eGFP-transduced cancer cells, or BM-DCs co-cultures at 350× g for 5 min at 4 °C.
Flip the plate in a fast movement to remove media.
Add 0.1 mL of antibody-viability dye staining solution per well.
Separately, prepare staining controls as follows (see Table 5):
Table 5. Single-color and FMO staining controls for flow cytometric analysis of T-cell activation and expansion
Sample/well Tube Fluorochrome
Unlabeled T cells
(from BM-DC OVA 323–339 wells E6-E11)
Unstained Not applicable
Viability dye Viability dye (FVD)
SC CD4 Only antibody for CD4
SC CD44 Only antibody for CD44
SC TCR Only antibody for TCR
FMO CTV All antibodies and viability dye
CTV-labeled T cells
(from BM-DC OVA 323–339 wells E1-E5)
SC CTV Not applicable
FMO CD4 All antibodies except anti-CD4 and viability dye
FMO CD44 All antibodies except anti-CD44 and viability dye
FMO TCR All antibodies except anti-TCR and viability dye
FMO viability dye All antibodies except viability dye
Single color staining for CD4, CD44, TCR, and FVD and FMO CTV are done using unlabeled T cells from BM-DCs OVA condition.
Single color staining for CTV and FMO CD4, CD44, TCR, and FVD are done using CTV-labeled T cells from BM-DCs OVA condition.
Reserve the supernatant of one well to use as unstained control.
Incubate plates for 30 min at 4 °C, protected from light.
Add 0.1 mL of cold MACS buffer to each well and centrifuge plates at 350× g for 5 min at 4 °C.
Flip the plates to remove staining solution and resuspend the cell pellets in the 96-well plates in 100 µL of cold MACS buffer before going to a flow cytometer.
Flow cytometry analysis can be done directly from the 96-well plate using a high-throughput screening device attached to the flow cytometer. Alternatively, pipette each sample to a polystyrene tube prior to acquisition.
OT-II proliferation is measured by gating within live, single CD4+ TCR+ CD44+ CTVlow population of CD4+ T cells, with T-cell division assessed by loss of CTV staining overtime (see Data analysis).
Note: This protocol can also be used for assessing antigen presentation through MHC-I to Naïve CD8+ T cells (Zimmermannova et al., 2023). Instead of using OVA 323–339, incubate APCs with OVA 257-264 (SIINFEKL) or OVA protein. Wash peptide before co-culture with Naïve T cells, as some cells naturally express MHC-I that leads to unspecific binding. Instead of using OT-II mouse strain, collect spleens from OT-I mouse strain (C57BL/6-Tg(TcraTcrb)1100Mjb/J) as described in steps D16–35, and isolate Naïve CD8+ T cells using appropriate kits. Co-culture with Naïve CD8+ T cells should be shortened to three days instead of four. In this scenario, anti-CD4 is substituted for anti-CD8 in the same color, PE-Cyanine7.
Human tumor-APCs can also be used for antigen presentation assays by pulsing tumor-APCs with peptides derived from cytomegalovirus (CMV) and melanoma-associated antigen recognized by T cells (MART-1), followed by co-culture with CD8+ T cells isolated from CMV+/MART-1+ donors. Flow cytometric analysis is done at day 8 of co-culture and read-out is obtained from tetramer staining against CMV or MART1 and intracellular staining for IFN-γ and TNF-α to assess activation (Zimmermannova et al., 2023). In addition to antigen presentation assays, tumor-APCs can be employed in killing assays, by co-culturing with activated T cells followed by assessing T cell–mediated cytolysis by flow cytometry or by electric impedance with the xCELLigence Real-time Cell Analysis Assay (Zimmermannova et al., 2023). Finally, tumor-APCs can be used for therapeutic intratumoral vaccination protocols. In all applications described here, tumor-APCs derived from cancer cell lines and primary samples can be used.
Data analysis
Mouse and human melanoma cells are amenable to direct cell reprogramming by overexpression of PU.1, IRF8, and BATF3 (Figure 1), imposing a cDC1 cell-like cell fate (Zimmermannova et al., 2023). Tumor-APCs can be harvested at days 3, 5, 8, or 9 of reprogramming to assess reprogramming efficiency by immunophenotyping and transcriptional profiling (Figure 1). Reprogramming efficiency is assessed at reprogramming day 9 by flow cytometry through the emergence of partial (CD45+ or MHC-II+) and completely reprogrammed (CD45+MHC-II+) populations within transduced cells (eGFP+) (Figure 2A–2B), reflecting the transition to the hematopoietic lineage and acquisition of antigen presentation machinery. Additionally, we tested the expression of immune checkpoint inhibitors CD274, ICOSL, TIM3, and VISTA at cell surface of reprogrammed cells (Figure 2C–2D). Overall, we show that reprogrammed cells express these molecules at levels in a similar range to natural cDC1 cells isolated from mouse spleens (live, lineage negative for B220, CD3, NK1.1, CD19, and Gr1, MHC-II+CD11c+CD8a+) and human peripheral blood (live, lineage negative for B220, CD3, CD56, and CD19, HLADR+CD11c+CD141+).
Figure 2. Immunophenotyping human and mouse-derived eGFP-transduced cancer cells by flow cytometry. A. Gating strategy to analyze the reprograming efficiency of mouse-derived eGFP-transduced cancer cells and B. human-derived eGFP-transduced cancer cells at day 9 post induction of PU.1, IRF8, and BATF3 (PIB). Live cells were gated on viability dye negative population (4,6-diamidino-2-phenylindole, DAPI) followed by singlet selection. Expression of CD45+MHC-II+ was quantified by flow cytometry following a gating strategy based on fluorescence minus one (FMO) staining. eGFP-transduced cells were used as controls. C. Mouse and D. human tumor-APCs were evaluated by flow cytometry for the expression of PD-L1, ICOSL, TIM-3, and VISTA at day 9 of reprogramming. The gating strategy was based on FMO for the respective fluorochrome. eGFP-transduced cancer cells and splenic (mouse, live, lineage negative for B220, CD3, NK1.1, CD19, and Gr1, MHC-II+CD11c+CD8a+) or peripheral blood (human, live, lineage negative for B220, CD3, CD56, and CD19, HLADR+CD11c+CD141+) cDC1 cells were used as controls (biological replicates n = 3–11). Mean ± SD is represented.
Population RNA-seq data can also be used to validate the successful acquisition of a tumor-APC transcriptional program in purified CD45+MHC-II+/HLA-DR+ cancer cells (Zimmermannova et al., 2023). For example, we have previously demonstrated that reprogrammed eGFP+ cancer cells upregulate endogenous gene expression PU.1, IRF8, and BATF3 as well as start expressing the APC machinery at the mRNA level. Gene set enrichment analysis using RNA-seq data can also be used to follow tumor-APC maturity. In contrast to eGFP transduced cancer cells, we observed that reprogrammed tumor-APCs derived from mouse melanoma and lung carcinoma cells activate the expression of Tlr1-6 (Figure 3). These findings are consistent with the expression of co-stimulatory molecules following TLR stimuli with Poly(I:C) and lipopolysaccharide (Zimmermannova et al., 2023).
Figure 3. Profiling PIB-eGFP-transduced cancer cells maturity by toll-like receptor (TLR) analysis. Genes encoding murine Tlr1-6 are expressed in mouse melanoma and lung cancer cells (B16, LLC, eGFP+CD45+MHC-II+) on reprogramming day 9 after PIB induction (red) and splenic conventional dendritic cells type 1 (cDC1, MHC-II+CD11c+CD8a+). After 9 days post transduction with PIB, tumor-APCs (eGFP+MHC-II+CD45+), eGFP-transduced cancer cells (eGFP+MHC-II-CD45-), and splenic cDC1 cells (MHC-II+CD11c+CD8a+) were FACS purified. Transcriptome was assessed by bulk RNA sequencing to quantify gene expression (Rosa et al., 2018; Zimmermannova et al., 2023, GSE103618, GSE184527). Gene expression is depicted as whisker-box plots with minimum, maximum, mean, and standard deviation shown. As an alternative, RT-qPCR can be executed to quantify the expression of TLR and assess the tumor-APCs maturation profile.
Tumor-APC function can be assessed by cytokine secretion, endogenous and exogenous antigen presentation capacity, CD8+ T cell–mediated killing assays, and enhanced in vivo anti-tumor activity (Figure 1). However, due to the number of cells required for these applications, enrichment for the reprogrammed population is required. Here, we describe an optimized protocol for enriching reprogrammed murine cancer cells using a MACS-based purification (Figures 4A–4B). Quality control is performed by flow cytometry through viability, total transduced cells (eGFP+), and total percentage of cells expressing either CD45, MHC-II, or both markers (partial and complete reprogrammed cells) (Figures 4C–4D). eGFP+ expression is confirmed in the majority of reprogrammed PIB-eGFP-transduced cancer cells after MACS isolation [post-MACS(+), % eGFP+ cells 73.94 ± 16.81], contrasting with 34.60 ± 23.09% and 32.62 ± 22.00% of successfully transduced cells found in pre-MACS and post-MACS(-) fraction (Figure 4E). More importantly, the enriched positive fraction [post-MACS(+)] contains 72.23 ± 12.74% of partially and complete reprogrammed cells (Figure 4F). This enrichment is supported by low percentages of reprogrammed cells in post-MACS-negative fraction [post-MACS(-), 5.82 ± 4.1%], which represents a 10-fold enrichment from pre-MACS (10.97 ± 7.45%) to the post-MACS(+) fraction (Figure 4F). Additional phenotypic analysis by evaluation of MHC-I expression illustrates that tumor-APCs express more MHC-I molecules according to higher mean intensity fluorescence (MFI) than eGFP-transduced cancer cells (Figure 4G, H). This optimized protocol for purification of tumor-APCs can yield approximately 2 × 106 total cells per 1 × 106 cells seeded cancer cells at day -1.
Figure 4. Enrichment of PIB-eGFP-transduced cancer cells by a magnetic activated cell sorting (MACS)-based method and antigen presentation assessment. A. Schematic overview of a MACS-protocol to enrich mouse melanoma–derived PIB-eGFP-transduced cancer cells based on the expression of CD45 and MHC-II. Reprogramming cultures were labeled with anti-CD45 and anti-MHC-II, followed by the introduction of magnetic beads targeting the previous antibodies. The cell suspension is first filtered in a nylon mesh before passing through a column under a strong magnetic field where positively labeled cells (PIB-eGFP-transduced cancer cells) will attach, letting the negative population flow through in the post-MACS-negative fraction (-). The column is then removed from the magnetic MACS MultiStand and placed in a centrifuge tube before elution and plunging of positively labeled PIB-eGFP-transduced cancer cells in the post-MACS-positive fraction (+). Quality control was assessed by flow cytometry after collecting samples pre-MACS, post-MACS(-), and post-MACS(+). The gating strategy follows viability dye negative population selection before duplet exclusion and evaluation of the total reprogrammed cell population that includes either CD45+, MHC-II+, or double positive cells (partial and complete reprogrammed cells). eGFP+ population is also evaluated as a measure of transduction efficiency. B. Photographs depicting tumor-APC enrichment in the magnetic columns (top) followed by elution outside the magnetic stand (bottom). C. Representative flow cytometry plots for expression of eGFP and D. representative flow cytometry plots of a successful MACS procedure. Pre-MACS and post-MACS(-) samples were included to demonstrate the enrichment efficiency. E. Quantification of eGFP+ cells representing total % of transduced cells in pre-MACS samples and post-MACS(-) (post -) and (+) fractions (post +) (biological replicates n = 4–9). F. Quantification of partial and complete reprogrammed cells in pre-MACS samples and post-MACS(-) and (+) fractions. eGFP-transduced cancer cells were included as controls (biological replicates n = 4–9). G. Representative flow cytometry histograms for MHC-I expression across eGFP-transduced cancer cells and tumor-APCs pre- and post-MACS fractions. H. Median intensity fluorescence (MFI) for MHC-I expressed by cancer cells and tumor-APCs pre- and post-MACS fractions (biological replicates n = 3). *** p-value < 0.001; **** p-value < 0.0001
To provide evidence for the function of MACS-enriched tumor-APCs, we demonstrated antigen presentation to Naïve CD4+ T cells (Figure 5A). We have previously shown that reprogrammed cancer cells present endogenous and exogenous antigens through MHC-I context to Naïve CD8+ T cells (Zimmermannova et al., 2023). Here, we assess their antigen presentation capacity through MHC-II pathway. For this, MACS-enriched reprogrammed cancer cells were fed OVA peptides 323–339, which are specifically presented in MHC-II to CD4+ T cells (Figure 5A). Splenic Naïve CD4+ T cells are co-cultured with tumor-APCs for four days before flow cytometric analysis (Figures 5A–5B). Melanoma-derived tumor-APCs in co-culture with Naïve CD4+ T cells induced the expression of CD44 and T-cell expansion at levels similarly elicited by BM-DCs (Figure 5C) (PIB %CD44+CTVlow: 88.48 ± 5.47, BM-DCs %CD44+CTVlow: 85.35 ± 3.86).
Figure 5. Evaluation of tumor-APCs exogenous antigen presentation to Naïve CD4+ T cells. A. Schematic representation of an antigen presentation assay using tumor-APCs as exogenous antigen presenters to CD4+ T cells. Enriched tumor-APCs are cultured with ovalbumin (OVA) peptides (323–339), which bind specifically to MHC-II molecules. After OVA 323–339 pulse, tumor-APCs are then co-cultured with Naïve CD4+ T cells isolated from OT-II mouse model. Naïve CD4+ T cells were labeled with CellTrace Violet (CTV) before the start of co-culture. T cell expansion is measured after four days of co-culture by flow cytometric analysis. B. Photographs depicting spleen isolation and collection from OT-II mouse strain. Splenocytes are isolated and enriched for Naïve CD4+ T cells. C. T cell activation was evaluated by CD44 expression (CD44+) and CTV dilution (CTVlow) (biological replicates n = 4). Mean ± SD is shown.
In summary, this protocol describes the reprogramming of murine and human cancer cells into tumor-APCs by ectopic expression of PU.1, IRF8, and BATF3. Tumor-APCs undergo stepwise acquisition of a mature APC transcriptional and immunophenotypic profile. The expression of CD45 and MHC-II allows MACS purification for in vitro and in vivo functional assays. Tumor-APCs are endowed with enhanced immunogenicity and can present exogenous antigens to Naïve CD8+ (Zimmermannova et al., 2023) and CD4+ T cells, as shown in this protocol. Moreover, tumor-APCs constitute a novel strategy to enhance anti-tumor immunity through cell reprogramming towards cDC1 cell fate by directly inducing antigen presentation in cancer cells, surpassing a central immune evasion mechanism. Finally, direct reprogramming of cancer cells can serve as a platform for alternative methods for neoantigen discovery and in vitro expansion of tumor infiltrating lymphocytes.
Validation of protocol
All experiments were repeated independently at least once. Data obtained by flow cytometry for quality control after MACS-enrichment (Figure 4E, 4F and 4H) were subjected to a normality test before the application of repeated measures one way ANOVA (matched pairs), with Geisser-Greenhouse correction enabled, followed by Tukey’s multiple comparison test in GraphPad Prism.
This protocol was validated in Zimmermannova et al. (2023)
General notes and troubleshooting
General notes
General note 1: Labeling for MACS procedure can be done using fluorescently labeled antibodies, i.e., PE-CD45. To recover positively labeled cells, use appropriately labeled microbeads against the fluorochrome, i.e., anti-PE microbeads. It is not recommended to perform this version of MACS if the cells will be used for in vitro assays with subsequent flow cytometry readouts, as the persistence of fluorescent antibodies in culture can interfere with the analysis.
General note 2: BM-DCs are used as reference samples for antigen presentation assays. Prepare BM-DCs according to Mayer et al. (2014) in advance. Freeze day-16 BM-DCs with a cell density of 2 × 106–3 × 106 cells per vial, to account for cell loss during freezing-thawing cycles, and thaw one vial the day before experimental setup.
General note 3: As an alternative to using BM-DCs as reference for antigen presentation assays, it is possible to use fresh isolated APCs, such as cDC1, cDC2 cells, or macrophages from C57BL/6j spleens.
Troubleshooting
Table 6. Troubleshooting
Step Problem Possible reason Solution
B4 and 5 High levels of cell death Initial virus load too high Confirm the minimum amount copy number necessary to transduce and reprogram efficiently by testing different volumes of the same virus concentrate for the cell type of choice. Use a eGFP reporter and evaluate the transduction efficiency at days 3 and 9 by flow cytometry. qPCR titration is recommended to calculate the number of particles used per seeded cell.
B4 Low transduction levels Initial virus load too low Functionally titrate the virus and calculate the minimum amount needed for efficient transduction and reprogramming without compromising cell viability.
B6 Low reprogramming efficiency The cell type of choice can be resistant If the cell type of choice is not amenable to reprogramming despite high transduction levels, check for literature descriptions of epigenetic alterations silencing MHC-II, MHC-I, and CD45 gene loci as well as hard mutations in the genome in known cDC1 cells–specific genes.
C3 High levels of cell debris and cell death Acidic media during reprogramming
Initial viral load too high
Cell type of choice is fragile
Remove cell debris and dead cells by using a dead cell removal kit to avoid unspecific binding to antibodies and poor enrichment yields.
C19 Cell clumping and column clogs Cells are large and sticky. Use EDTA as a chelating agent in the MACS buffer throughout the entirety of the protocol.
C28 Poor enrichment yield–post-MACS(+) fraction has more than 20% contaminating untransduced cells
Cell volume/concentration loaded in the column was too high for the cell type of choice
Insufficient column washing
Avoid loading too many cells onto the column.
If cells are large, decrease the volume loaded onto one column and divide the volume of cell suspension by more columns.
Thoroughly wash the columns with the indicated volume and ensure that the liquid passes through completely before loading more volume.
Ensure all liquid has passed through before removing the column from the magnetic stand. Repeat MACS procedure.
C28 The post-MACS(+) fraction has a high number of dead cells
A plunger was inserted too fast in the column
The plunger was pushed up and down in the column
When plunging the positively labeled cells from the column, do it slowly.
Avoid pushing the plunger up and down in the column once the cells are inside as it can cause suction and damage cells.
Wash column twice with 5 mL of MACS buffer, only plunging slowly on the last wash outside of the magnetic stand.
C28 The post-MACS(-) fraction has high levels of reprogrammed cells Insufficient or improper staining before microbead addition
Ensure that the quantity of antibodies used is adequate. This might need optimization.
Repeat enrichment on the post-MACS(-) fraction.
Acknowledgments
This project has received funding from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program (Grant agreement No. 866448). This project was also funded by Cancerfonden (20 0939 PjF), the Swedish Research Council (2020-00615), NovoNordisk Fonden (0056527), the Mats Paulsson Foundation, SweLife and Medtech4Health (2020-04744), Eurostars-2 Joint Program (2021-03371), and FCT (2022.02338.PTDC), and Plano de Recuperação e Resiliência de Portugal pelo fundo NextGenerationEU (C644865576-00000005). The Knut and Alice Wallenberg foundation, the Medical Faculty at Lund University, and Region Skåne are acknowledged for financial support. A.G.F is supported by an FCT scholarship (SFRH/BD/133233/2017).
This protocol was adapted from the publication of Zimmermannova et al. (2023).
Competing interests
C.-F.P has an equity interest and serves in a management position at Asgard Therapeutics, AB, which develops cancer immunotherapies based on reprogramming technologies. C.-F. P., A.G.F, O.Z. and E.A are inventors on US patent 11,345,891, patent applications WO 2018/185709 and WO 2022/243448 held by Asgard Therapeutics, AB, which covers the cell reprogramming protocol described here.
Ethical considerations
Animal care and procedures for strains OT-II and C57BL/6j were performed in accordance with Swedish guidelines and regulations after approval from the local ethical committee (ethical permit: 5.8.18-11845/2019). Human leucocyte concentrates from healthy donors were provided by the Clinical Immunology and Transfusion Medicine at Skåne University Hospital (ethical permit: 2022:11). Donor consent and anonymity are ensured by the hospital.
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Immunology > Immune cell function > Antigen-specific response
Cell Biology > Cell engineering > Partial reprogramming
Cancer Biology > Tumor immunology
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Human Schwann Cells in vitro II. Passaging, Purification, Banking, and Labeling of Established Cultures
PM Paula V. Monje
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4882 Views: 588
Reviewed by: Vivien J. Coulson-ThomasDjamel Eddine ChafaiRomina Vuono
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Original Research Article:
The authors used this protocol in Scientific Reports Oct 2020
Abstract
This manuscript describes step-by-step procedures to establish and manage fresh and cryopreserved cultures of nerve-derived human Schwann cells (hSCs) at the desired scale. Adaptable protocols are provided to propagate hSC cultures through serial passaging and perform routine manipulations such as enzymatic dissociation, purification, cryogenic preservation, live-cell labeling, and gene delivery. Expanded hSCs cultures are metabolically active, proliferative, and phenotypically stable for at least three consecutive passages. Cell yields are expected to be variable as determined by the rate of growth of individual batches and the rounds of subculture. The purity, however, can be maintained high at >95% hSC regardless of passage. The cells obtained in this manner are suitable for various applications, including small drug screens, in vitro modeling of neurodevelopmental processes, and cell transplantation. One caveat of this protocol is that continued expansion of same-batch hSC populations is eventually restricted due to senescence-linked growth arrest.
Keywords: Ensheathing glia Peripheral nerve Cell culture Mitogenic factors Laminin Serial passaging Fibroblast contamination Immunopanning Magnetic-activated cell sorting Cryopreservation Scalability Proliferation Senescence
Background
Human Schwann cells (hSCs) from nerve tissues are fairly amenable cells for in vitro culture. Whereas preparing primary cells requires labor-intensive procedural steps, standard, general practices can be applied to manage the cultures successfully once the hSCs are established. hSCs are expandable under adherent conditions. Substantial scaling up of same-prep cultures can be achieved by supplementing the culture medium with specific mitogenic factors and plating the cells on matrix-coated dishes, preferably with laminin [reviewed in Monje (2020a and 2020b)].
The steady expansion of individual hSC populations is feasible up to the second or third passage because the rate of hSC proliferation diminishes thereafter in most preparations. However, a single harvest of adult nerve-derived fascicles may yield cultures able to expand at a >103 amplification rate and total yields may surpass 100 million cells within two or three rounds of subculture. Established hSC cultures maintain key characteristics of lineage-committed SCs, as evidenced by the expression of SC-specific markers such as NGFR, S100B, Nestin, and Sox10 (Peng et al., 2020). Aberrant growth or spontaneous hSC transformation has not been observed (Emery et al., 1999). Indeed, hSCs from expanded batches are deemed safe for transplantation partly due to their phenotypic stability and resistance to transform (Bastidas et al., 2017). In managing established hSC cultures, researchers should be mindful that the cells achieve an irreversible state of growth arrest or senescence by or around the fifth passage, or the equivalent of 20 population doublings, after their initial isolation from an adult nerve source (Levi et al., 1995; Levi, 1996).
This protocol describes our optimized approaches to propagate hSC cultures and perform various routine manipulations in vitro. A thorough description of materials, methods and procedures is presented here to obtain scalable hSC cultures from both fresh isolates and cryogenic stocks. The banking of hSCs is described in detail due to its multiple benefits for deferred experimentation, long-term storage, and transfer of cell cultures. Several purification and cell labeling methods are introduced along with recommendations on choosing certain methods for specific applications. The following sections describe step-by-step procedures to facilitate replication in other labs. Figure 1 presents a generic representation of suggested interventions at any given step of the culture workflow. This paper does not include comprehensive data sets. The microscopy image data in the figures and videos are provided for qualitative purposes only. Our publications contain more information about laboratory-scale experimentation using established hSC cultures (Monje et al., 2018; Peng et al., 2020). Investigators can use our papers to gather additional details on experimental design, use of positive and negative controls, and analysis of results from diverse assays. The hSC cultures prepared as described herein are reliable models for the development of cell-based platforms, including reconstituted co-culture systems, and studies of xenotransplantation. Lastly, these methodologies are intended for basic research, since they differ substantially from clinical protocols (Khan et al., 2021), and may be effective only when using hSC cultures prepared from normal nerve fascicles or ganglia. We have not used hSC cultures from unconventional sources.
Figure 1. Generation and management of established human Schwann cells (hSC) cultures: passaging, purification, transfer, and in vitro modifications.The diagram summarizes the suggested steps described in each protocol. Primary, passage-zero (P0) hSC cultures (drop-plated dish, left panel) can be expanded at a 1:10 ratio (controlled passaging, upper panel) or an unfixed ratio (routine passaging) until enough cells are obtained or the cultures become senescent, as indicated. Purification, banking, and gene delivery (or labeling) can be attempted at any level of passage but preferably no later than passage-2 or -3, which is the optimal time for experimentation using cells that are both proliferative and pure. The indicated cell yields represent accumulated cell numbers in each passage as predicted by the expansion rate of a typical hSC culture containing 1 million cells at P0.
Materials and reagents
All materials, reagents, and solutions should be sterile and cell culture grade. If possible, consistently use materials and reagents of the same brand and maintain a record of lot numbers in case signs of cell toxicity are observed. The list below is not intended to be fully comprehensive or limit the use of products from certain manufacturers. The product information is provided for reference only. Follow the manufacturer’s recommendations and use best practices in cell culture to avoid microbial contamination and ensure the reproducibility of the results.
Supplies and consumables
Disposable serological pipettes (5, 10, and 25 mL), polystyrene, plugged, sterile, and individually wrapped (VWR, catalog numbers: 76201-710, 75816-100, and 75816-090)
Polystyrene Pasteur (transfer) pipettes, sterile and individually wrapped (VWR, Argos Technology, catalog number: 10122-560)
Petri dish, bacteriological grade, 10 cm, not tissue culture treated (Corning, catalog number: 351029)
Centrifuge tubes, 15 and 50 mL, polypropylene, conical-bottom (Corning, catalog numbers: 430791 and 430290)
Cell culture dishes (35, 60, and 100 mm), polystyrene, tissue culture treated (Corning, catalog numbers: 353001, 353002, and 353003)
Multi-well plates (6, 12, and 24-well), tissue culture treated, flat bottom (Corning, catalog numbers: 3506 and 3524)
Cell culture flasks (T25 and T75), canted neck with plug seal caps (Corning, catalog numbers: 430168 and 430720U)
Cryogenic vials, 2 mL, internal thread (Corning, catalog number: 430489)
Polycarbonate freezing container with a tube holder, Nalgene “Mr. Frosty” Freezing Container (VWR, catalog number: 55710-200)
Cell lifter, individually wrapped, polyethylene, 2 cm length blade, 18 cm long (VWR, catalog number: T-2443-4)
Ice pans (VWR, catalog number: 89233) containing wet ice and dry ice
Media, supplements, and other cell culture products
Distilled water, cell culture grade (Fisher Scientific, Gibco, catalog number: 15-230-147)
Hank’s balanced salt solution (HBSS), formulated without calcium or magnesium and containing phenol red, pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 14170-112)
Leibovitz’s L15 Medium (L15) (Thermo Fisher Scientific, Gibco, catalog number: 11415064)
Dulbecco’s Modified Eagle’s Medium (DMEM), with high glucose and phenol red, pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 11965092). (Optional) Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12) (Thermo Fisher Scientific, Gibco, catalog number: 11320033)
Gentamycin 1,000× provided as 50 mg/mL stock solution (Thermo Fisher Scientific, Gibco, catalog number: 15750-060). (Optional) Penicillin-Streptomycin, 10,000 U/mL stock solution (Thermo Fisher Scientific, Gibco, catalog number: 15140-148)
De-complemented fetal bovine serum (FBS) (HyClone, catalog number: SV 30014.03). Stored in aliquots at -80 °C; used in all applications requiring serum, including cryopreservation of cell stocks
GlutaMAX supplement (100×) consisting of 200 mM L-alanyl-L-glutamine dipeptide in 0.85% NaCl (Thermo Fisher Scientific, Gibco, catalog number: 35050061)
Heregulin-β1, 177-244 amino acid peptide (Preprotech, catalog number: G-100-03); instructions on the preparation, storage, and use of heregulin-β1 stock solution (25 µM) can be found in our protocol Andersen and Monje (2018). The stock solution is best preserved when kept in aliquots at -80 °C and used only for media preparation, avoiding freezing/thawing cycles. Heregulin-β1 is the primary mitogenic factor for hSCs; it is referred to as “heregulin” in the text and figures
Forskolin powder (Sigma-Aldrich, catalog number: F68861); instructions on the preparation, storage, and use of forskolin stock solution (15 mM) can be found in our protocol Andersen and Monje (2018). Keep the forskolin stock solution in the -80 °C freezer for long-term preservation of the cAMP-inducing activity. Forskolin is a synergistic enhancer of heregulin-dependent hSC proliferation
0.5% Trypsin/EDTA (TE) solution (10×), without phenol red (Thermo Fisher Scientific, Gibco, catalog number: 15400054). (Optional) TrypLETM select enzyme, 1× (Thermo Fisher Scientific, Gibco, catalog number: 12563011)
Laminin stock, consisting of a sterile 1 mg/mL laminin solution from Engelbreth-Holm-Swarm murine sarcoma basement membrane (Sigma-Aldrich, catalog number: L2020), stored at -80 °C in aliquots for single use. Prepare laminin-coated dishes as described in Andersen and Monje (2018). Always use freshly coated laminin dishes for seeding hSCs. Nerve fibroblasts can be plated in regular cell culture–treated dishes without coating
Poly-L-lysine (PLL) powder (Sigma, catalog number: P-263). Prepare a PLL stock solution and use it for coating dishes as described in Andersen and Monje (2018). Store air-dried PLL-coated dishes at 4 °C for up to one month
Dulbecco’s phosphate-buffered saline (DPBS), pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 14190)
Dimethyl sulfoxide (DMSO) (Invitrogen, catalog number: D12345). Used as a component of the freezing medium and solvent of lyophilized fluorophores
(Optional) RecoveryTM cell culture freezing medium, ready to use, stored in aliquots at -80 °C (Thermo Fisher Scientific, Gibco, catalog number: 12648010)
Isopropyl alcohol (Fisher Chemical, catalog number: A416P-4L)
Tris base, powder (Thermo Fisher Scientific, Invitrogen, catalog number: 15504020)
Hydrochloric acid (HCl) solution, 1 N, bioreagent suitable for cell culture (Sigma, catalog number: 7647-01-0); used to balance pH in buffers and cell culture medium
Antibodies, dyes, and assorted commercially available products
Anti-NGFR mouse IgG monoclonal antibody, produced from the HB-8737 hybridoma cell line (American Type Culture Collection, ATCC). See Ravelo et al. (2018) for technical details on our culture protocols for this and other hybridoma cell lines
Anti-O4 mouse IgM monoclonal antibody, produced from the O4 hybridoma cell line (Sommer and Schachner, 1981), kindly provided by Dr. Melitta Schachner. (Optional) Use purified O4 antibodies from a commercial source (Novus Biologicals, catalog number: NL637)
Anti-mouse immunoglobulins, goat polyclonal antibody (anti-IgG, IgA, and IgM), affinity purified, liquid, unconjugated (ICN/Cappel, catalog number: 55486)
CellTrackerTM Green CMFDA (5-chloromethylfluorescein diacetate) powder (Invitrogen, Molecular Probes, catalog number: C7025)
(Optional) Pluronic F-127 (Molecular Probes, catalog number: P-6866)
Hoeschst-34580 (Molecular probes, catalog number: H21486) prepared in water at 1 mg/mL
Basic NucleofectorTM kit for primary mammalian glial cells (Lonza, catalog number: VPI-1006)
GFP plasmids. The pmaxGFPTM expression vector (0.5 µg/µL in 10 mM Tris pH 8.0) can be used as provided in the Basic NucleofectorTM kit or expanded in house
Lentiviral particles, aliquoted and stored at -80 °C for up to six months. For a reference, we have used lentiviral expression vectors for the fluorescent reporters EGFP and mCherry (Monje et al., 2018) acquired as ultra-purified, ready-to-use, packaged lentiviruses from the Viral Vector Core Facility, The Miami Project to Cure Paralysis, Miami, FL. Follow the manufacturer’s recommendations on the safe storage and use of viral stocks, including the MOI determination for each virus lot. Multiple freeze/thaw cycles of the viral particles are not recommended
(Optional) Polybrene infection/transfection reagent, 10 mg/mL stock solution (Santa Cruz Biotechnology, catalog number: sc-134220)
(Optional) Eukaryotic antibiotics for the selection of virally-infected cells. Puromycin dihydrochloride (Santa Cruz Biotechnology, catalog number: sc-108071); Blasticidin S HCl solution (Santa Cruz Biotechnology, catalog number: sc-495389); Hygromycin B solution (Santa Cruz Biotechnology, catalog number: sc-29067)
Solutions
Low proliferation medium (LP) (see Recipes)
High proliferation medium (HP) (see Recipes)
TE dissociation solution (see Recipes)
Freezing medium (see Recipes)
Immunoglobulins solution (see Recipes)
Equipment
Biosafety cabinet, BL2 level (Thermo Fisher Scientific, model: 1300 Series A2)
CO2 cell incubator set up at 37 °C and 8%–9% CO2 (Thermo Fisher Scientific, model: Forma Steri-Cycle)
Benchtop centrifuge (Beckman Coulter, model: Allegra X-I2R)
Inverted phase contrast microscope with an attached digital camera (VWR, model: V5MP)
Cell counter for automated counting of cells in suspension (Bio-Rad, TC20). (Optional) Hemocytometer for manual cell counting
4 °C refrigerator and -80 °C laboratory freezer (Thermo Fisher Scientific, model: RLE Series)
Liquid nitrogen storage tank (Thermo Fisher Scientific, model: Locator JR Plus)
Single cuvette-based NucleofectorTM I device and certified cuvettes (formerly Amaxa Biosystems, now Lonza)
Inverted fluorescence microscope equipped with a UV, FITC, and TRITC filter sets (Olympus IX71) and attached digital camera
(Optional) Live-cell imaging system IncuCyte ZOOMTM (Essen BioScience, MI) for time-lapse microscopy of cultured cells plated in multi-well dishes
Software
(Optional) IncuCyte Zoom 2015A, Rev 1 for image capture and video assembly of time-lapse microscopy data
Image analysis software, ImageJ-FIJI (NIH), free-access software available at https://imagej.net/software/fiji/. Used to estimate the covered area (confluency) in selected phase contrast images using the area measuring tool
Procedure
Protocol 1: Trypsinization, plating, and propagation
Human Schwann cells isolated from nerve tissues are highly proliferative in the presence of added mitogens. At least two distinct mitogenic factors, namely heregulin (an ErbB agonist also known as neuregulin/NRG and glial growth factor/GGF) and forskolin (an agent that elevates intracellular cAMP), are needed for effective propagation (Bunge et al., 2017). For this reason, standard protocols for hSC culturing use a DMEM-based medium formulation containing heregulin (usually provided as a recombinant peptide at a nanomolar dose), forskolin (provided in the micromolar range), and serum (provided in the form of FBS) (Casella et al., 1996). This supplemented formulation (herein referred to as high proliferation or HP medium) supports 3–4 consecutive rounds of subculture of adult nerve-derived hSCs (Figure 1). The combination of heregulin and forskolin is optimal for hSCs. cAMP-elevating agents are known to both potently drive heregulin-dependent hSC proliferation and concomitantly reduce fibroblast growth (Rutkowski et al., 1992 and 1995).
The following sections explain how to propagate cultures from a confluent plate containing >90% hSCs, preferably from primary cells, as shown in the first paper of the series (Aparicio and Monje, 2023). A discrimination is made between routine (Protocol A) and controlled (Protocol B) serial passaging. Routine passaging at a non-constant ratio is practical for small-scale experimentation, but the lifespan of donor-relevant cultures may be hard to predict. Controlled passaging is suggested for preparing cryogenic stocks and comparing cultures from different donors (or lots), to ensure that cells have undergone roughly an equivalent number of cell divisions in culture by the time of experimentation. We recommend starting this protocol using cells at P0 to allow substantial propagation by passages-2 or -3. In all cases, researchers should pay attention to the growth attributes of each cell batch by performing daily observations and running appropriate tests. For reference, the growth characteristics of a representative, proliferative hSC culture are depicted in Figure 2. Videos 1 and 2 illustrate the distinctive dynamics of early-passage (non-senescent) and late-passage (senescent) hSC cultures, respectively.
Figure 2. Progression of a proliferative human Schwann cell (hSC) culture. Images A–F and insets i–iii provide an example of the temporal course of changes in a typical, low-passage hSC culture plated in HP medium (Protocol 1A). Notice that the cells divide quickly and asynchronously as soon as they are plated (H). The development of a pattern of alignment (C–F), a defining characteristic of SC cultures in vitro, is time and density dependent under these conditions. Cell–cell alignment usually occurs at or around confluency. The time of arrival to confluency (C, F, G) depends on various factors intrinsic to the hSC populations and the extracellular environment. The phase contrast images were selected from automated video-imaging microscopy analysis using IncuCyte ZOOMTM (see Video 1). The covered area per time point (G, graph) was calculated from the respective mask images (D–F) generated with ImageJ software.
Video 1. Growth of a typical low-passage human Schwann cell (hSC) culture. A single cell suspension of adult hSCs (proliferative) obtained by trypsin dissociation was plated on a laminin-coated dish and incubated in HP medium up until confluency, as described in Protocol 1A. Notice the fast adhesion of the cells, the changes in cell morphology (including process extension and alignment), the rapid cell migration, and the asynchronous appearance of mitotic figures. This culture can be considered confluent and ready to use by the third day of culture. The cells were imaged using IncuCyte ZOOMTM using 20× objective lenses. Individual images were taken every 20 min for a total time of 84 h from the onset of plating. Some relevant elements were highlighted in this and other videos.
Video 2. Growth of a typical high-passage human Schwann cell (hSC) culture. A single cell suspension of adult hSCs was obtained, plated, and imaged as described in Video 1. The cells adhere to the substrate and extend processes but most cells in this culture can be considered senescent. Senescent hSCs either fail to undergo cell division or do so at a very low rate, as revealed by the sporadic appearance of mitotic figures. Some senescent hSCs contain conspicuous vacuoles that are clearly observable at early time points and seem to be reduced over time. Whereas some individual cells develop an expanded cytoplasm and acquire a fibroblast-like morphology, others are bipolar and extend long processes resembling the phenotype of proliferative cells. Nevertheless, stable cell-to-cell alignment and pattern formation may not be achieved, as shown in this image series. Floating debris (high contrast puncta) are usually seen in senescent hSC cultures.
Routine passaging
Starting with a plate containing hSC cultures that have reached confluency (see Figure 2C–2F), remove the HP medium (see Recipe 2), and rinse the cultures with 10 mL of HBSS without calcium and magnesium.
Obtain a single cell suspension by controlled dissociation using TE dissociation solution (see Recipe 3). Use 5 mL of the 1× TE solution for a 10 cm tissue culture dish. Monitor the progression of the trypsinization by phase contrast microscopy to avoid overexposure of the cells to the action of the trypsin.
When the cells detach from the dish, add 10 mL of LP medium (see Recipe 1) directly onto the cultures to stop trypsinization.
Collect the cell suspension and transfer it to a 50 mL conical centrifuge tube. Rinse the dish with 5 mL of LP medium to collect the remaining cells. Confirm that no cells remain in the dish by phase contrast microscopy.
Centrifuge the cells at 200× g for 8–10 min at 4 °C and resuspend the cell pellet in HP medium for re-plating onto laminin-coated dishes.
Count the cells using an automated cell counting device or a hemocytometer and estimate viability according to the method of choice (e.g., Trypan blue or propidium iodide exclusion assays). The percentage of live cells should be high (>90%) after this and other routine operations. Obtaining a single-cell suspension is desirable to accurately estimate cell counts and allow an even distribution of the cells inside the culture dish.
Prepare laminin-coated dishes to be used fresh on the next day (applicable to all protocols). Briefly, to coat one 10 cm culture dish, thaw the laminin stock slowly at 4 °C and prepare a working solution containing 55 μg of laminin in 10 mL of DPBS. Incubate the plate ON at 4 °C on a flat surface up until use. Scale the volume of laminin solution according to the surface of the dish or well. It is recommended to use 0.7–1 μg of laminin per cm2 of coated surface. Sequential coating of dishes with PLL-laminin increases cell adhesion and recovery, e.g., after purification or transfection. Double coating is not needed for regular passaging if the goal is to simply expand the cultures.
Plate the cells in laminin dishes by homogeneously dispersing them at the ratio that best suits the experimental needs. A 1:3–1:5 plating ratio in HP medium is suitable for most applications. This density can render a confluent plate in <5 days (Figure 2), especially when using low-passage cells. Consider that batch variability can occur in this and other cellular responses.
Place the dishes inside a CO2 incubator immediately after seeding. hSCs normally attach within one hour after plating; use phase contrast microscopy to confirm that the cells have attached properly and extended processes after plating (Video 1) (see Note a).
Replace the medium two to three times per week with HP medium until the cultures reach confluency (see Notes b–c).
Repeat steps A1–A10 if further expansion is desired. Use the same conditions described above for trypsinization, plating, and growth regardless of the round of subculture. Monitor arrival to confluency as shown in Figure 2.
Controlled serial passaging
Starting with a confluent 10 cm plate of cells at P0, prepare a single-cell suspension, count the cells, and estimate their viability, as described in Protocol A. Next, plate 5 × 105 viable cells in a 10 cm dish in 10 mL of HP medium (see Note d).
Culture the cells in a CO2 incubator up until arrival to confluency. A confluent plate obtained in this manner can be considered an established culture at passage-1 (P1).
Proceed as described above to passage the cells but maintain the initial plating density at 5 × 105 cells per dish and per passage, which is the equivalent of a 1:10 expansion ratio in each round in proliferative cultures. If passaging is performed in this manner, each batch can be propagated effectively usually up to passage-4 (P4) (Figure 2).
Notes:
hSCs tend to spontaneously adhere to one another and form clumps of various sizes while in suspension. Cells within small clumps usually disperse evenly once the clumps attach to a laminin substrate, but big clumps may fail to attach. It is recommended to change the medium at this time only if dead cells, clumps, or excessive floating debris are observed.
It is recommended to allow cultures to reach confluency before sub-culturing them, so as to maximize cell yields in each passage. A confluent 10 cm plate usually contains 4 × 106–6 × 106 hSCs. The time needed to reach confluency can range from 4 to 10 days depending on the initial density, passage number, donor’s age, and other factors.
Two hallmarks of confluent cultures are the observation of an aligned configuration (explained above) and the disappearance or drastic reduction of mitotic figures (Figure 2). Confluent cultures must be used quickly or passaged to a new dish to prevent cell detachment.
Controlled passaging maximizes the efficiency of cell expansion per round of subculture. This protocol recommends using a plating density of 5 × 105 cells in each round (or a 1:10 expansion ratio) preferably starting with a confluent P0 culture. If passaging is performed as suggested, yields in the order of >108 cells can be obtained as soon as the second round of passage (Figure 1). The total cells obtained per batch are mainly linked to the cell yields from the initial harvest.
Protocol 2: Cryopreservation and transfer
This section describes the preparation of cryopreserved stocks of hSCs (Protocol 2A) and their re-plating after thawing (Protocol 2B). Cryogenic storage of hSCs is a standard practice that greatly facilitates in-house experimentation and the transfer of cell stocks to other laboratories. For this reason, this section describes simple protocols for transferring of cells in the form of live (adherent cultures in flasks) or banked stocks (frozen cells in cryovials) (Protocol 2C).
hSCs can be cryopreserved at any passage without detrimental effects on viability or adhesion post-recovery (our empirical observations). Cryopreserved hSCs have been used in various in vitro and in vivo approaches by our group (Monje et al., 2018) and others (Kohama et al., 2001; Bastidas et al., 2017). It is recommended to use healthy hSC cultures harvested in the logarithmic or exponential growth phase for optimal preservation and recovery. Another important recommendation is to prepare a master stock of hSCs at passages-1 or -2, as well as working stocks of the derivatives from these cells at passages-2, -3, and -4. Maintaining sufficient cryogenic stocks of cells collected at low passages can allow researchers to reinitiate the cultures at any time and use cells from the same batch and passage in independent experimental rounds (Monje et al., 2018).
Preparation and storage of cryogenic stocks
Once the hSC cultures are approximately 80% confluent, harvest the hSCs via trypsinization and collection by centrifugation (see Protocol 1, steps A1–A5).
Discard the supernatant and gently resuspend the cell pellet in ice-cold freezing medium (see Recipe 4) at a density of 1 × 106–2 × 106 cells/mL (see Note a).
Gently resuspend the cells by pipetting up and down using a 10 mL pipette (only once or twice) to obtain a homogeneous cell suspension.
Aliquot 1.5 mL of the cold cell suspension directly into properly pre-labeled cryogenic vials on ice. Work as fast as possible. This is a time- and temperature-sensitive step that can seriously affect the survival and recovery of the cells after thawing.
Immediately transfer the cryogenic vials to an isopropanol-filled polycarbonate freezing container to be placed in a -80 °C freezer.
Twenty-four hours later, transfer the cryogenic vials to a liquid nitrogen tank for long-term storage.
Retrieval of cryopreserved hSCs
Transfer the cryogenic vials directly from the liquid nitrogen tank to a safe container filled with dry ice for transportation to the biosafety cabinet.
Thaw the cells quickly by placing the vials in a 37 °C water (or bead) bath up until ~70% of the liquid volume has melted.
Transfer the cell suspension to a 50 mL conical centrifuge tube containing at least 15–20 mL of ice-cold LP medium.
Collect the cells by centrifugation at 200× g for 8 min at 4 °C. Remove the supernatant and resuspend the cell pellet in 10 mL of HP medium. Gently pipette the cells up and down no more than twice using a 10 mL pipette to obtain a homogeneous cell suspension.
Plate the cells in a 10 cm laminin-coated dish, incubate them in a CO2 incubator, and culture them as described in Protocol 1. Monitor the condition of the cells by phase contrast microscopy 1–3 h after plating. Most cells are expected to be attached to the substrate by this time. Lack of attachment may indicate poor viability or a problem with the substrate.
The following day, use a phase contrast microscope to confirm that the cells have attached properly and extended processes. Change the medium at this time only if floating dead cells or excessive debris are observed.
Monitor the progression of the culture daily. The increase in cell density should be obvious as soon as two days after plating (see Figure 2).
Packaging and shipping of cultured hSCs
To ship cryogenic stocks, retrieve the stocks from the liquid nitrogen tank and place them in a leak-proof plastic bag or conical tube (secondary container). Transfer the cells as soon as possible into a Styrofoam container filled with dry ice (see Note b).
To ship live cells, prepare a cell culture of hSCs in HP medium at 40%–60% confluency in a T25 or T75 culture flask double-coated with PLL and laminin for stronger adhesion. Use flasks with a plug-seal cap (not vented caps). On shipment day, fill the flasks with warm HP medium to full capacity. Tighten the cap and seal it with parafilm. Wrap the flasks with absorbent paper towels and place them inside a leak-proof bag or container. Position the flasks horizontally inside a small Styrofoam box surrounded by bubble wrap or soft paper, ensuring that the flask remains in place during transit. Ship the cells at room temperature (see Notes b–c).
Send the package using an express courier service for delivery between 24 and 72 h, preferably a next-day delivery.
Once the cells arrive at the destination, retrieve the cells as described in Protocol 2B for frozen stocks and Protocol 1A for live cell cultures.
Notes:
Ready-made cell freezing medium is available from various commercial sources. We have tested RecoveryTM with good results in the cryopreservation of rat SCs (Andersen and Monje, 2018). This medium is also suitable for hSCs and can be used in replacement of our home-made freezing media (Recipe 4).
Take all possible precautions to prevent leakages during transit by double-bagging and adding absorbent paper around the flasks. General recommendations for safe shipping of biohazards, such as using leak-proof packaging materials, must be followed, according to the requirements of postal service or customs authorities, as applicable. If possible, send a second cryogenic tube or flask as a backup.
Cells growing in flasks should be shipped at room temperature. Add thermal protection (insulation) to ensure safe transport. Do not add cold pads. hSCs proliferate even at room temperature in HP medium, so avoid shipping cultures that are close to reaching confluency.
Protocol 3: Purification methods
Various types of non-glial cells from the epi- and perineurial layers, the vasculature, and the endoneurial matrix can get introduced into the hSC cultures despite precautions taken during the dissection of the fascicles. Non-glial cells do not negatively influence hSC behavior in culture. Yet, contaminating cells may be undesirable experimentally or hamper data interpretation in some in vitro and in vivo studies. For instance, it has been argued that transplantation of hSCs containing a higher than acceptable number of fibroblasts leads to excessive collagen matrix deposition in the central nervous system (Brierley et al., 2001).
The hSC culture medium containing forskolin and heregulin selectively promotes hSC growth over that of fibroblasts and often causes a progressive enhancement in hSC purity (Levi et al., 1995). However, the selective pressure of media components is insufficient to prevent the spread of fibroblasts (Peng et al., 2020). Antibody-based technologies such as magnetic-activated cell sorting (MACS) and fluorescence-activated cell sorting are among the most efficient methods available for fibroblast removal (Morrissey et al., 1995b; Weiss et al., 2016). MACS is highly selective, scalable, and adaptable for direct purification of hSC cultures (Peng, Sant et al. 2020). However, there are cost-effective and simpler methods seemingly efficacious for fibroblast removal. Immunopanning, which utilizes a solid surface coated with a specific antibody (or protein ligand) immobilized onto the surface of a cell culture plate, is a traditional way to purify SCs from mice (Lutz, 2014) and humans (Fregien et al., 2005). The difference in cell size and adhesion properties between hSCs and fibroblasts, which leads to more expedited sedimentation and attachment of the latter cells, has been exploited to separate them from hSCs. This method is advantageous in clinical applications because it does not introduce reagents or chemicals that can pose a risk to patients (Khan et al., 2021). Controlled trypsinization and a shock of cold medium are also suitable for hSC enrichment (Haastert et al., 2007; Weiss et al., 2016). However, complement-mediated killing of Thy1+ positive fibroblasts, a widely used method to purify rat SCs (Brockes et al., 1979), is ineffective for hSC cultures because certain contaminating cells do not seem to express cell surface Thy1 (our empirical observations).
This section features two distinct panning protocols (i.e., cell culture plastic panning and immunopanning) and briefly introduces our nanobead-assisted MACS protocols, described previously in Ravelo et al. (2018), to help researchers balance options in dealing with fibroblast overgrowth. All protocols start with preparing hSCs in suspension (Protocol 1). Best results are obtained using established, myelin-free cell cultures (passage-1 or higher) that result in a clean, highly viable single-cell suspension whose numbers can be estimated properly. The methods described below are unsuitable for separating hSCs after nerve tissue dissociation because of interference with myelin and other tissue-derived debris. Before starting the purification procedures, make sure the cultures contain typical hSCs, as judged by cell surface expression of NGFR and/or O4 (Peng et al., 2020). For details on our suggested staining protocols, see the accompanying paper Monje (2023).
Fibroblast depletion by cell culture plastic panning
Rinse the hSC cultures with HBSS, obtain a single-cell suspension using a 1× TE solution, and collect them by centrifugation in LP medium, as described in Protocol 1. In this and all purification procedures, estimate the total number and viability of the cell suspensions before purification (Ravelo et al., 2018). The number of cells to be purified (step 2) should be based on the viable cell counts (see Note a).
Plate no more than 3 × 106 total cells suspended in a 7–10 mL of HP medium in an uncoated, cell culture-treated 10 cm dish and transfer the cells immediately to the CO2 incubator.
Incubate the cell suspensions at 37 °C for 15 min to allow large-diameter cells, mainly fibroblasts, to be deposited on the bottom of the dish. Do not leave the cells in the incubator for a longer time. Otherwise, hSCs will simultaneously attach to the plastic.
Remove the dish from the incubator and slowly aspirate the supernatant, which contains a cell suspension consisting mainly of hSCs. (Optional) Slowly add 5 mL of LP medium to one side of the culture dish and tilt the plate to collect the hSCs that are not attached, with the caveat that this procedure may also detach some of the fibroblasts.
Transfer the supernatant containing hSCs into a PLL-laminin-coated 10 cm dish for recovery in LP medium or use the cells directly in experimentation. Alternatively, transfer the cells into a 50 mL conical tube, collect them by centrifugation (200× g for 8 min) and plate them at the desired density in the medium of choice.
hSC enrichment by immunopanning
Immunopanning plates are prepared by pre-adsorbing secondary antibodies directly to the surface of a plastic Petri dish before the adsorption of primary antibodies (Protocol 3B1). A single-cell suspension is then plated onto the coated plates for antibody-mediated hSC binding (Protocol 3B2).
B1. Preparation of antibody-coated plates
Prepare 10 mL of anti-mouse immunoglobulins solution (unconjugated secondary antibody, see Recipe 5) for each dish to be coated.
Coat one or more 10 cm Petri dishes (non-cell culture treated plastic) with 10 mL of immunoglobulins solution each and incubate them ON in a 4 °C refrigerator. Place this dish on a flat surface ensuring that all areas are well-covered with liquid. Safeguard the dishes from contamination while stored at 4 °C by placing them inside a sterile container. Consider that the surface of untreated plates is hydrophobic compared with regular tissue culture plates, and movement of the dish will be needed to spread the liquid evenly.
The next day, remove the immunoglobulins solution by aspiration and rinse the dishes three times with cold L15 medium to remove unbound antibodies.
Immediately after, add 10 mL of the primary monoclonal antibody solution and incubate the plates for at least 2 h in the 4 °C refrigerator. As source of monoclonal antibodies, we use the culture supernatant (undiluted media) produced in house from hybridoma cell lines, HB-8737 (NGFR) and O4 (see Note b).
Remove the primary antibody solution by aspiration and rinse the dishes three times with cold L15 medium to remove unbound primary antibodies. These dishes can be stored ON at 4 °C on a flat surface. Do not remove the medium from the last wash up until the cells are ready for panning (Protocol B2).
B2. Cell purification
Gently resuspend a single cell preparation of hSCs in ice-cold L15 medium right before the panning experiment. For instance, prepare 3 × 106–5 × 106 cells in 10 mL of L15 medium. Maintain these cells on ice making sure no clumps are formed before panning.
Remove the L15 medium from the last wash (Protocol B1, step 5) and plate the cell suspensions on the antibody-coated dishes immediately after.
Place these dishes on a flat surface at 4 °C for 20 min without disturbing them. Use phase contrast microscopy to confirm that a proportion of the cells have adhered to the substrate by gently moving the plate from side to side. This movement of fluid also dislodges loosely bound cells. The time of incubation is determined empirically by visual observation. If attached cells are not observed after 20 min, incubate the dishes for an additional 10 min at 4 °C to allow more time. Do not prolong the incubation unnecessarily. Make sure the hSCs have settled on the plate but still maintain a rounded shape before proceeding with step 4.
Remove the media manually with a transfer pipette and gently wash the cells three times with L15 medium to rinse off non-attached cells (e.g., these cells will be mostly fibroblasts in HB-8737/NGFR immuno-panning experiments).
Add 10 mL of LP or HP medium and gently scrape the attached cells off the panning dish. Detach the cells gently from the 10 cm plate into the LP medium with a 2 cm blade cell lifter. Slightly tilt the dish and scrape the cells from the edges to the center to ensure all areas are covered. Confirm that the cells were detached by phase contrast microscopy. This is a time-sensitive step. Caution should be taken during the scraping procedure to prevent mechanical damage to the plasma membrane. Use a gentle dissociation reagent such as TrypLETM select to lift the cells if poor viability is observed while optimizing this step.
Transfer the cell suspension to a conical 15 mL tube for collection by centrifugation (200× g for 8 min) or plate them directly onto a PLL-laminin-coated dish for recovery, as explained in Protocol 3A, step 5.
hSC enrichment by magnetic-activated cell sorting
Prepare a single cell suspension of hSCs. Allocate at least 3 × 106 viable cells for each purification step.
Follow the instructions in our step-by-step MACS protocol for positive selection of NGFR+ and O4+ hSCs (Ravelo et al., 2018).
Analyze the resultant cell products by phase contrast and fluorescence microscopy to confirm the purity of the hSCs. Representative results are shown in Figure 3 and Videos 3, 4, and 5.
Figure 3. Purification of human Schwann cell (hSC) cultures. The phase contrast images compare a representative adult nerve-derived hSC culture before (mixed culture, left panels) and after magnetic-activated cell sorting (MACS)-assisted cell separation (purified hSCs, middle panels), as per Protocol 3C. The fibroblast cultures are shown in parallel for comparison (purified fibroblasts, right panels). The phase contrast images were selected from IncuCyteTM Videos 3, 4, and 5, and the respective mask images (ImageJ, panels in magenta) were generated to denote the degree of confluency within the first- and third-days post-purification, as indicated. The hSCs were identified as NGFR+ cells (green) by immunofluorescence microscopy analysis (lower panels). NGFR- cells can be regarded generically as fibroblasts. Total cell nuclei were labeled with DAPI (blue), and proliferating nuclei were labeled with EdU (red). Notice that the post-MACS cell products (i.e., hSCs and fibroblasts) are both highly viable and proliferative cells.
Video 3. Characteristics of unpurified human Schwann cell (hSC) cultures. A single-cell suspension of adult hSCs (proliferative) was plated and imaged as described in Video 1. Notice the fast arrival to confluency but delayed or impaired cell–cell alignment (compare with Video 4), likely because of the presence of abundant intermixing fibroblasts in these populations.
Video 4. Characteristics of magnetic-activated cell sorting (MACS)-purified human Schwann cell (hSC) cultures. The MACS-purified hSCs (retained fraction) were derived from the mixed cultures shown in Video 3. The cells were plated immediately after purification. This video shows the fast adhesion, proliferation, and alignment of the hSCs, which can be considered evidence in support of the viability and biological activity of the purified cells.
Video 5. Characteristics of magnetic-activated cell sorting (MACS)-purified fibroblast cultures. The MACS-purified human fibroblasts (eluted fraction) were obtained from the cell cultures shown in Video 3. The cells were plated immediately after purification. Notice that the fibroblasts are highly proliferative, and their phenotype is clearly distinguishable from that of hSCs. Most fibroblasts exhibit a flat and expanded morphology with a conspicuous reticulated cytoplasm at confluency.
Notes:
Perform all necessary controls during the optimization phase. For instance, it is useful to set out a sample of the original cell suspension and plate it in a multi-well dish to estimate the purity of the original populations in comparison to those obtained at the end of the purification procedure. One way to confirm the effectiveness of cell purification is to perform an immunostaining analysis using antibodies against hSC-specific markers before and after purification. An example of such analysis is provided in Figure 3. Our publication by Peng et al. (2020) provides additional experimental data on hSC purification methods, cell-based assays, and analysis of results.
The hybridoma ATCC #HB8737 was selected because it produces a monoclonal antibody against the extracellular domain of NGFR, a stable cell membrane marker for cultured hSCs (Peng et al., 2020). This antibody does not recognize rodent NGFR (Ross et al., 1984) and is suitable for live-cell labeling and immunopanning of hSCs. An alternative panning antibody is O4 (Bansal et al., 1989), which can be used for positive selection of stage-specific, O4-expressing hSCs (Ravelo et al., 2018).
Protocol 4: Cell labeling and gene delivery
SCs can be labeled directly with vital fluorophores or indirectly by introducing plasmids and viral vectors while retaining their normal phenotype (Mosahebi et al., 2000 and 2001; Hoyng et al., 2015). Like many other primary cells, hSC cultures are hard to transfect. In our experience, transient transfection of adherent nerve-derived hSCs with conventional lipid-based transfection reagents seldom overpasses 5% transfection efficiency regardless of the brand or formulation used (unpublished). However, we and others have found that Nucleofection technology is appropriate for introducing plasmid DNA into hSCs in suspension with efficiencies that can average 40%–60%, as determined by the levels of GFP expression from reporter plasmids (Haastert et al., 2007; Monje et al., 2008). Stable, long-term fluorescent labeling of cultured hSCs can be accomplished by lentiviral or retroviral infection (Monje et al., 2018). However, the initial transduction efficiency with retroviruses is generally low even when using low-passage cultures, as only hSCs in cell division can be infected (unpublished).
The issue of toxicity in relation to the efficiency of labeling or genetic modification is the most important challenge to overcome in any transfection or infection protocol. hSCs are particularly sensitive to changing environmental conditions. The additives needed for efficient transfection or transduction or cell selection (e.g., with antibiotics) can cause substantial cell loss. In certain cases, transient labeling with membrane-permeable fluorescent dyes is sufficient for the visualization of cells over a period of time, e.g., to monitor the morphology of SCs in vitro (Monje et al., 2009) or after transplantation in experimental animal models (Li et al., 2003).
The following sections describe our recommended methods for transient and long-lasting fluorescent labeling of hSC cultures for live-cell imaging and tracing. Our transfection methods are suitable to achieve overexpression of reporter genes and membrane receptors (Monje et al., 2008), and possibly other forms of genetic modification.
Transient labeling with vital fluorophores
One practical way to fluorescently label hSCs with minimal cytotoxicity is with CellTrackerTM. This fluorescent dye is well-suited to detect dynamic changes in cell size and shape by fluorescence microscopy. CellTrackerTM freely passes through the plasma membrane and transforms into a cell membrane-impermeant fluorescent product that is maintained for several days in the hSCs’ cytoplasm. The labeling procedure involves the addition of the reagent in serum-free culture medium followed by washes to remove the soluble reagent. Cells are ready to use soon after the labeling treatment. The fluorescence intensity is strong even after fixation with aldehyde-based fixatives and can be combined with antibody-based staining or staining with nuclear dyes.
Prepare a 10 cm plate of cultured hSCs in HP medium, preferably at confluency (Figure 2).
Dissolve the lyophilized CellTrackerTM powder with high-quality DMSO to a concentration of 10 mM, as suggested by the manufacturer, immediately before use.
Rinse adherent hSCs with serum-free DMEM to remove traces of serum.
Dilute the freshly prepared CellTrackerTM stock solution to a working concentration of 6.5 μM in pre-warmed (37 °C) serum-free DMEM and directly add it to the cells. For 10 mL of medium, use 6.5 μL of 10 mM CellTrackerTM stock together with 30 μL of 20% (v/v) Pluronic F-127, a non-ionic detergent used as dispersing agent to enhance the labeling intensity. The addition of Pluronic is optional. Higher concentrations of Pluronic are toxic to hSCs.
Incubate the cells for 30 min in the CO2 incubator. Check the labeling efficiency by fluorescence microscopy before proceeding with the next step. Be strict with the labeling time so as not to compromise the health of the cells.
Rinse the cells with a sufficient volume of pre-warmed serum-free DMEM to remove the background fluorescence. Next, add 10 mL of HP medium and incubate the cells for at least 30 min at 37 °C (recovery) before experimentation.
(Optional) Include an additional incubation step with Hoeschst-34580 (1:1,000 dilution in LP medium) to concurrently stain the cell nuclei. Hoeschst-34580 is well-tolerated by hSC cultures. For reference, Figure 4A shows typical hSC cultures (live-cell imaging) after combined CellTrackerTM/Hoeschst-34580 staining.
(Optional) Re-stain the cells 48–72 h after by repeating steps A2–A6 if the fluorescence intensity declines.
Figure 4. Fluorescent cell labeling of cultured human Schwann cells (hSCs). (A) Transient cell labeling with CellTrackerTM showing homogeneous staining of all viable cells. (B) Persistent cell labeling via infection with EGFP-encoding lentiviral vectors. (C–F). Transient transfection of GFP-encoding plasmid vectors via Nucleofection. In A, cells were treated for three days with combined heregulin (10 nM) and forskolin (2 µM) in the absence of serum, labeled with CellTrackerTM and Hoescht-34580, and imaged 1 h post-labeling. In B, cells were infected at passage-2 and subjected to another round of expansion in HP medium before imaging. Notice the nonhomogeneous levels of EGFP expression in individual cells. Lentiviral infection does not significantly alter the phenotype or the proliferation of hSCs in vitro even after several passages. In C–D, hSCs and rat SCs (control) were transfected via Nucleofection (program A-33, using pmaxGFP reporter plasmid), and visualization was performed four days after the procedure. Expression of pmaxGFP is maintained for at least one week. Yet, transfected hSCs can show signs of stress, as evidenced by the unusual stellate morphology of the cells (C–D).
Transient transfection using Nucleofection
Nucleofection is a technology that applies an electrical pulse to momentarily create small pores in the plasma and nuclear membranes to enable rapid delivery of nucleic acids into the nucleus. This technology has shown superior performance for transfecting various primary cells, including hSCs. The conditions outlined below were set up to transfect expanded, donor-derived hSCs in suspension using cuvettes and reagents provided by the manufacturer (Amaxa Biosystems, now Lonza). This low-throughput method, which uses high cell numbers (106–107 cells) per transfection reaction, is suitable for performing biochemistry (e.g., gene reporter assays and western blotting) and fluorescence microscopy studies using transfected hSCs (Monje et al., 2008). Variants on the original Nucleofection technology are currently available but have not been tested in our laboratory.
Prepare a PLL-laminin-coated 6-well plate containing 1.5 mL of HP medium and incubate it in the CO2 incubator for plating the hSCs immediately after transfection.
Starting with a confluent 10 cm plate of hSCs, harvest the cells by trypsinization, count the cells, and estimate their viability. Use a healthy hSC culture preferably collected at passage 1–3, as shown in Figure 2C–2F.
Prepare an aliquot of 5 × 106 viable cells in LP medium and collect them by low-speed centrifugation at room temperature to obtain a very loose cell pellet. This is a sensitive step. Centrifugation should not exceed 150× g (for up to 8 min) in a swinging bucket centrifuge.
Remove the supernatant completely, resuspend the cell pellet in Nucleofector Solution from the Basic NucleofectorTM kit for primary mammalian glial cells (100 µL per sample), and add 1 µg of pmaxGFPTM Vector (positive control) or 1–4 µg of plasmid DNA of choice (experimental) following the instructions provided by the manufacturer (see Note a).
Immediately transfer the cell/DNA suspension into a certified cuvette, remove air bubbles, and close the cuvette with the cap.
Select the appropriate Program (A-33 or O-17) for the NucleofectorTM I device or an equivalent program on other models. Insert the cuvette into the holder and apply the selected program. Program A-33 is preferred for better recovery of viable hSCs post-transfection. Program O-17 renders a higher transfection efficiency (>40% as determined by pmaxGFP expression) with the caveat of increased cell loss.
Promptly add 500 µL of pre-equilibrated HP medium directly to the cuvette and gently transfer the sample in a drop-by-drop manner directly into the wells of the 6-well plate. Work quickly, as the cells are very sensitive at this stage. Plate the product of one transfection reaction (initially 5 × 106 cells) into two wells of a 6-well plate, as substantial cell death is expected due to the electrical shock. Transfer the plate to the CO2 incubator for stabilization without delay.
Observe the plate 3–4 h post-transfection to confirm cell attachment and change the medium to remove floating (dead) cells and debris.
Assay the cells in the 6-well plate or re-plate them into a new multi-well dish after they have recovered for at least 24 h (see Note b). Substantial expression of pmaxGFP is expected in the positive control condition (Figure 4C).
Infection with lentiviral vectors
Lentiviral particles can be used to achieve stable overexpression of a transgene of interest in virtually any cell type, including non-proliferating, terminally differentiated cells. Once integrated into the DNA of the target cells, long-term constitutive expression of a gene product can be achieved. We have used lentiviruses to transduce rat and human SCs with vectors encoding fluorescent proteins for direct visualization of cells in isolation and in co-culture with neurons (Monje et al., 2018). Transducing hSC cultures at a low passage (e.g., P1) and expanding them for at least another round (e.g., 1:10 ratio) is feasible and recommended to create a working batch of transduced cells for experimentation and/or storage by cryopreservation.
Standard practices for viral transduction are applicable for the infection of hSCs. The first step is to empirically determine the suitable multiplicity of infection or MOI (i.e., the number of viral particles needed per cell to effectively achieve transduction) to optimize gene delivery in hSCs, as the MOI directly correlates with the number of integration events and the expression levels of the transgene. A MOI of 1 (one viral particle per cell) is commonly used, but a higher MOI (usually 5–10) is often needed to achieve >90% GFP expression in rat and human SCs (our empirical observations). In addition, it should be noted that high infection levels can lead to reduced hSC viability possibly linked to transgene overexpression.
We routinely perform preliminary experiments to assess the functional titer of viral stocks for each new virus type and lot (Protocol C1). We do so in replicate samples using at least two independent hSC cultures because the infection efficiency (and associated toxicity) can vary from batch to batch and donor to donor. Testing a range of viral concentrations in small wells (multi-well plates) can aid in determining the volume of viral stock needed for larger cultures. Once the infection conditions are optimized, the desired quantity of hSCs can be infected and the cells used as such or after being selected with antibiotics to enrich in the infected population (Protocol C2).
C1. Determination of the optimal virus dose
For the virus titration curve, seed the hSCs in a 24-well dish coated with PLL-Laminin and plate ~50,000 hSCs per well in HP medium to obtain a sub-confluent culture.
The next day, replace the medium with 300 µL of transduction medium (TM) containing increasing concentrations of the viral particles for EGFP, mCherry, or the virus of choice (see Note c).
Incubate the cells overnight at 37 °C in the CO2 incubator.
The following day, remove the viral particles by replacing the TM with 500 µL of HP medium per well for optimal growth and recovery of the cells.
Three days after infection, observe the expression of reporter proteins by fluorescence microscopy imaging.
Determine the optimal MOI to be used in subsequent experiments by calculating the percentage of cells expressing the gene reporter vs. the total number of cells by image analysis or other methods.
C2. Amplification of transduced hSCs
Plate hSCs at a density of 1 × 106–2 × 106 cells in a 10 cm plate double-coated with PLL and laminin in 10 mL of HP medium. (Optional) Scale the cell density and volume of medium up or down for other plate formats.
When the cells reach ~40%–50% confluency (usually within two days post-plating), transduce them by replacing the medium with 6 mL of TM at the desired MOI, as determined in Protocol C1.
Proceed as described above for managing virus-transduced cells.
Analyze the cells three days after infection to confirm expression of the gene reporters. By this time, >90% of the cells should display high levels of the reporter gene (Figure 4B). These cells are ready for use in experimentation as such, after additional expansion (step 5), or after antibiotic selection (step 6).
(Optional) Subculture the virally transduced cells as recommended in Protocol 1A or B to generate larger batches of cells for experimentation or cryogenic storage. Transgene expression is expected to be stable after 2–3 additional rounds of expansion.
(Optional) Enrich the infected populations at an early passage by treating the cells with an appropriate antibiotic as determined by the selection gene encoded in the viral vector. For reference, we have used 0.5 μg/mL puromycin, 100 μg/mL hygromycin, and 3 μg/mL blasticidin to select virally transduced hSCs established at passage-1 (unpublished). The antibiotic should be provided in HP medium for as long as needed to eliminate all non-transduced cells, per visual inspection under the phase contrast microscope.
Notes:
Strictly follow the instructions provided by the manufacturer for the preparation, storage, and use of reagents (including the quantity and quality of plasmid DNA) and the use of the equipment. During the optimization phase, perform pilot studies with at least two batches of cultured hSCs, as transfection efficiency varies from batch to batch. Introduce the necessary controls to estimate the transfection rate while observing cytotoxicity due to the procedure, the expressed gene product, or other variables.
Transfected cells can be harvested for analysis or assayed directly in the 6-well plates. To better control the cell density and obtain accurate replicas for experimentation with the genetically modified cells, re-plate the transfected hSCs into 96-, 12-, or 24-well assay plates. Perform cell-based assays preferably within three days post-transfection since the levels of transgene expression decline thereafter.
The TM consists of HP medium containing viral particles at a given MOI. Estimate the MOI based on the information provided by the manufacturer of viral particles (e.g., as it relates to the concentration of p24 capsid protein in the stock) and the number of cells plated in the test well. If there is no prior experience with hSCs or the virus, start by testing a broader range of MOIs (e.g., 0, 0.5, 1, 2, 5, 10, 15, 30). Transduction enhancers such as polybrene (8 μg/mL) may be added to the TM with caution because additives can be toxic to hSCs. Use duplicate or triplicate cultures for testing each MOI condition.
Recipes
Low proliferation (LP) medium
Reagent [Stock concentration] Final concentration Amount
DMEM (or DMEM/F12) n/a 445 mL
FBS [100%] 10% 50 mL
GlutaMAX [100×] 1× 5 mL
Gentamycin [1,000×] 1× 0.5 mL
Total n/a 500 mL
Note: Balance the pH of the LP medium to make it slightly acidic (pH = 6) using a cell culture grade HCl solution; then, sterilize it by filtration using a 0.22 µm filtration unit and store it at 4 °C for up to one month. Gentamycin is the preferred antibiotic but can be replaced by penicillin/streptomycin. Do not include antifungal reagents in this or any other culture media because of cytotoxicity to the hSCs.
High proliferation (HP) medium
Reagent [Stock concentration] Final concentration Amount
Low proliferation medium n/a 500 mL
Heregulin-β1 [25 µM] 10 nM 200 µL
Forskolin [15 mM] 2 µM 69.25 µL
Total 500 mL
Note: Balance the pH, sterilize it by filtration, and store it as indicated for the LP medium. The HP medium induces optimal hSC proliferation when stored appropriately at 4 °C within a month of preparation. Do not freeze the HP medium.
TE dissociation solution
Reagent [Stock concentration] Final concentration Amount
Trypsin/EDTA [10×] 1× 1 mL
HBSS n/a 9 mL
Total 10 mL
Note: Dilute the 10× TE stock solution in ice-cold, sterile HBSS, and use it without delay to dissociate the cells. Do not freeze. Follow the instructions provided in our generic trypsinization protocol for storage and use of the TE stocks and working solutions (Andersen and Monje, 2018).
Freezing medium
Reagent [Stock concentration] Final concentration Amount
Decomplemented FBS 90% (v/v) 90 mL
DMSO [100%] 10% (v/v) 10 mL
Total 100 mL
Note: This medium is prepared fresh every time and maintained on ice up until the preparation of cryogenic hSC stocks. More information can be found in our generic cryopreservation protocol (Andersen and Monje, 2018).
Immunoglobulins solution
Reagent [Stock concentration] Final concentration Amount
Tris base, pH = 9.5
Immunoglobulins
0.05 M
n/a
99 mL
1 mL
Total 100 mL
Note: To prepare the 1 M Tris-buffer, dilute the Tris-base powder (121.14 g) in 800 mL of water, adjust the pH to 9.5 with 6 N or 1 N HCl (as needed), and add water to a final volume of 1,000 mL. Next, prepare 100 mL of a 0.05 M working solution from the 1 M Tris stock using sterile, cell culture grade water, and dilute the immunoglobulins at 1:100. Sterilize this solution by filtration and use it fresh to coat the immunopanning dishes.
General notes and troubleshooting
This section provides additional conceptual and technical information to understand the procedures, identify potentially problematic issues, and interpret the results. Implementing appropriate controls to verify the identity and function of the hSCs is highly recommended, as the cells that are passaged in vitro are expected to change (Figure 5, upper panel). Likewise, it is equally important to verify that all laboratory materials, including cell culture media, consist of endotoxin-free, sterile products optimal for hSC growth (Figure 5, lower panel).
Figure 5. Suggested quality control assessments in the propagation of human Schwann cells (hSCs). The recommended routine controls for the cell cultures (above) and the materials (below) are highlighted in the diagram. Conducting purity checks for cellular (e.g., fibroblasts) and non-cellular (e.g., myelin) impurities is most critical during the establishment of the cultures (P0–P1). Late-passage hSCs cultures (P4–P5) should be tested for senescence as soon as the rate of proliferation begins to decline. Viability assays are valuable indicators of cell health after routine manipulations (e.g., enzymatic dissociation) and during the post-thaw recovery. Though rare, appropriate assays may be needed if growth becomes abnormal (e.g., cells lose contact inhibition or proliferate in the absence of heregulin) or propagation is extended over passage-5, as these observations may indicate transformation or contamination with proliferative cell lines. Researchers can refer to the accompanying paper (Monje, 2023) for detailed protocols on how to perform identity, purity, bioactivity, and authentication assays.
Expansion and subculture
Substrate. hSCs should be plated on freshly prepared laminin-coated dishes because they do not properly attach to non-coated dishes or dishes coated with simple substrates such as PLL. Laminin is unreplaceable for hSC adhesion and proliferation in vitro, which is an important difference when compared to rat SCs. Other groups have used alternative substrates such as fibronectin and collagen for the culturing of hSCs (Vleggeert-Lankamp et al., 2004). However, their effectiveness to sustain hSC expansion through passaging (this protocol) has not been determined experimentally.
Culture media. Effective expansion of hSCs in the absence of neurons strictly relies on the addition of media supplements. The mitogenic factors heregulin and forskolin are equally required for optimal propagation of hSCs. Serum factors do not promote extensive proliferation of isolated hSCs. Yet, a 10% concentration of FBS should be used consistently to maintain the hSCs viable and adherent for long periods of time. We have not identified additional factors with strong mitogenic activity for cultured hSCs that could replace or complement the abovementioned ones (Monje et al., 2018). Bovine pituitary extract, a source of heregulin-like activity, is used to supplement the culture medium of rat SCs (Brockes et al., 1979). However, pituitary extract is not mitogenic for hSCs (our unpublished observations). The source of cAMP can be variable. Forskolin is preferred because it is a non-cytotoxic and reversible inducer of cAMP that can be provided alone or together with cholera toxin for optimal cAMP elevation and proliferation in hSC cultures (Rutkowski et al., 1992; Levi et al., 1995; Morrissey et al. 1995b). It has been argued that cholera toxin should be avoided in clinical hSC preparations due to its irreversible effects on adenylyl cyclase activity [discussed in Bunge et al. (2017)].
Certain experimental procedures, such as kinase and proliferation assays, require manipulations to be performed in mitogen- and serum-free media. However, removing defined or undefined media components is not recommended for routine culture because it leads to substratum detachment and hSC apoptosis. We have found that stepwise mitogen and FBS deprivation over a 2–3-day period is effective in eliciting cell cycle withdrawal (quiescence) without significant cell loss (Monje et al., 2006). Overall, changing the concentration of serum and mitogenic factors is discouraged during the propagation, purification, and labeling of hSC cultures.
Good practices and routine controls. hSC cultures change daily (Videos 1–5). For this reason, the periodic inspection of the cultures by phase contrast microscopy is an unreplaceable practice to detect abnormalities such as cell clumping and detachment, fibroblast growth, and microbial contamination. The morphology of hSCs varies under standard growth conditions, and dynamic changes in cell size and shape are expected. Yet, the steady progression towards confluency should be clearly observable (Figure 2 and Video 1). Researchers should consider that a bipolar cell morphology and the formation of paralleled bundles of aligned cells may or may not occur in healthy hSC cultures, as this pattern of growth is often evident in confluent cultures only (Figure 2 and Video 1). Indeed, cell–cell alignment may be widespread across the dish or formed in local areas where the cell density is appropriate [for an example, see Monje (2020)]. In addition, variability should be expected in the expansion rate and other properties of hSC cultures due to cell intrinsic and extrinsic factors. Therefore, each batch- or donor-specific culture should be handled and investigated individually from the onset.
Always follow best cell culture practices and the recommendations from manufacturers in the preparation, storage, and use of culture media, supplements, buffers, and other reagents (Figure 5). Controlling the pH in buffers and media is critical to maintain hSC viability. hSCs are stable in acidic pH but are severely damaged if medium turns alkaline, especially while they are in suspension. Recalibrating the pH of buffers and media as frequently as needed and working fast while operations are conducted in the biosafety cabinet can avoid potentially harmful pH fluctuations. It is useful to set up the incubators at 8%–9% CO2 and adjust the pH of all solutions to be slightly acidic to prevent media alkalization during the handling of cells (see Recipes).
Limits to expandability. Passaging is the most defining factor affecting the expansion rate of established adult hSC populations because it eventually leads to hSC senescence. The age of the donor can be influential rather than decisive on the expandability of hSC stocks. Whereas cells from younger donors tend to proliferate faster (Boyer et al., 1994), advanced age does not preclude the derivation of proliferative hSC cultures (Levi, 1996). We have not discovered how to experimentally overcome hSC senescence in vitro but found that early passage hSCs can become senescent due to in vitro–induced stress rather than genetic influence (Monje et al., 2021). Senescent cultures may be discarded. A good practice is to establish and maintain enough cryogenic stocks (Protocol 2) to reinitiate the culture once signs of senescence are manifested.
Importantly, we have not observed spontaneous or induced (oncogene mediated) immortalization of cultured hSCs (unpublished). In fact, the creation of genetically modified hSC lines is restricted due to the hSCs’ resistance to immortalize in vitro by genetic ablation of tumor suppressors (Petrilli and Fernandez-Valle, 2018). There is precedent for hSC immortalization by ectopic co-expression of hTERT and SV40 large T-antigen (Lehmann et al., 2012). Yet, most available hSC lines are derived from peripheral nerve sheath tumors (Lee et al., 2004; Dilwali et al., 2014) rather than primary, normal hSCs. Perform an authentication analysis in case signs of aberrant growth (e.g., uncontrolled proliferation) become evident (Figure 5).
Cryogenic stocks. Cryopreservation is feasible and effective when using hSC cultures collected at any passage number, except for P0. P0 cultures are not yet established and usually contain abundant intracellular myelin and/or debris detrimental to the viability of the stocks (our empirical observations). A good practice is to create master stocks using purified hSCs from passages 1–2 along with working stocks using purified hSCs from passages 2–3 as per Protocol 1B, from each and all donors. Cultures established from cryopreserved hSCs are nearly identical to those obtained from fresh isolates as evidenced by their morphology, expression of SC-specific markers, and proliferation rates (Bastidas et al., 2017).
Standard safety practices for cell cryopreservation should be used to minimize the risk of microbial contamination, avoid temperature fluctuations, and maintain cell viability. Transfer the cells to the liquid nitrogen tank, preferably within 24–28 h, as the viability of hSCs declines if the stocks are stored at -80 °C for one week or longer. In addition, prevent unnecessary exposure of cells to the toxic effects of the DMSO during the freezing procedure and after thawing. It is good practice to dilute the freezing medium with LP medium (at 1:10 or a higher ratio) while thawing the cells to readily reduce the concentration of the cryo-protector before centrifugation and plating. Performing a stability study of the cryogenic stocks is recommended when cryopreservation methods are implemented for the first time.
Cell purification. Knowing the constitution of the cultures beforehand can help experimenters rationally choose an appropriate purification method. The panning protocols are relatively inexpensive and can be applied to reduce the content of contaminating cells in hSC cultures containing a low proportion (i.e., 30% or less) of fibroblasts. MACS is superior for the purification of heavily contaminated hSC cultures. Nevertheless, fibroblast elimination may only be achieved by performing repeated rounds of purification or combining physical (e.g., differential adhesion to plastic) and/or chemical (immunological) methods.
An important consideration is that cell culture plastic panning is a non-cell-type selective method. A proportion of hSCs is usually lost due to retention to the plastic and a proportion of fibroblasts is separated together with the hSC suspensions. If possible, collect the cells that adhere to the non-coated plates and analyze the expression of hSC- and fibroblast-selective markers to determine the efficiency of the separation. The immunopanning technique is easy to implement but may be suboptimal if the percentage of cells that exhibit the expression of the antigen used for panning is low in the cell population. Re-assess and optimize the panning conditions if a new batch of antibodies is used. During the optimization phase, it is important to run negative controls using non-coated plates (i.e., plates without pre-adsorbed antibodies) and plates coated with secondary antibodies only to determine non-specific cell attachment under the selected panning conditions.
Cell labeling and gene delivery
Vital fluorophores. hSC cultures can be stained with CellTrackerTM while adherent (monolayers) or in suspension immediately after enzymatic dissociation. This vital dye rapidly labels all cells except for dead cells with a homogeneous staining throughout the cytoplasm (Figure 4A). The more active the cells are, the higher the staining intensity will be. The hSCs typically stain with similar intensity but differences among treatments or among cell types, e.g., hSCs vs. fibroblasts, are evident. The fluorescent signal is maintained for at least 72 h inside the cells and decays thereafter. Importantly, the cultures can be re-stained without significantly affecting the viability, morphology, or migration of hSCs.
Transfection. The transfection efficiency can vary from prep to prep. Perform gene reporter controls to estimate the efficiency of transfection in each Nucleofection attempt. The pmaxGFP vector expresses GFP from the copepod Potellina sp. and is highly recommended because the fluorescence intensity is very strong and can be observed as soon as 5–6 h post-transfection in a proportion of the hSCs. The transfection efficiency would appear lower if other reporter constructs are used as transfection controls. The hSCs maintain high levels of pmaxGFP expression for at least 2–3 days with evenly distributed, mainly cytoplasmic signal localization. Although the fluorescence intensity declines with time, it is possible to observe pmaxGFP-expressing hSCs at 5–8 days post-transfection. Fading of fluorescence and granular expression of pmaxGFP can be revealed at later time points, possibly due to intracellular removal of excess pmaxGFP protein. The expression levels of other recombinant proteins are variable and likely determined by the characteristics of the protein (e.g., smaller proteins are usually expressed at higher levels), the type of vector used, and possibly environmental factors.
Lentiviral transduction. Follow best practices in handling viral particles to maintain their bioactivity and protect the operators. All operations involving lentiviruses should be performed in a BL2 biosafety cabinet. Infection of human cells with lentiviruses requires BL2+ or enhanced BL2 practices. Follow institutional guidelines to inactivate and dispose of unused viral agents, virally transduced cells, cell culture fluids, and other associated biohazardous products.
Ideally, the optimal MOI will be the minimum one that achieves 100% infection of the target hSCs. Experimentally, the most appropriate MOI may be limited by the levels of associated cytotoxicity. We have observed cytoplasm vacuolization and detachment in hSCs transduced at higher MOIs. Therefore, perform viability tests in the virus-treated conditions. We also recommend monitoring cultures for at least seven days post-infection to determine whether the transgene is toxic to the cells before seeding them for experimentation or preparing cryogenic stocks. If possible, perform the same titration experiment using more than one batch of hSCs to grasp donor-variability and select the best batches for future experiments. Additionally, the infection conditions should be re-tested in the following cases: (1) when a new stock of virus or hSC culture is used; (2) when the viral particles have been stored in the -80 °C freezer for >6 months; and (3) when the viral particles have been subjected to freezing-thawing.
Fluorescent proteins are useful gene reporters for direct image analysis of transduced cells. EGFP and mCherry are generally well tolerated by hSCs, but an excess of protein may be detrimental to cell physiology. For instance, the transduced cells may be more susceptible to die than wild-type cells upon changing environmental conditions (e.g., serum starvation) or manipulations, such as cryogenic storage. The expression of fluorescent proteins in cells infected at an appropriate MOI is not expected to change the proliferation, differentiation, and SC–axon interactions typical of non-infected SCs (Monje et al., 2018). Confirm the transgene expression by an appropriate method for those viral vectors that do not encode for fluorescent proteins.
Antibiotic selection can be used to enrich the number of transduced cells. For this, perform preliminary experiments (killing curve) to determine the minimum concentration of antibiotic that kills all non-infected cells. The optimal antibiotic concentration should be optimized in each cell preparation due to batch variability.
Acknowledgments
Natalia Andersen, Ketty Bacallao, and Kaiwen Peng provided expert technical assistance. Gabriela Aparicio assisted with data analysis and figure preparation, Lingxiao Deng with lab support, Valeria Nogueira with illustrations, Thomas Dolan with video editing, and Louise Pay with English editing. Patrick M. Wood and Mary Bunge contributed guidance and critical materials, including hybridoma cell lines and human Schwann cells, during our early investigations. The protocols described here were developed during a >15-year period supported by funding (to P.V.M.) from the National Institutes of Health NIH-NINDS (NS084326), the Craig H Neilsen Foundation, The Miami Project to Cure Paralysis and the Buoniconti Foundation from University of Miami, and the Indiana State Department of Health (grants 33997 and 43547). Grant NIH/NS009923 (to M.B., P.M.W., and P.V.M.) provided seminal funding for cell labeling and banking studies. P.V.M. is currently affiliated to and receives support from the Department of Neurosurgery from UK. The contents of this article are the responsibility of the author and do not necessarily represent the official views of the funding agencies. We are greatly indebted to the generosity of the anonymous individuals who donated their tissues for research.
Competing interests
P.V.M. is the founder of GliaBio LLC, a consulting company that focuses on glial cell research. The author declares that the research described here was conducted in the absence of commercial or financial relationships that could be construed as a potential conflict of interest.
Ethical considerations
The experiments were performed with human Schwann cell cultures (anonymized, from non-pathological, post-mortem tissues) without regard to the gender or age of the donor. This research was deemed to constitute non-human subjects research by the Institutional Human Subjects Research Offices from University of Miami (years 2003–2018) and Indiana University (years 2019–2022). Procedures were further reviewed and approved by the Institutional Biosafety Committee of Indiana University. The original cell cultures used to optimize our expansion, purification, labeling, and cryopreservation methods were derived from nerve biospecimens made available by the Life Alliance Organ Recovery Agency (LAORA) of the University of Miami Miller School of Medicine to the laboratory of Patrick Wood (University of Miami). Requests for information and materials should be directed to P.V.M. Investigators are encouraged to provide feedback on the reported protocols.
References
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Lee, P. R., Cohen, J. E., Tendi, E. A., Farrer, R., Vries, D. E. G. H., Becker, K. G. and Fields, R. D. (2004). Transcriptional profiling in an MPNST-derived cell line and normal human Schwann cells. Neuron Glia Biol 1(2): 135-147.
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Cell Biology > Cell isolation and culture > Cryopreservation
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A Microfluidic Platform for Screening Gene Expression Dynamics across Yeast Strain Libraries
ES Elizabeth Stasiowski *
RO Richard O’Laughlin *
SH Shayna Holness
NC Nicholas Csicsery
JH Jeff Hasty
NH Nan Hao
(*contributed equally to this work)
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4883 Views: 474
Reviewed by: Chiara AmbrogioNona Farbehi Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Oct 2022
Abstract
The relative ease of genetic manipulation in S. cerevisiae is one of its greatest strengths as a model eukaryotic organism. Researchers have leveraged this quality of the budding yeast to study the effects of a variety of genetic perturbations, such as deletion or overexpression, in a high-throughput manner. This has been accomplished by producing a number of strain libraries that can contain hundreds or even thousands of distinct yeast strains with unique genetic alterations. While these strategies have led to enormous increases in our understanding of the functions and roles that genes play within cells, the techniques used to screen genetically modified libraries of yeast strains typically rely on plate or sequencing-based assays that make it difficult to analyze gene expression changes over time. Microfluidic devices, combined with fluorescence microscopy, can allow gene expression dynamics of different strains to be captured in a continuous culture environment; however, these approaches often have significantly lower throughput compared to traditional techniques. To address these limitations, we have developed a microfluidic platform that uses an array pinning robot to allow for up to 48 different yeast strains to be transferred onto a single device. Here, we detail a validated methodology for constructing and setting up this microfluidic device, starting with the photolithography steps for constructing the wafer, then the soft lithography steps for making polydimethylsiloxane (PDMS) microfluidic devices, and finally the robotic arraying of strains onto the device for experiments. We have applied this device for dynamic screens of a protein aggregation library; however, this methodology has the potential to enable complex and dynamic screens of yeast libraries for a wide range of applications.
Key features
• Major steps of this protocol require access to specialized equipment (i.e., microfabrication tools typically found in a cleanroom facility and an array pinning robot).
• Construction of microfluidic devices with multiple different feature heights using photolithography and soft lithography with PDMS.
• Robotic spotting of up to 48 different yeast strains onto microfluidic devices.
Keywords: Yeast library Protein aggregation Saccharomyces cerevisiae Yeast Microfluidics Microfabrication High-throughput screening Gene expression dynamics
Background
Numerous strain libraries of Saccharomyces cerevisiae have been created, including those with genome-wide GFP tagging of open reading frames (Huh et al., 2003), single-gene deletions (Shoemaker et al., 1996), multi-gene deletions (Tong et al., 2001; Kuzmin et al., 2018), and inducible gene expression (Arita et al., 2021). High-throughput screens using these libraries have led to significant advancements in our understanding of genetic function and regulation in budding yeast (Costanzo et al., 2010; Giaever and Nislow, 2014). Recently, Newby et al. constructed a library of strains called yTRAP (yeast transcriptional reporting of aggregating proteins) in which strains have been engineered to express genetic sensors that report on the protein aggregation status of a gene (Newby et al., 2017). This library includes a set of 158 yTRAP sensors for RNA binding proteins as well as several other sensors for different prion proteins such as Sup35, Rnq1, and New1 (Newby et al., 2017). Since protein aggregation as a result of dysregulated proteostasis has been linked to cellular aging in a number of organisms (Lopez-Otin et al., 2013; 2023), we sought to use this library to identify proteins that may be prone to aggregation during yeast replicative aging.
Experiments in microfluidic devices have revealed that single cells proceed along one of two paths during aging, where one path (termed Mode 1 aging) exhibits loss of rDNA chromatin silencing and the other exhibits diminished mitochondrial function and reduced heme levels (termed Mode 2 aging) (Li et al., 2017; Jin et al., 2019; Li et al., 2020; O’Laughlin et al., 2020). Furthermore, we observed a decline in proteostasis in Mode 1 aging cells that coincides with losses of Sir2 activity and rDNA silencing (Paxman et al., 2022). Many RNA-binding proteins contain low complexity domains and are hence aggregation prone. We reasoned that aging or loss of Sir2 activity may lead to aggregation of RNA-binding proteins and hence interfere with their functions. To test this, we developed a high-throughput microfluidic device to screen for RNA-binding proteins that aggregate in response to a sustained loss of Sir2 activity caused by a nicotinamide (NAM) treatment, which induces rDNA silencing loss and mimics the later phases of Mode 1 aging. We used the device to simultaneously monitor the response of multiple yTRAP library strains to NAM treatment and to assess their aggregation state via fluorescence microscopy. Building off of previous high-throughput microfluidic technology developed by our group for screening bacterial libraries of more than 2,000 strains, termed Dynomics (Graham et al., 2020), we adapted this platform for use in yeast by utilizing design principles our group had previously used to trap single yeast cells (Li et al., 2017; Baumgartner et al., 2018). This protocol describes the steps for building and applying a yeast Dynomics microfluidic device that can continuously culture 48 different yeast strains. We used this device to perform a screen of the yTRAP library to identify RNA-binding proteins that aggregate in response to a loss of Sir2 activity (Paxman et al., 2022), however, this microfluidic platform can be used to perform a variety of dynamic screens on different kinds of yeast libraries.
Materials and reagents
Materials
SU-8 2002 (Kayaku Advanced Materials, catalog number: Y111029)
SU-8 2005 (Kayaku Advanced Materials, catalog number: Y111045)
SU-8 2007 (Kayaku Advanced Materials, catalog number: Y111053)
SU-8 2075 (Kayaku Advanced Materials, catalog number: Y111074)
SU-8 Developer (Kayaku Advanced Materials, catalog number: Y020100)
Edge Bead Remover (EBR) PG (Kayaku Advanced Materials, catalog number: G050200)
Dow Corning Sylgard 184 kit (PDMS) (Dow, catalog number: 2646340 or Fisher Scientific, catalog number: NC9285739)
0.5 mm biopsy puncher (e.g., World Precision Instrument Reusable Biopsy Punch, catalog number: NC0815069)
Scotch tape (3M, catalog number: 50-190-9521)
100 mm (4 inch) Silicon wafers (University Wafer, catalog number: 452)
5 inch × 5 inch × inch glass plate (e.g., McMaster-Carr, catalog number: 8476K15)
1 inch by 3 inch glass slide (e.g., Ted Pella, catalog number: 26007)
Plus Plates (Singer Instruments, catalog number: PLU-003)
Singer RePads, 96 Short (Singer Instruments, catalog number: REP-002)
Singer RePads, 1536 Short (Singer Instruments, catalog number: REP-005)
Singer RePads, 6144 Short (Singer Instruments, catalog number: REP-006)
SBS-format Acrylic Tool (custom made)
Wafer tweezers (e.g., EMS Rubis Style 39S-4, Fisher Scientific, catalog number: 50-239-33)
Weigh boats (e.g., Fisherbrand, Fisher Scientific, catalog number: S67090A)
96 well plates, non-tissue culture treated (e.g., Corning, Fisher Scientific, catalog number: 08-772-53)
Reagents
Tween 20 (e.g., Calbiochem, Millipore Sigma, catalog number: 655204-100 mL)
Yeast extract (BD Bacto, catalog number: 21270)
Peptone (BD Bacto, catalog number: 211820)
Glucose (e.g., Fisher Scientific, catalog number: D16-500)
Agar (e.g., Teknova, catalog number: A7774)
SC Powder (Sunrise Science Products, catalog number: 1300-030)
Yeast nitrogen base (YNB) (BD Difco, Fisher Scientific, catalog number: DF0919-07-3)
10 N NaOH (Fisher Scientific, catalog number: SS255-1)
YNB without riboflavin nor folic acid (Sunrise Science Products, catalog number: 1535-250)
Helmanex III (Fisher brand, Fisher Scientific, catalog number: 14-385-864)
Sulforhodamine B (Invitrogen, catalog number: S1307)
Solutions
Standard YPD media (see Recipes)
Standard SC agar (see Recipes)
Microfluidic media (1 L)
2% Helmanex III solution (see Recipes)
Recipes
Standard YPD media (1 L)
10 g of yeast extract
20 g of peptone
20 g of glucose
Standard SC agar (1 L)
6.7 g of yeast nitrogen base
2 g of SC powder
900 mL of deionized water
Adjust pH to 5.6 using 10 N NaOH and a pH meter
Add 16 g of agar and autoclave at 121 °C for 20 min
Add 20 g of glucose when media has cooled to approximately 50 °C
Microfluidic media (1 L)
6.7 g of YNB without riboflavin nor folic acid
2 g of SC powder
20 g of glucose
0.5 mL of Tween 20
2% Helmanex III solution
98 mL of deionized water
2 mL of Helmanex solution
Equipment
Mask Aligner (Optical Associates Inc (OAI) Hybralign Series 200, MDL: 204-092997)
Hot plate (e.g., Corning Hot Plate, model: PC-600 D, catalog number: 07-770-109)
Profilometer (e.g., Dektak, model: DektakXT)
Sonicator (e.g., Branson 8510, model: 8510R-DTH)
Oven (e.g., Fisher Scientific, model: 6901)
Vacuum desiccator (see Ferry et al. (2011) for vacuum system setup and parts)
Singer ROTOR (Singer Instruments, model: ROTOR)
Singer Stinger (Singer Instruments)
Oxygen Plasma machine (Glow Research AutoGlow plasma system)
pH Meter (e.g., Mettler-Toledo AG, SevenCompact pH/Ion S220, catalog number: 01-915-101)
Inverted microscope (e.g., Nikon Ti-1 model)
Software
Nikon NIS-Elements (catalog number: MQS31000)
Fiji/ImageJ (version 2.1.0/1.53c or another version)
RStudio (version 1.3.1093)
Procedure
Photolithography for wafer fabrication
See Figure 1 for the device layout and design. The silicon wafer was fabricated using standard photolithography techniques previously described by our group (Ferry et al., 2011). We used standard spin protocols from the SU-8 manufacturer and all layers were spun at the spin speeds listed below for 30 s after an initial 10 s photoresist spreading step. The conduit layer was fabricated using 2002 SU-8 photoresist with a spin speed of 1,000 rpm and had a resulting height of 2.4 μms, the spotting region layer was fabricated using 2005 SU-8 photoresist with a spin speed of 2,600 rpm and had a resulting height of 4.8–5 μms, the HD biopixel layer was fabricated using 2007 SU-8 photoresist with a spin speed of 1,000 rpm and had a resulting height of 12 μms, the minor channel layer was fabricated using 2075 SU-8 photoresist with a spin speed of 4,000 rpm and had a resulting height of 60–65 μms, and the major channel layer was fabricated using 2075 SU-8 photoresist with a spin speed of 2,400 rpm on top of an undeveloped major channel layer including a 1 h edge bead removal step (Lee et al., 2011) at 40 °C on a hot plate using Edge Bead Remover (EBR) PG, resulting in a final height of 145–170 μms. For additional details related to multi-layer photolithography, refer to the accompanying Bio-protocol manuscript for Paxman et al., “Fabrication of Microfluidic Devices for Monitoring Yeast Aging” (O’Laughlin et al., 2023).
Figure 1. 48 Strain yeast microfluidic chip. A. Overall design of the 48-yeast-strain Dynomics microfluidic chip. B. Close-up of single-strain culturing units. The regions robotically spotted by the robot are the green regions (~5 μm tall), the cell trapping areas are the cyan regions (~12 μm tall), the dark-blue areas are the conduits (~2.4 μm tall), the dark-gray features are the minor channels (~60–65 μm tall), and the light-gray features are the major channels (~145–170 μm tall).
Soft lithography for fabrication of polydimethylsiloxane (PDMS) devices
Dispense 70 g of Dow Corning Sylgard 184 elastomer base into a weigh boat and mix with 7 g of Dow Corning Sylgard 184 curing agent. Thoroughly mix together using a clean glass stir rod (approximately 3–5 min).
Place the PDMS mixture in a vacuum desiccator to remove bubbles. The vacuum chamber can be periodically vented to facilitate this process and ensure no overflow of PDMS outside the weigh boat. Remove the mixture from the chamber only when all bubbles have been removed.
Cut out a 16 inch ×16 inch piece of aluminum foil and fold in half twice to produce an 8 inch × 8 inch piece. Place the 5 inch × 5 inch × ⅛ inch glass plate in the middle of the aluminum foil and gently fold up the edges, ensuring that the foil does not tear, to create a bowl or dish around the plate. Overlap the aluminum foil over the edges of the glass to minimize PDMS leaking underneath the glass dish.
Place the photopatterned silicon wafer in the center of the glass plate with surrounding aluminum foil walls and pour on the bubble-free PDMS mixture.
Degas the PDMS again by placing the wafer in a vacuum desiccator, ensuring that it is level. Remove only once all bubbles have been removed (approximately 30 min–1 h) (Figure 2A).
Critical step: It is vital that the vacuum desiccator is completely level so that a flat PDMS device is produced. This is a requirement for uniform cell spotting in subsequent steps.
Figure 2. Soft lithography with polydimethylsiloxane (PDMS) for making microfluidic chips. A. PDMS is poured onto the silicon wafer in an aluminum foil boat. B. The wafer is centered using pipette tips. C. After baking, aluminum foil is removed from the wafer edges first. D. Aluminum foil fully removed from wafer. E. Removing PDMS around the wafer. F. Using a razor blade to detach PDMS from the front surface of the wafer. G. Removing PDMS from the back of the wafer. H. Peeling PDMS off of the wafer. I. Cutting out individual chips from the PDMS. J. Punching the cell loading and waste ports from the PDMS. K. Cleaning PDMS chips with 70% ethanol. L. Cleaning chips with Scotch tape.
After all bubbles are cleared, remove the wafer from the vacuum chamber. If needed, use two pipette tips to re-center the wafer. Carefully push down on opposite sides of the wafer to ensure it is in even contact with the glass plate and to push out any PDMS that may have seeped underneath the wafer (Figure 2B).
Place the wafer stack into a level oven and bake at 95 °C for 1 h.
Critical step: The oven must be level in order to obtain a flat PDMS device. Note that PDMS shrinks slightly during baking, and 95 °C must be used as the cell traps are designed to accommodate the amount of shrinkage at this temperature.
Peel off the aluminum foil from the wafer stack and remove any excess PDMS around the wafer using a razor blade (Figure 2C–2E).
Gently slide a razor blade horizontally between the wafer and glass plate and then remove it. Repeat this around the circumference of the wafer until the wafer separates from the glass plate (Figure 2F).
Critical step: The razor blade must slide horizontally between the glass and the wafer. Wafers are extremely fragile; if the razor blade is angled, then the wafer will break.
Using a razor blade, remove any excess PDMS from the bottom of the wafer (Figure 2G). Peel the PDMS off of the feature side of the wafer in the direction of the major channels (Figure 2H).
Place the PDMS on a cutting mat with the feature-side up to keep the PDMS clean. Using a razor blade, cut out each PDMS device (Figure 2I). Punch out the inlet and outlet channels with an 0.5 mm biopsy puncher (Figure 2J).
Rinse devices with 70% ethanol and blow dry with compressed air or pressurized nitrogen gas (Figure 2K).
Use scotch tape to clean devices. Clean the feature side four times and the non-feature side twice. Use forceps to gently press the tape into the features to remove all debris (Figure 2L). Leave a fresh piece of tape on each side to keep the devices clean or store them for later use. Note that PDMS devices can be made well in advance of experiments and stored in this condition; however, it is best practice to use PDMS that is no more than two months old.
Cleaning glass slides
Sonicate glass slides in a 2% Helmanex III solution for 30 min at 40 °C.
Rinse glass slides with deionized water, rubbing them with a clean nitrile glove.
Completely dry the glass slides with pressurized nitrogen gas and ensure that no streaks are visible.
Pause point: Clean glass slides can be stored in a clean, dust-free Petri dish until used.
Cell preparation and device loading using singer ROTOR agar plate preparation
Dispense 42 mL of SC agar into Singer Plus Plates on a level surface.
Critical step: The plates must be flat to ensure full cell transfer to plates and microfluidic devices.
Allow plates to dry on the benchtop with the lids covered for 48 h before adding Parafilm and putting in a 4 °C fridge. Plates can be stored at 4 °C for up to six months.
Cell preparation (3–4 days)
Fill each well of a 96-well plate (e.g., 200–300 μL) with a strain from the yTRAP library and allow cells to grow in Standard YPD media overnight in a 30 °C shaker.
Using the Singer ROTOR, spot the liquid plate onto an SC agar Plus Plate. Use the default pinning settings for both the source and target plates. Allow yeast strains to grow up on the agar plates for two days at 30 °C.
Note: The Singer robot is compatible with plates of 96, 384, 1536, and 6144-density. The 6144 density has a 1.125 mm spacing between arrays, matching the design of our microfluidic device. While the Singer robot can array directly from 96 to 6144 density, the number of cells deposited is variable, and 6144 plates cannot be stored in the fridge for reuse. Therefore, we array strains from a 96-density to 4×1536 density plates that are combined to the microfluidic-matching 6144-density. The number of strains placed onto the 1536-well plates is dependent on the size of the microfluidic device. In this context, we array 12 strains per each 1536 plate including replicates. Furthermore, we array up to eight devices per plate. See Figure 3 for a summary of the arraying procedure.
Figure 3. Overview of robotic spotting process. A. Plates are spotted using the Singer ROTOR array pinning robot. B. Individual pins allow strains to be arrayed in rectangular grids on agar plates. C. Arraying process for the Dynomics microfluidic device.
Using the Singer Stinger single colony picker, re-array the 96-agar plate onto a set of four 1536-density SC agar plates that matches the array of the microfluidic device(s).
Grow cells overnight at room temperature.
Pause point: 1536-density plates can be stored in the fridge and continually used as source plates for up to six months.
6144-density plate and acrylic tool preparation
Using the Singer ROTOR, replicate the four 1536-density agar source plates onto four 1536-density SC agar source plates using the Replicate program. Grow the plates at 37 °C overnight.
Critical Step: S. cerevisiae must be heat-shocked overnight at 37 °C in order to be viable when revived on the microfluidic device.
Using the Singer ROTOR, combine the four 1536-density agar source plates onto one 6144-density SC agar plate using the 1:4 array program and the pinning settings listed in Table 1.
Table 1. Arraying parameters for spotting yeast
Pinning pressure (%) Pinning Speed (mm/s) Pinning overshoot (mm)
Source Target Source Target Source Target
1536 agar to 6144 agar 58 64 19 10 2 1
6144 agar to acrylic 50 100 10 10 0.6 0.6
6144 agar to microfluidic device for S. cerevisiae 50 64 10 10 0.6 0.6
Using the Replicate program on the Singer ROTOR and the pinning setting listed in Table 1, spot cells from 6144-density target plate onto the clean acrylic alignment tool. These cells will be used as alignment markers for the PDMS device.
Aligning the PDMS to the acrylic tool
Using a mask aligner, set the acrylic tool on top of the mask holder with the alignment cells facing up. Bring the cells of one device into focus on the optics.
Remove the scotch tape from one PDMS device, avoiding touching the feature side of the device.
Gently place the PDMS device on top of the alignment cells, feature-side up, such that the center of the spotting regions is centered over the cells.
Place tape on top of the PDMS, pressing the PDMS down to ensure adhesion of the PDMS onto the acrylic tool.
Repeat this process for each device on the acrylic tool.
Oxygen plasma exposure
Expose the clean 1 inch × 3 inch glass slide and the PDMS acrylic stack to 30 W of oxygen plasma for 30 s.
Blow any dust off the glass slide and PDMS acrylic stack with compressed nitrogen.
Cell preparation for S. cerevisiae spotting
Using the Singer ROTOR, combine the four heat-shocked 1536-density SC agar source plates onto one 6144-density SC agar plate using the 1:4 Array program and the pinning settings listed in Table 1.
Critical Point: This plate should be used immediately to spot cells onto the oxygen plasma–exposed PDMS device.
Loading and bonding the device
Using the Singer ROTOR and the parameters listed in Table 1, spot the cells from the 6144-density SC agar plate to the oxygen plasma–exposed PDMS acrylic stack.
Peel the spotted PDMS off the acrylic piece and gently place it face down on the center of the oxygen plasma–exposed glass slide. Gently tap the top of the PDMS, ensuring that the device bonds to the glass.
Incubate the device at 37 °C for at least two hours.
See Figure 4 for information on how spotted cells load into the cell trapping chambers that are used for experimental analysis.
Critical Point: Spotted chips should be wetted with media and set up on the same day to ensure complete cell revival.
Figure 4. Loading of robotically spotted cells into downstream traps. A. Each of the 48 spotted strains resided in a unique bulb-shaped region of the device (gray region). B. During experiments, as cells grow up in the bulb region, they flow into the four downstream traps, which we refer to as hydrodynamic biopixel traps. Cells in these traps are used for experimental analysis. Red arrows indicate the direction of media flow. C. Filling of downstream biopixel traps during experiments. Scale bar = 100 μm.
Setting up the microfluidic experiment
Place the spotted chip into the vacuum desiccator for 20 min.
Wet the chip by adding at least 20 µL of microfluidic media to both the inlet and outlet port.
Note: Additional details for setting up microfluidic experiments from this point forward are explained at length in (Ferry et al., 2011). Further, additional details about the experimental and imaging parameters used for the yTRAP library screen are given in Paxman et al. (2022).
Data analysis
Example images of yTRAP strains growing in the device and responding to nicotinamide (NAM) exposure are shown in Figure 5. For analysis, background subtraction using a rolling ball radius of 50 pixels was done in ImageJ on the GFP channel, which contained the fluorescence signal of each yTRAP strain. Using the phase channel as a guide, biopixels were manually outlined using the polygon tool in ImageJ and used to create regions of interest (ROIs). These ROIs were then used to extract fluorescence information from the corresponding biopixel in the GFP channel (Figure 6A). Fluorescence signals from all four biopixels were averaged together to create a response curve for each strain (Figure 6B).
Figure 5. Example response of the SCP160 strain to nicotinamide (NAM). A. Strain fluorescence before NAM exposure. B. Strain fluorescence during NAM exposure. Sulforhodamine B dye was added to the NAM-containing media (1:10,000 ratio) to aid in visualization. C. Strain fluorescence after NAM exposure. Scale bar = 100 μm.
Figure 6. Analysis of yTRAP strain responses to nicotinamide (NAM). A. Extraction of the fluorescence signals in the GFP channel using manually drawn regions of interest in the corresponding frame in the Phase channel. The NOP15 yTRAP strain is used as an example. Scale bar = 25 μm. B. Mean NOP15 yTRAP GFP signal across all four biopixels. Light-blue shaded area represents the standard deviation. Gray region of the plot represents the NAM exposure window.
To determine yTRAP strains that exhibited aggregation during the NAM induction, the average response curves for each strain were normalized between 0 and 1. Normalized traces were smoothed using the loess() function in R, with the “span” parameter set to 0.2. Using these normalized traces, we then approximated the first derivative for each fluorescence trace by calculating the forward difference. We then filtered the dataset to only include those strains that displayed both a decreasing fluorescence signal for more than half of the NAM induction as well as at least a 75% decrease in fluorescence during the induction (as measured relative to the average fluorescence value of that strain 3 h before the NAM induction). From this list, images were manually verified to ensure decreases in fluorescence were not affected by cell washout from the traps or similar artifacts. This produced a list of 15 strains for further analysis.
Acknowledgments
Figure 2 as well as the text for protocol steps for this manuscript was originally written as part of and appears in E.S.’s PhD thesis “Multiplexed Microfluidics Utilizing Genome-scale Dynamics for Biosensing and Fermentation Monitoring” (Stasiowski, 2019). This work was supported by NIH R01 AG068112 (to NH), AG056440 (to NH, JH), GM144595 (to NH, JH), and GM111458 (to NH).
This protocol was derived from the original work of Paxman et al. (2022).
Competing interests
The authors declare that no competing interests exist.
References
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Article Information
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4,884 | https://bio-protocol.org/en/bpdetail?id=4884&type=0 | # Bio-Protocol Content
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This is an update notice. See the updated protocol.
Peer-reviewed
Update Notice: Computational Analysis of Plasma Lipidomics from Mice Fed Standard Chow and Ketogenic Diet
AS Amy L. Seufert *
JH James W. Hickman *
JC Jaewoo Choi
BN Brooke A Napier
(*contributed equally to this work)
Published: Oct 20, 2023
DOI: 10.21769/BioProtoc.4884 Views: 229
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After official publication in Bio-protocol (https://bio-protocol.org/e4819), we would like to add the following source of funding to the Acknowledgments section of the article: “The QTOF mass spectrometer was funded by an instrument grant from the NIH: ABSciex Triple ToF 5600 NIH #1S10RR027878-01.”
Article Information
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Efficient Differentiation of Human Induced Pluripotent Stem Cell (hiPSC)-Derived Mesenchymal Progenitors Into Adipocytes and Osteoblasts
MD Martha Elena Diaz-Hernandez
NK Nazir M. Khan
HD Hicham Drissi
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4885 Views: 667
Reviewed by: Chiara AmbrogioHaneen Abdulkarim NurVinit SharmaDemosthenis Chronis
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Original Research Article:
The authors used this protocol in eLIFE Jan 2023
Abstract
Human induced pluripotent stem cells (hiPSCs) hold immense promise in regenerative medicine as they can differentiate into various cell lineages, including adipocytes, osteoblasts, and chondrocytes. Precisely guiding hiPSC-derived mesenchymal progenitor cells (iMSCs) towards specific differentiation pathways is crucial for harnessing their therapeutic potential in tissue engineering, disease modeling, and regenerative therapies. To achieve this, we present a comprehensive and reproducible protocol for effectively differentiating iMSCs into adipocytes and osteoblasts. The differentiation process entails culturing iMSCs in tailored media supplemented with specific growth factors, which act as cues to initiate adipogenic or osteogenic commitment. Our protocol provides step-by-step guidelines for achieving adipocyte and osteoblast differentiation, ensuring the generation of mature and functional cells. To validate the success of differentiation, key assessment criteria are employed. For adipogenesis, the presence of characteristic lipid droplets within the iMSC-derived cells is considered indicative of successful differentiation. Meanwhile, Alizarin Red staining serves as a marker for the osteogenic differentiation, confirming the formation of mineralized nodules. Importantly, the described method stands out due to its simplicity, eliminating the need for specialized equipment, expensive materials, or complex reagents. Its ease of implementation offers an attractive advantage for researchers seeking robust and cost-effective approaches to derive adipocytes and osteoblasts from iMSCs. Overall, this protocol establishes a foundation for exploring the therapeutic potential of hiPSC-derived cells and advancing the field of regenerative medicine.
Key features
• iMSC derivation in this protocol uses embryonic body formation technique.
• Adipogenesis and osteogenesis protocols were optimized for human iPSC-derived iMSCs.
• Derivation of iMSC from hiPSC was developed in a feeder-free culture condition.
• This protocol does not include human iPSC reprogramming strategies.
Graphical overview
Schematic representation of induced pluripotent stem cell (iPSC) differentiation into adipocytes and osteoblasts via mesenchymal progenitors as intermediates
Keywords: Osteogenesis Adipogenesis hiPSC Cell derivation Mesenchymal progenitors Trilineage differentiation Regenerative medicine
Background
Mesenchymal stromal/stem cells (MSCs) hold immense promise in regenerative medicine as they possess the capability to differentiate into mature specialized cells. Their therapeutic utility is underscored by their replenishment property in adult tissues, although inherent variations in MSC potency due to donor age, tissue selection, and isolation methods necessitate further improvement for its efficient use as a therapeutic treatment. The International Society for Cell Therapy has proposed minimal criteria to identify MSCs, including adherence to plastic under standard culture conditions and the expression of specific surface markers such as CD105+, CD73+, CD90+, CD34-, CD14-, CD11b-, CD79-, CD19-, or HLA-DR (Dominici et al., 2006; Uder et al., 2018). Recently, adult MSCs have gained attention for their immunoregulatory properties, intrinsic regenerative capacity, and multilineage potential (Hoang et al., 2022; Kim et al., 2022). Although MSC derivation poses an intrinsic therapeutic utility, heterogeneity derived from donor to donor, tissue isolation, collection, and expansion methods need to be improved for their efficient use as a therapeutic treatment (Hwang et al., 2009).
Interestingly, the derivation of MSCs from human induced pluripotent stem cells (hiPSCs) offers a viable alternative (iPS-MSCs or iMSCs), presenting an avenue to study activation pathways and differentiation routes during cell commitment and terminal differentiation. Differentiation methodologies for adipocytes, cartilage, and bone have been developed by recapitulating molecular cues present in the embryo, given MSCs’ origin from the mesodermal layer (Hoang et al., 2022; Humphreys et al., 2022). Selective induction of mesodermal cells and specific iMSC subpopulations expressing distinct surface markers has been reported using culture conditions. It has been reported that iMSC subpopulations expressing CD29+, CD44+, CD73+, CD90, and CD105+ CD11b-, CD14-, CD31-, CD45-, and HLA-DR- markers have a faster proliferative capability and restrictive adipogenic ability compared to BM-MSC (Kang et al., 2015).
Here, we present detailed step-by-step protocols for the differentiation of osteoblasts and adipocytes from human iPSCs, starting from a naive pluripotent state and via the derivation of a scalable, mesenchymal-like progenitor intermediate, as described in our previous work (Khan et al., 2023). The protocols include the derivation of iMSCs from hiPSCs through embryoid bodies (EBs) formation, followed by osteoblast and adipocyte differentiation. Various assessment methods are also described to validate the maturation of functional cells. The standardized protocols provided in this study offer valuable insights into the differentiation potential of hiPSC-derived iMSCs, contributing to a deeper understanding of their therapeutic applications. By elucidating the intricacies of cell commitment and maturation processes, these methodologies may lead to further improvements in the development of regenerative therapies and disease modeling.
Materials and reagents
Biological materials
HDFa-YK27-hiPSC human dermal fibroblast line (University of Connecticut cell and Genome Engineering Core)
YK27-iPSC–derived iMSCs (Emory Musculoskeletal Center, Emory University)
Reagents
mTeSR-1 media (Stemcell Technologies, catalog number: 85871)
Accutase in DPBS without Ca++ or Mg++ (Innovative Cell Technologies, catalog number: AT-104)
Dulbecco’s Modified Eagle medium (DMEM) (1×) (Gibco, catalog number: 11965-092)
HyClone fetal bovine serum (FBS) defined, heat-inactivated (Cytiva, catalog number: SH30070.03HI)
Bovine serum albumin (BSA) solution 7.5% in DPBS (Sigma-Aldrich, catalog number: A8412-100mL)
Penicillin streptomycin (Pen/Strep) (Gibco, catalog number: 15070-063)
Non-essential amino acid solution (MEM NEAA) (100×) (Gibco, catalog number: 11140-050)
Gelatin 0.1% in water (Stemcell Technologies, catalog number: 07903)
Dulbecco’s Ca2+ Mg2+ free phosphate buffered saline (DPBS) (10×) (Gibco, catalog number: 14080055)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D8418)
StemLine II (Sigma-Aldrich, catalog number: SLCM2573)
Dulbecco’s Ca2+ Mg2+ free phosphate buffered saline (DPBS) (1×) (Gibco, catalog number: 14040117)
Trypsin-EDTA 0.25% (1×) phenol red (Gibco, catalog number: 25200056)
Trypan Blue solution (0.4%) (Gibco, catalog number: 152500-061)
GlutaMAX 200 mM supplement (Gibco, catalog number: 35050061)
Collagen I, rat tail (Gibco, catalog number: 1048301)
2-propanol or Isopropanol (Sigma-Aldrich, catalog number: 67-63-0)
IBMX (3-isobuty-l-methyl-xanthine) (Sigma-Aldrich, catalog number: 28822-58-4)
Alkaline phosphatase (ALP) (Sigma-Aldrich, catalog number: 85L2-1kt)
10× phosphate buffered saline (PBS) (Gibco, catalog number: 14200-075)
bFGF (146 a.a.) (Peprotech, catalog number: 100-18C)
Recombinant human BMP4 (Peprotech, catalog number: 120-05ET)
Dexamethasone (Stem Cell Technologies, catalog number: 72092)
Indomethacin (Stem Cell Technologies, catalog number: 73942)
β glycerol phosphate (Sigma-Aldrich, catalog number: 13408-09-8)
Recombinant human insulin solution (Santa Cruz Biotechnology, catalog number: 11061-68-0)
Oil red stock solution (Sigma-Aldrich, catalog number: O0625)
Recombinant human BMP4 (Peprotech, catalog number: 120-05)
Recombinant Human VEGF (Peprotech, catalog number: 100-20)
Sodium hydroxide (NaOH) (Sigma, catalog number: 484024)
Ascorbic-acid-2-phosphate (AA2P) (Sigma, catalog number: A8960)
Solutions
10 μg/mL of bFGF (see Recipes)
25 μg/mL recombinant human BMP4 (see Recipes)
10 mM dexamethasone solution (see Recipes)
10 mM indomethacin (see Recipes)
20 mM IBMX (see Recipes)
1 N NaOH (see Recipes)
3% Oil red stock solution (see Recipes)
4% paraformaldehyde (PFA) solution (see Recipes)
iMSC maintenance media (see Recipes)
50 mg/mL ascorbic acid (AA2P) (see Recipes)
1 M β glycerol phosphate (see Recipes)
Pre-adipocyte basal media (see Recipes)
iMSC expansion media (see Recipes)
Adipocyte media (see Recipes)
Insulin induction media (see Recipes)
Osteogenic media (see Recipes)
iMSC frozen media (see Recipes)
Collagen I rat tail coated plates (see Recipes)
2% Alizarin Red staining solution (see Recipes)
Recipes
10 μg/mL of bFGF (146 a.a.)
Reagent Final concentration Quantity
5 mM HCl containing 0.1% BSA NA 1 mL
bFGF 10 ng/μL 10 μg
Centrifuge vial prior to opening. Avoid repeated freeze-thaw cycles. Store aliquots at -80 °C.
25 μg/mL recombinant human BMP4
Reagent Final concentration Quantity
5 mM HCl containing 0.1% BSA NA 400 μL
BMP4 25 ng/μL 10 μg
Centrifuge vial prior to opening. Avoid repeated freeze-thaw cycles. Store aliquots at -80 °C.
10 mM dexamethasone solution (F.W: 392.4 g/mol) (stock solution)
Reagent Stock concentration Quantity
DMSO NA 2.54 mL
Dexamethasone 10 mM 10 mg
As a general guide, small molecules in DMSO solution are recommended to be stored in small aliquots at -20 °C. Aliquot the solutions into working volumes to avoid freeze-thaw cycles. Protect from prolonged exposure to light.
10 mM indomethacin (F.W: 357.9 g/mol) (100× stock solution)
Reagent Stock concentration Quantity
DMSO NA 2.79 mL
Indomethacin 10 mM 10 mg
It is recommended to store DMSO aliquots at -20 °C. Aliquot the solutions into working volumes to avoid freeze-thaw cycles. Protect from prolonged exposure to light.
20 mM IBMX (F.W: 222.2 g/mol) (stock 20×)
Reagent Stock concentration Quantity
DMSO NA 22.5 mL
IBMX 20 mM 100 mg
Sterile filter with 0.2 μM pore microfilter. Aliquots are stored at -20 °C.
1 N NaOH (F.W: 40 g/mol)
Reagent Final concentration Quantity
H2O NA 1,000 mL
NaOH 1 N 40 g
3% Oil red stock solution
Reagent Final concentration Quantity
Isopropanol NA 100 mL
3% Oil red NA 3 mg
0.3% Oil red working solution: mix three parts of 3% oil red stock solution and two parts of distilled water and allow to sit for 10 min. Final solution is Whatman paper filtered. This solution is stable for 2 h. 3% Oil red stock solution is stable for one year.
4% paraformaldehyde (PFA) solution
Reagent Final concentration Quantity
1× PBS NA 70 mL
PFA
NaOH
4%
1 M
4 g
1 mL
Dissolve PFA in a heating and stirring block (60 °C) and in a chemical hood. Adjust to 7.4 pH with 1 M HCl. Adjust to 100 mL final volume. Aliquots can be stored at -20 °C.
iMSC maintenance media
Reagent Final concentration Quantity
DMEM
Pen/Strep
MEM NEAA
bFGF 100 μg/mL
NA
1×
1×
5 ng/mL
440 mL
5 mL
5 mL
25 μL
FBS defined 10% 50 mL
50 mg/mL AA2P
Reagent Stock concentration Quantity
Ascorbic-acid-2-phosphate (AA2P)
Milli-Q water
50 mg
1 g
20 mL
To dissolve the AA2P, prewarm the water at 37 °C.
1 M β glycerol phosphate (F.W: 306.11 g/mol)
Reagent Stock concentration Quantity
β-Glycerol phosphate
Milli-Q water
1 M
1 g
3.26 mL
Pre-adipocyte basal media
Reagent Final concentration Quantity
DMEM
FBS defined
GlutaMAX
Pen/strep
NA
5%
2 mM
1×
218 mL
2.25 mL
2.25 mL
2.25 mL
iMSC expansion media
Reagent Final concentration Quantity
StemLine II
FBS defined
Pen/strep
VEGF©
hrBMP4©
NA
10%
1×
25 ng
25 ng
45 mL
5 mL
0.5 mL
vary©
vary©
©Supplement the iMSC media with 25 ng/mL of hrBMP and 50 ng/mL of VEGF only to the media required for media change.
Adipocyte media
Reagent Final concentration Quantity
DMEM NA 218.2 mL
FBS
10 mM Dexamethasone
20 mM IBMX
Insulin solution
10 mM indomethacin
10%
1 μM
0.5 mM
10 μg/mL
100 μM
25 mL
25 μL
6.25 mL
25 μL
2.5 mL
Insulin induction media
Reagent Final concentration Quantity
DMEM
Pen/Strep
Insulin solution
NA
1×
10 μg/mL
225 mL
2.25 mL
22.5 μL
Osteogenic media
Reagent Final concentration Quantity
DMEM
FBS defined
10 mM Dexamethasone
50 mg/mL ascorbic acid
1 M β-glycerophosphate
Pen/Strep
NA
10%
0.1 μM
50 μg/mL
10 mM
1×
43.5 mL
5 mL
0.5 μL
50 μL
0.5 mL
0.5 mL
iMSC frozen media
Reagent Final concentration Quantity
iMSC maintenance media
FBS
DMSO
10%
80%
10%
1 mL
8 mL
1 mL
Collagen I rat tail coated plates
Plates are rinsed twice with 1× PBS before use and kept at room temperature.
Reagent Final concentration Quantity
Collagen 1 rat tail 3 mg/mL
Acetic acid
10× PBS
2 μg/cm2
20 mM
Vary
Vary
Perform manipulation of the collagen on ice (2–8 °C) as gelling may occur rapidly at room temperature.
Calculate the Collagen 1 rat tail volume needed:
20 μg/mL collagen × final volume = stock collagen I rat tail (μg/mL)
Calculate the volume of 20 mM acetic acid needed:
Final volume needed-Volume of Collagen1 stock.
100 μL of the solution is added to each well of the 24-well plate.
Add the solution to the plates. Incubate at room temperature for 1 h. Do not freeze the Collagen 1 rat tail.
2% Alizarin Red staining solution
Reagent Final concentration Quantity
Alizarin Red
Milli-Q water
40 mM
6.846 g
500 mL
Adjust the pH to 4.2 with 1 N NaOH.
Laboratory supplies
15 mL conical centrifuge tubes (Thermo Scientific, catalog number 339650)
50 mL conical centrifuge tubes (Thermo Scientific, catalog number 339652)
6 well-cell culture plate polystyrene (Costar, catalog number: 3506)
96 well-cell culture plate polystyrene (Costar, catalog number: 3595)
24 well-cell culture plate polystyrene (Costar, catalog number: 3524)
Cell culture dish sterile 60 mm ×15 mm (Greiner bio-one, catalog number: 628 160)
EZ Flow syringe filter pore size 0.22 μm (Foxx Life Sciences, catalog number: 378-2415-OEM)
Cell lifter polyethylene (Costar, catalog number: 3008)
2.0 mL cryogenic vials (Corning, catalog number: 430661)
Whatman Grade 1 filter paper (Sigma-Aldrich, catalog number: WHA1001125)
BD 10 mL syringe Luer-Lok tip (BD, catalog number: 309604)
EZFlow® 33 mm STERILE syringe filter, 0.2 µm (Stellar Scientific, catalog number: 379-2415-OEM)
Equipment
Bright-line chamber and 0.4 mm cover glass (Electron Microscopy Sciences, catalog number: 63514-11)
Corning LX cool cell freezing system for cryogenic vials (Corning, catalog number: 432004)
Plate reader (SpectraMax® iD3, Molecular Devices, catalog number: 735-0391)
Tissue culture incubator, 5% CO2 atmosphere (Forma Scientific, model: 3110)
Laminar flow hood (StreilGARD III Advance, The Baker Company, model: SG603)
Light microscope for live imaging (Nikon, model: TMS 215135)
Procedure
MSC derivation from hiPSC using 3D embryonic body formation method
In this protocol, we use the formation of embryonic bodies (EBs) to facilitate the commitment of the hiPSCs (HDFa-YK27-hiPSC) to the mesodermal lineage. Once EBs have formed, iMSC enrichment is achieved by induction with mesodermal factors and later selection of specific MSC population (YK27-iPSC–derived iMSCs).
To achieve proper cell density for iMSC derivation, we strongly recommend using three wells of the 6-well plate or one 6 cm dish of hiPSC culture. Wells and dish(es) must be 80% confluent.
To perform EB formation, dissociation of hiPSCs colonies is achieved by Accutase-PBS diluted (1:1 ratio) treatment. Use 1 mL of this solution (Accutase) for one well of the 6-well plate, whereas 3 mL is needed for a 6 cm dish.
Colonies are partially dissociated at room temperature for 3 min. After 3 min, slow down cell dissociation by adding 1 mL of mTeSR-1 media; then, lift clumps of hiPSCs cells and transfer into a 15 mL conical tube.
Centrifuge tubes at 300× g for 5 min.
Aspirate the supernatant from the tube; the pellet should be visible at the bottom. Then, add 0.5 mL of mTeSR-1 media to each well of a 6-well plate (for each 6 cm dish, add 1 mL of mTeSR-1 media).
Gently resuspend the pellet. Clumps of 2–3 cells together are still expected.
Perform cell counting (see the hiPSC counting method in General notes and troubleshooting).
A centrifugation of 300× g is required to adjust cell number to 15,000/15 μL in mTeSR-1 media. We recommend having 3.0 × 105 cell density as a minimum to obtain 20 EBs.
Dispense 15 μL of cell suspension into a 10 cm Petri dish. The use of a 20 μL pipette tip is preferred for easy cell dispensing. iPSC drops should be at a certain distance from each other to prevent them from fusing together (Figure 1A).
Figure 1. Graphical representation of step-by-step method used for embryonic body (EB) formation. A. Human induced pluripotent stem cells (hiPSCs) are dissociated and seeded in high-density drops in a 10 cm Petri dish. B. The Petri dish is inverted upside down immediately after preparing drops and incubated for 24 h at 37 °C. C. Plates are inverted back and cultured for an additional 24 h to allow the formation of EBs, which are counted manually and transferred to gelatin plates.
Close the lid of the Petri dish and then flip the plate over. Keep the dish inside the incubator for 16 h at inverted position (Figure 1B).
After incubation, flip the Petri dish back and carefully add 8 mL of mTeSR-1 media. Return the plate to the CO2 incubator for an additional 24 h at 37 °C (Figure 1C).
Transfer the EBs into a 6 cm dish coated with 1% gelatin. The use of the 1 mL pipette tip is preferred for easy handling of EBs.
Replace the mTeSR-1 media with iMSC expansion media (see Recipe 14) and supplement it with VEGF and BMP4 factors (see Recipe 2).
EBs have adhered to the plate after 24 h. However, some EBs will still be floating. A mix of MSC precursors should appear in the next days. Maintain the media and factors for a total of three days.
Replace the iMSC expansion media with iMSC maintenance media (see Recipe 10) for the next 5–7 days.
Once the dish is confluent, pass the iMSCs mix population using the Accutase-PBS method, as previously described. At this point, there is no need to dissociate EBs in single-cell suspension. Transfer EB aggregates and floating cells to two dishes coated with gelatin to expand. Label dishes as passage number 0.
iMSCs should be passaged when they reach confluency. From P1, they should be passaged by the trypsinization method (see General notes and troubleshooting). Once they reach P4, perform gene expression and osteo, adipo, and chondrogenesis and cell surface analyses to select specific cell populations, if required.
iMSCs are cryo-preserved for further validation and expansion (See General notes and troubleshooting, Recipes).
Recovery and maintenance of human iMSCs
Transfer iMSCs from cryovials to the hood in dry ice. Cell thawing should be completed in a short time to maximize cell survival. Immerse frozen cryovials in a clean water bath at 37 °C; once the chunk of ice is melted (approximately 1 min), gently transfer the cells to a sterile 15 mL conical tube containing 9 mL of DMEM media.
Centrifuge the tubes at 400× g for 3 min to pellet the cells.
Aspirate the media and resuspend the cells in maintenance media.
For cell culture maintenance, use a 10 cm culture dish without any coating.
Change media every three days until iMSCs have reached 80% of confluency.
iMSC cryopreservation
iMSC frozen vials are made in early passages at 1 × 106/mL cell density.
Prepare the iMSC frozen media (see Recipe 18) in advance and keep at room temperature.
Briefly, remove maintenance media from culture dishes and wash once with 1× PBS.
Perform trypsinization and cell counting (see General notes and troubleshooting).
Adjust cell density to 1 × 106 /mL in iMSC maintenance media.
Centrifuge cell suspension at 400× g for 5 min.
Resuspend pellet in frozen media to have 1 × 106/mL, immediately transfer into labeled cryovials, and place on the Corning LX cool cell freezing system. Keep at -80 °C overnight.
Cryovials are ready for long-term storage at the liquid nitrogen tank.
Preadipocyte growth
To promote adipogenic differentiation, treatment should start once MSCs are in the arrest phase of the cell cycle (Zebisch et al., 2012). Although there is no specific recommendation to the plate size, 24-well plates are preferred for the adipocyte assay and 4,000 cells/cm2 is recommended as initial seeding density. Collagen I from rat tail is used to coat the plates (see Recipe 19). Keep in mind that experimental duplicates or triplicates and control wells (no-treatment group) should be included for all experiments.
Growth and preadipocyte expansion are achieved by using pre-adipocyte basal media (see Recipe 13) with media changes every other day until cells reach 100% confluency.
During preadipocyte growth, iMSCs do not show major changes in morphology. At this stage, iMSCs are seen as fibroblast-like cells. Maintain iMSCs, preadipocytes, and adipocytes in 5% CO2, 95% humidity and 37 °C.
Adipocyte induction
Adipocyte induction is promoted at iMSC and preadipocytes 100% confluency and in intervals/cycle between the adipocyte media (see Recipe 15) for 3–4 days, followed by insulin induction media (see Recipe 16) for an additional three days.
Perform three repeated interval-cycles (adipocyte media: insulin media); pay special attention during media changes to avoid drying and detachment of cells from the plastic surface between media changes. During adipocyte induction, preadipocytes change their morphology to a spherical shape, and lipid droplets inside of the cytoplasm are seen. Multiple induction series can be performed until 95% of the cells are committed to adipogenic lineage.
Changes in iMSC morphology are observed approximately at day 7, and cells become highly responsive to lipogenic and lipolytic hormones and accumulate high levels of lipogenic enzymes (Smith et al., 1988). Monitor terminal differentiation by brightfield microscopy, through changes from an elongated to a round shape and an evident presence of lipid vacuoles, which increase in number and size covering the whole well.
Multiple induction treatments resulted in more than 95% of the cells committing to this lineage, and the lipid vacuoles continued to develop over time, coalesced, and eventually filled the cell. These adipocytes remained healthy in culture for at least three months.
Oil red staining
Once the terminal adipocyte maturation is achieved, stain lipid vacuoles of the adipocytes with 0.3% oil red solution.
Fix adipocytes in 4% PFA for 15 min at room temperature, followed by one wash with 1× PBS.
Remove the PBS, add 250 mL of 60% isopropanol, wait for 30 s, and aspirate the solution. Repeat the 60% isopropanol wash and then let the plate dry completely at room temperature (takes approximately 3 min).
Add 200 μL of 0.3% oil red working solution to each well, including experimental and control groups (non-induced group).
Incubate the plate at room temperature and monitor it under the microscope for red color precipitates, which can be visualized after 15 min of 0.3% of oil red staining (Figure 2).
Figure 2. Differentiation of human induced pluripotent stem cell (hiPSC)-derived mesenchymal progenitor cells (iMSCs) into adipocytes was determined by oil red staining. Accumulation of lipid droplets as shown by the pointed arrows indicate adipocyte differentiation of iMSCs at 21 days of differentiation. Scale bar = 100 μm.
Red precipitates’ area and dots (lipid vesicles) are strong in color compared to the other cell compartment areas and are an indication to stop the reaction (approximately 20 min). After staining, remove the 0.3% oil red staining solution and wash the plate once with distilled water; visualize under a microscope to be sure about proper staining. If staining is not optimal, incubate the plate with 0.3% Oil red solution for some additional time.
After proper staining, remove the 0.3% oil red staining solution and wash the plate with distilled water until the water is colorless.
Visualize the plate under the microscope for visible dark red color oil droplets (Figure 2).
Osteogenic differentiation of iMSCs
Osteogenic differentiation of iMSC can be achieved in monolayer cultures at 4,000 cells/cm2. 6-well plates are preferred for the osteogenic assay, but 12-well plates can also be used. From our experience, coating wells with Collagen I rat tail (see General notes and troubleshooting) decreased cell peeling during the differentiation process.
After plating the iMSCs, culture the cells in MSC maintenance media (see Recipe 10) for 24 h at 37 °C, 5% CO2. As in adipogenesis, the treatment is preferred once MSCs are in cell cycle arrest.
After 24 h of incubation, change MSC media to osteogenic media (see Recipe 17) and maintain for three days.
Change induction media every 3–4 days for a total of 14–21 days.
Maintain osteogenic cultures until changes in cell morphology are visualized under brightfield microscopy (after the third week of osteogenic treatment).
Keep in mind to have experimental groups in duplicate or triplicates. Additionally, an undifferentiated control group (no-treatment group) should be included in the experiments.
Alizarin Red staining
Extracellular calcium deposits stain positive for Alizarin Red solution.
Fix the osteogenic cultures with 4% PFA (see Recipe 9) for 15 min at room temperature.
Wash the wells three time with distilled water for 5 min each.
Add 700 μL of filtered 0.2% Alizarin Red staining solution to each well and incubate for 30 min at room temperature. After 30 min, monitor the staining under the microscope; cells will change from bright orange to red. If required, increase the incubation time for additional staining and periodically monitor cells until a complete red color develops (Figure 3). Keep in mind that undifferentiated control cells appear slightly reddish.
Figure 3. Osteoblast differentiation of human induced pluripotent stem cell (hiPSC)-derived mesenchymal progenitor cells (iMSCs) was determined by Alizarin Red S staining. Microscopic view of staining on day 21 of osteogenic differentiation. A. Undifferentiated control did not show any staining with Alizarin Red. B. Osteogenic media induced a strong red color staining, indicating the differentiation into osteoblast. The mineralization is shown by the arrow pointing towards nodule formation. Scale bar = 500 μm.
After proper staining, remove the staining solution and wash the wells three times for 5 min each with distilled water.
Keep the wells with distilled water for image acquisition.
Alkaline phosphatase staining
Osteoblasts form aggregates or nodules, which show increased expression of alkaline phosphatase. We followed the protocol from Sigma-Aldrich with some modifications:
Culture the cells as described for osteogenesis.
Prepare diazonium salt solution: dissolve one capsule of Fast violet B in 48 mL of pre-warmed water (18–26 °C).
Add 2 mL of Naphthol AS-MX phosphate alkaline solution to the previous diazonium salt solution (ALP staining solution).
Carefully, remove the media from the wells and fix the wells with 4% PFA (see Recipe 9).
Add 1 mL of ALP staining solution to each well and incubate in the dark for 15–30 min at 37 °C.
Rinse the wells two times for 5 min with tap water.
Image using a light microscope.
Mineralization can also be assessed by colorimetric detection at 405 nm.
Data analysis
Colorimetric measurements are done in triplicates using a plate reader at 405 nm.
Present the data as mean ± S.E.M. of at least three independent samples. Perform statistical comparisons between untreated and growth factor–treated groups using a two-tailed Student’s t-test. Significance is assigned to P values < 0.05.
General notes and troubleshooting
General notes
Determination of the cell number:
Cell counting is performed in 100 μL aliquots of cell suspension and diluted in 100 μL of trypan blue staining solution. 10 μL of stained cells is transferred into a Neubauer chamber and cell counting is performed at the 10× objective. Counting is repeated in the second chamber, and cells average is used to calculate total number of cells per milliliter. Cell concentration is adjusted by centrifugation or dilution to obtain the desired cell density for adipogenesis or osteogenesis outcomes.
Determination of the hiPSC cell number:
Cell counting is performed in a 100 μL aliquot of the hiPSCs suspension. A 100 μL aliquot of hiPSC clumps (3–4 cell aggregates) is completely dissociated by pipetting up and down to get a single-cell suspension and then diluted in 100 μL of trypan blue staining solution to a 1:1 dilution. 10 μL of the stained single cells is transferred into a Neubauer chamber, and cell counting is performed at the 10× objective. Counting is repeated in the second chamber, and cells average is used to calculate total number of cells per milliliter. Cell concentration is adjusted by centrifugation or dilution to obtain the desired cell density for EBs formation.
Cell trypsinization:
Briefly, maintenance media is removed from culture dishes and washed once with 1× PBS; then, Trypsin-EDTA solution is added and incubated for 3–5 min at 37 °C. After cell detachment, trypsin is inactivated by adding maintenance media. Cells are transferred to a 50 mL conical tube. iMSCs are centrifugated at 400× g for 5 min and resuspended.
iMSC cryopreservation:
iMSCs frozen vials are made in early passages at 1 × 106 /mL cell density and passage number is documented.
Troubleshooting
Problem 1: EB formation. Embryonic body is disintegrated.
Possible cause: issues with cell density. A high number of cells can prevent cell aggregation. Meanwhile, a low cell number can form small EBs that will take more time to grow, which can affect the number of cells committed to the mesodermal lineage.
Solution: confirm initial cell density. Also, different cell densities can be tested to confirm the adequate cell number required for EBs formation. For iMSC derivation, we strongly recommend using three wells of the 6-well plate or one 6 cm dish for proper cell density. Wells and dish(es) must be 80% confluent.
Problem 2: Recovery and maintenance of human iMSC. Low rate of cell recovery.
Possible cause: iMSCs were harvested at the stationary phase of the cell cycle.
Solution: cells need to be collected in the growing phase to have good cell quality after thawing. Cryopreservation of iMSCs should be done at 80%–90% confluency.
Be sure that iMSCs are completely mixed with the cryopreservation media.
Dispensing of cells needs to be done gently by slowly dropping to avoid cell death excess.
Problem 3: Adipogenesis and osteogenesis peeling off in monolayer cultures.
Possible cause: excessive force during media changes and dryness of the monolayer cells.
Solution: to avoid disturbing the seeded cells, place the plate in a horizontal position during transfer from hood to CO2 incubator.
Aspirate the media from the edges using the manual pipette.
We also recommend leaving some media (just to protect the cells from dryness), e.g., approximately 250 μL for one well of the 6-well plate. Avoid handling the plate in a vertical position.
Acknowledgments
This work was supported by funds from Veteran Affairs and Emory University School of Medicine. This research was funded by Georgia CTSA/REM Pilot Project 00080502 to HD, Veteran Affairs CaReAP Award (I01-BX004878) to HD.
This protocol was mainly adapted from the publication of Khan et al. (2023).
Competing interests
There are no competing interests.
References
Dominici, M., Le Blanc, K., Mueller, I., Slaper-Cortenbach, I., Marini, F., Krause, D., Deans, R., Keating, A., Prockop, D. and Horwitz, E. (2006). Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4): 315–317.
Hoang, D. M., Pham, P. T., Bach, T. Q., Ngo, A. T. L., Nguyen, Q. T., Phan, T. T. K., Nguyen, G. H., Le, P. T. T., Hoang, V. T., Forsyth, N. R., et al. (2022). Stem cell-based therapy for human diseases. Signal Transduct. Target. Ther. 7(1): 272.
Humphreys, P. A., Mancini, F. E., Ferreira, M. J., Woods, S., Ogene, L. and Kimber, S. J. (2022). Developmental principles informing human pluripotent stem cell differentiation to cartilage and bone. Semin. Cell. Dev. 127: 17–36.
Hwang, N. S., Zhang, C., Hwang, Y. and Varghese, S. (2009). Mesenchymal stem cell differentiation and roles in regenerative medicine. WIREs Syst. Biol. Med. 1(1): 97–106.
Kang, R., Zhou, Y., Tan, S., Zhou, G., Aagaard, L., Xie, L., Bünger, C., Bolund, L. and Luo, Y. (2015). Mesenchymal stem cells derived from human induced pluripotent stem cells retain adequate osteogenicity and chondrogenicity but less adipogenicity. Stem. Cell. Res. Ther. 6(1): 144.
Khan, N. M., Diaz-Hernandez, M. E., Chihab, S., Priyadarshani, P., Bhattaram, P., Mortensen, L. J., Guzzo, R. M. and Drissi, H. (2023). Differential chondrogenic differentiation between iPSC-derived from healthy and OA cartilage is associated with changes in epigenetic regulation and metabolic transcriptomic signatures. eLife 12: e83138.
Kim, H., Park, K., Yon, J. M., Kim, S. W., Lee, S. Y., Jeong, I., Jang, J., Lee, S. and Cho, D. W. (2022). Predicting multipotency of human adult stem cells derived from various donors through deep learning. Sci. Rep. 12(1): 21614.
Smith, P. J., Wise, L. S., Berkowitz, R., Wan, C. and Rubin, C. S. (1988). Insulin-like growth factor-I is an essential regulator of the differentiation of 3T3-L1 adipocytes. J. Biol. Chem. 263(19): 9402–9408.
Uder, C., Brückner, S., Winkler, S., Tautenhahn, H. and Christ, B. (2018). Mammalian MSC from selected species: Features and applications. Cytometry Part A 93(1): 32–49.
Zebisch, K., Voigt, V., Wabitsch, M. and Brandsch, M. (2012). Protocol for effective differentiation of 3T3-L1 cells to adipocytes. Anal. Biochem. 425(1): 88–90.
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Simultaneous Profiling of Chromosome Conformation and Gene Expression in Single Cells
YC Yujie Chen *
HX Heming Xu *
ZL Zhiyuan Liu *
DX Dong Xing
(*contributed equally to this work)
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4886 Views: 1000
Reviewed by: Rajesh RanjanVartika Sharma Anonymous reviewer(s)
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Bio-protocol journal peer-reviewed
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Original Research Article:
The authors used this protocol in Science Jun 2023
Abstract
Rapid development in single-cell chromosome conformation capture technologies has provided valuable insights into the importance of spatial genome architecture for gene regulation. However, a long-standing technical gap remains in the simultaneous characterization of three-dimensional genomes and transcriptomes in the same cell. We have described an assay named Hi-C and RNA-seq employed simultaneously (HiRES), which integrates in situ reverse transcription and chromosome conformation capture (3C) for the parallel analysis of chromatin organization and gene expression. Here, we provide a detailed implementation of the assay, using mouse embryos and cerebral cortices as examples. The versatility of this method extends beyond these two samples, with the potential to be used in various other cell types.
Key features
• A multi-omics sequencing approach to profile 3D genome structure and gene expression simultaneously in single cells.
• Compatible with animal tissues.
• One-tube amplification of both DNA and RNA components.
• Requires three days to complete.
Graphical overview
Schematic illustration for the Hi-C and RNA-seq employed simultaneously (HiRES) workflow
Keywords: Single-cell sequencing Hi-C RNA-seq Multi-omics Three-dimensional genome Transcriptome
Background
The acquisition of distinct cellular identities in multicellular organisms during development requires precise regulation of gene expression across various levels. The progress of single-cell multi-omics sequencing technologies has provided new ways for the comprehensive profiling of the transcriptome in conjunction with various other features, including DNA methylation, cellular proteins, chromatin accessibility, and histone modifications [1–4]. Emerging evidence has suggested that transcription is also closely related with higher-order spatial organization of chromatin in the nucleus [5–7]. Nevertheless, there is a lack of techniques allowing for simultaneous profiling of chromatin conformation and gene expression in individual cells. We have recently developed a method named Hi-C and RNA-seq employed simultaneously (HiRES) [8]. Here, we provide a step-by-step guide on how to perform the assay on mouse embryos and brains. This assay can also be performed on other cell lines or tissues from which single-cell suspensions can be obtained.
Materials and reagents
Materials
Tools and supplies for animal dissection (e.g., forceps, tweezers, eye scissors)
DNA low-bind disposable tips (Axygen, catalog numbers: TF-1000-L-R-S, TF-200-L-R-S, TF-20-L-R-S, TF-300-L-R-S)
Multi-channel pipette tips (Rainin, catalog numbers: 30374581, 30374584)
DNA LoBind tube 1.5 mL (Eppendorf, catalog number: 022431021)
0.5 mL thin-wall PCR tubes (Axygen, catalog number: PCR-05-C)
Thermo-fast 96-well plates (Thermo Fisher, catalog number: AB-1400-L)
40 μm cell strainer (Falcon, catalog number: 352340)
0.22 μm filter unit, sterile (Millipore, catalog number: SLGP033RB)
Hemacytometer (INCYTO, catalog number: DHC-N015)
Dounce tissue grinder, 1 mL (Wheaton, catalog number: 357538)
DNA Clean & Concentrator-5 (Zymo Research, catalog number: D4014)
AMPure XP beads (Beckman Coulter, catalog number: A63881)
Adhesive PCR plate seal (Bio-Rad, catalog number: MSB1001)
Reagents
RNase ZapTM (Invitrogen, catalog number: AM9780)
0.4% Trypan Blue solution (Sigma, catalog number: T8154-20ML)
Nuclease-free water (not DEPC-treated) (Thermo Fisher, catalog number: AM9937)
Phosphate-buffered saline (PBS) (Thermo Fisher, catalog number: 10010049)
Fetal bovine serum (FBS) (Gibco, catalog number: 10099141C)
TrypLETM Express (Gibco, catalog number: 12604013)
Sucrose, reagent grade, 99% (Sigma, catalog number: V900116-500G)
HEPES, BioUltra (Sigma, catalog number: 54457-250G-F)
1 M potassium chloride (KCl) (Sigma, catalog number: 60142-100ML-F)
1 M magnesium chloride (MgCl2) (Thermo Fisher, catalog number: AM9530G)
0.1 M DL-Dithiothreitol solution (DTT) (Invitrogen, catalog number: 18090050)
RNase inhibitor (TaKaRa, catalog number: 2313B)
SUPERaseInTM RNase inhibitor (Invitrogen, catalog number: AM2694)
10% Igepal CA 630 (Sigma, catalog number: I8896-100ML)
Protease inhibitor cocktail (Sigma, catalog number: P8340-5ML)
16% formaldehyde solution (Thermo Fisher, catalog number: 28906)
Bovine serum albumin, powder (BSA) (Sigma, catalog number: B2064)
Bovine serum albumin (20 mg/mL BSA) (NEB, catalog number: B9000S)
Glycerol (Sigma, catalog number: 49767-250ML)
1 M Tris pH 8.0 (Invitrogen, catalog number: AM9855G)
5 M sodium chloride (NaCl) (Invitrogen, catalog number: AM9760G)
Sodium dodecyl sulfate solution, BioUltra, ~10% in H2O (10% SDS) (Sigma, catalog number: 71736-100ML)
TritonTM X-100 solution, BioUltra, ~10% in H2O (10% Triton X-100) (Sigma, catalog number: 93443)
10 mM deoxynucleotide (dNTP) solution mix (dNTPs) (NEB, catalog number: N0447L)
100 mM magnesium sulfate solution (MgSO4) (NEB, catalog number: M0259L)
Betaine solution, 5 M, PCR reagent (Sigma, catalog number: B0300-5VL)
5× SuperScriptTM IV RT buffer (Invitrogen, catalog number: 18090050)
SuperScriptTM IV reverse transcriptase (Invitrogen, catalog number: 18090050)
10× NEBuffer 2 (NEB, catalog number: B7002S)
25 U/μL MboI (NEB, catalog number: R0147M)
10× T4 DNA ligase reaction buffer (NEB, catalog number: B0202S)
T4 DNA ligase (1 U/μL) (Invitrogen, catalog number: 15224-025)
QIAGEN protease (30 AU) (QIAGEN, catalog number: 19157)
0.5 M Ethylenediaminetetraacetic acid, pH 8.0 (EDTA) (Invitrogen, catalog number: AM9260G)
10× ThermoPol® reaction buffer (NEB, catalog number: M0259L)
Deep Vent® (exo-) DNA polymerase (NEB, catalog number: M0259L)
TruePrep DNA Library Prep kit V2 for Illumina®, 5× TTBL (Vazyme, catalog number: TD501)
TruePrep DNA Library Prep kit V2 for Illumina®, TTE Mix V50 (Vazyme, catalog number: TD501)
KAPA HiFi PCR Kit, 5× KAPA HiFi GC buffer (Roche, catalog number: KK2102)
KAPA HiFi PCR Kit, KAPA HiFi DNA polymerase (1 U/μL) (Roche, catalog number: KK2102)
High Sensitivity NGS Fragment Analysis kit (Agilent, catalog number: DNF-474-0500)
i5 and i7 unique dual (UD) index (Illumina, catalog numbers: 20091654, 20091656, 20091658, 20091660)
Primers (synthesized by Sangon Biotech with PAGE purification)
GAT5-RT: GTAGGTGTGAGTGATGGTTGAGGTAGTATTGCGCAATGNNNNNNNN TTTTTTTTTTTTTTTVN
GAT5-7N: GTAGGTGTGAGTGATGGTTGAGGTAGTNNNNNNN
GAT5: GTAGGTGTGAGTGATGGTTGAGGTAGT
RNA-P7: GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGGGTTGAGGTAGT ATTGCGCAATG
Solutions
2% BSA (see Recipes)
1.5 M Sucrose (see Recipes)
60 mg/mL Qiagen protease (see Recipes)
100 mM HEPES buffer pH 8.0 (see Recipes)
2× SC lysis buffer (see Recipes)
Recipes
2% BSA (5 mL)
Reagent Final concentration Quantity
1 M Tris-HCl pH 8.0
1 M KCl
500 mM EDTA
H2O
BSA (powder)
40 mM
200 mM
0.2 mM
n/a
2% (w/v)
200 μL
1 mL
2 μL
3,798 μL
0.2 g
Total n/a 5 mL *
*Note: Filter through a 0.22 μm filter. Add an equal volume of glycerol and store at -20 °C (for long-term storage, keep at -80 °C).
1.5 M sucrose (10 mL)
Reagent Final concentration Quantity
Sucrose
H2O
1.5 M
n/a
5.13 g
10 mL
Total n/a 10 mL *
*Note: Filter through a 0.22 μm filter. Aliquot and store at -20 °C. H2O should be added in increments of 10 mL.
60 mg/mL Qiagen protease
Reagent Final concentration Quantity
7.5 AU Qiagen protease
H2O
60 mg/mL
n/a
One vial
2.78 mL
Total n/a 2.78 mL *
*Note: The minimum package volume to be purchased was 7.5 AU/tube. Add water to dissolve directly. Mix well and filter through a 0.2 μm filter. Aliquot and store at 4 °C (maximum storage time is two months).
100 mM HEPES buffer pH 8.0 (10 mL)
Reagent Final concentration Quantity
HEPES
H2O
100 mM
n/a
0.24 g
see Note*
Total n/a 10 mL *
*Note: First, add 9 mL of H2O. Allow the solution to mix completely before adding more. At this time, the liquid is acidic; adjust the pH of the solution to 8.0 with 1 M NaOH and then bring the volume up to 10 mL with H2O. Filter through a 0.22 μm filter. Aliquot and store at -20 °C.
2× SC lysis buffer (5 mL)
Reagent Final concentration Quantity
1 M Tris-HCl pH8.0
5 M NaCl
10% Triton X-100
500 mM EDTA
1 M DTT
H2O
40 mM
100 mM
0.3% (v/v)
2 mM
50 mM
n/a
200 μL
100 μL
150 μL
20 μL
250 μL
4,280 μL
Total n/a 5 mL *
*Note: Aliquot and store at -20 °C.
Equipment
Eppendorf ThermoMixer® C (Eppendorf, model: ThermoMixer C)
Vortex mixer (Scientific Industries, model: Vortex-Genie 2)
Centrifuge (Eppendorf, model: 5424)
Tube revolver rotator (Thermo, model: 88881002)
Thermocycler (Bio-Rad, model: C1000)
Magnetic separator (Invitrogen, model: DynaMagTM -2)
Fluorometer (Invitrogen, model: Qubit 4)
Flow cytometer (BD FACSAriaTM Cell Sorter, model: 65011040)
Bioanalyzer (Agilent Fragment Analyzer, model: 5200)
Software and datasets
Snakemake (v5.20.1, access date: Oct 19, 2023) [9] (Conda: https://anaconda.org/bioconda/snakemake)
hickit (v0.1.1, access date: Oct 19, 2023) (GitHub: https://github.com/lh3/hickit/)
CHARMtools (v0.1, access date: Oct 19, 2023) (GitHub: https://github.com/skelviper/CHARMtools)
Other software or packages listed in https://github.com/skelviper/HiRES/blob/main/HiRES_preprocess_pipeline/envs/main_env.yaml
Procedure
Animal maintenance
Raise Cast/EiJ male mice (purchased from Jackson Laboratory) and C57BL/6J female mice (purchased from Beijing HFK Bioscience Co., Ltd and Institutional Animal Care of Peking University) under 12:12 h brightness/darkness. Eight-week-old mice were mated naturally; the morning of plug detection was defined as embryonic day 0.5 (E0.5).
Tissue dissection and single-cell/nucleus isolation (0.5–2 h, depending on the number of mice and embryos)
Mouse embryos
Collet post-implantation embryos in PBS containing 10% FBS from E7.0 to E11.5 (see General note 1).
For E7.0–E8.5 embryos, collect all embryos from one mouse as a biological replicate. Dissociate embryos in 100 μL of TrypLETM Express with 200 μL tips pipetting up and down for 5 min.
For E9.5–E11.5 embryos, collect one or two embryos from one mouse as a biological replicate. Dissociate embryos in 200 μL of TrypLETM Express with 1 mL tips for 5 min and then with 200 μL tips, pipetting for another 5 min (see General note 2).
Add a 5-fold volume of PBS (e.g., 500 μL of PBS for 100 μL of TrypLETM Express) to inactivate TrypLETM Express.
Filter the cell suspension with a 40 μm cell strainer to a new 1.5 mL tube.
Centrifuge at 300× g for 5 min at 4 °C.
Remove supernatant and wash pellet with 500 μL of PBS.
Centrifuge at 300× g for 5 min at 4 °C.
Remove supernatant and resuspend pellet with 500 μL of PBS.
Stain a 5 μL aliquot of the cell suspension with 5 μL of 0.4% Trypan Blue. Pipette to mix thoroughly.
Transfer to a disposable hemocytometer to estimate the number of intact cells.
Adjust the number of cells to ~100,000 in 200 μL of PBS.
Mouse brains
Prepare homogenization buffer (1 mL for each sample), vortex to mix, and keep on ice.
1.5 M sucrose 167 μL
100 mM HEPES buffer pH 8.0 100 μL
1 M KCl 25 μL
1 M MgCl2 5 μL
1 mM DTT 1 μL
RNase inhibitor 10 μL
SUPERaseIn 10 μL
10% Igepal CA 630 10 μL
Protease inhibitor 100 μL
H2O 572 μL
Precool the Dounce homogenizer and pestles on ice. Once cooled, fill the homogenizer with 1 mL of cold homogenization buffer and keep it on ice.
Dissected left cortices from 8–9-week-old F1 hybrid mice in ice-cold PBS.
Transfer the cortex from one mouse to a Petri dish (on ice) and cut out a ~10 mm3 section using a chilled scalpel. Immediately transfer the tissue section into the precooled Dounce homogenizer (see General note 3).
Homogenize the tissue with five strokes of the loose pestle, followed by 15 strokes of the tight pestle.
Filter the homogenate through a 40 μm cell strainer to a new 1.5 mL tube.
Centrifuge at 300× g for 5 min at 4 °C and remove supernatant.
Prepare nuclei storage buffer (1.5 mL for each sample), vortex to mix, and keep on ice.
1.5 M sucrose 166.5 μL
100 mM HEPES buffer pH 8.0 150 μL
1 M KCl 37.5 μL
1 M MgCl2 7.5 μL
H2O 1,138.5 μL
Combine 500 μL nuclei storage buffer with 5 μL of RNase inhibitor and 50 μL of protease inhibitor and resuspend pellet.
Centrifuge at 300× g for 5 min at 4 °C.
Remove supernatant and resuspend pellet with 500 μL of nuclei storage buffer.
Stain a 5 μL aliquot of the cell suspension with 5 μL of 0.4% Trypan Blue. Pipette to mix thoroughly.
Transfer to a disposable hemocytometer to estimate the number of intact cells.
Adjust the number of nuclei to ~100,000 in 200 μL with the remaining nuclei storage buffer.
Cell fixation and permeabilization (1–1.5 h)
To fix 200 μL of cell or nucleus suspension, add 24.56 μL of 16% formaldehyde (final 1.75%) directly. Rotate with the tube revolver rotator at 10 rpm for 10 min at room temperature.
Add 20 μL of 2% BSA and invert to mix to quench formaldehyde.
Centrifuge at 1,000× g for 5 min at 4 °C.
Prepare wash buffer (200 μL for each sample), vortex to mix, and keep on ice.
1 M Tris-HCl pH 8.0 2 μL
5 M NaCl 0.4 μL
20 mg/mL BSA 1 μL
RNase inhibitor 2.6 μL
Protease inhibitor 20 μL
H2O 174 μL
Remove supernatant and resuspend cells with 200 μL of ice-cold wash buffer. Mix well by pipetting.
Centrifuge at 1,000× g for 5 min at 4 °C.
Prepare Hi-C lysis buffer (200 μL for each sample), vortex to mix, and keep on ice.
1 M Tris-HCl pH 8.0 2 μL
5 M NaCl 0.4 μL
10% Igepal CA 630 4 μL
RNase inhibitor 2.6 μL
Protease inhibitor 20 μL
H2O 171 μL
Remove supernatant and resuspend cells with 200 μL of ice-cold Hi-C lysis buffer. Mix well by pipetting.
Incubate on ice for 15 min.
Centrifuge at 1,000× g for 5 min at 4 °C.
Prepare 0.5% SDS buffer (100 μL for each sample) and vortex to mix.
10% SDS 5 μL
10 mM dNTPs 5 μL
H2O 90 μL
Remove supernatant and resuspend cells with 100 μL of 0.5% SDS buffer.
Incubate at 65 °C for 10 min and then immediately place on ice for 3 min.
Prepare SDS quench buffer (100 μL for each sample), vortex to mix, and keep on ice.
10% Triton X-100 50 μL
RNase inhibitor 2.6 μL
H2O 47.4 μL
Add 100 μL of ice-cold SDS quench buffer, mix by vortex, and place on ice for 5 min.
Centrifuge at 2,500× g for 5 min at 4 °C.
Reverse transcription (~0.5 h)
Prepare reverse transcription buffer (75 μL for each sample), vortex to mix, and keep on ice.
5 M Betaine 20 μL
10 mM dNTPs 5 μL
100 mM DTT 5 μL
50 μM GAT5-RT primer 5 μL
RNase inhibitor 1.5 μL
SUPERaseIn 1.5 μL
10% Triton X-100 1 μL
100 mM MgSO4 1 μL
H2O 35 μL
Remove supernatant and resuspend nuclei with 75 μL of ice-cold reverse transcription buffer.
Caution: In this step, the cell pellet becomes transparent. Care should be taken to aspirate the supernatant from the opposite side of the pellet to avoid loss.
Critical note: If a salt-containing buffer, such as 5× SSIV buffer, is introduced immediately after SDS treatment, it will result in cell clumping, which is difficult to reverse without cell lysis. Therefore, prior to adding 5× SSIV buffer, it is recommended to resuspend the pellet as thoroughly as possible with reverse transcription buffer to achieve a single-cell suspension.
Add 20 μL of 5× SuperScriptTM IV RT buffer and 5 μL of 200 U/μL SuperScriptTM IV reverse transcriptase (final 10 U/μL). Mix thoroughly by pipetting (see General note 4).
Incubate at 55 °C for 15 min.
Centrifuge at 2,500× g for 5 min at 4 °C.
Digestion and ligation (~17 h)
Prepare digestion buffer (600 μL for each sample) and vortex to mix.
10× NEB buffer 2 60 μL
10% Triton X-100 6 μL
H2O 534 μL
Remove supernatant and resuspend nuclei with 200 μL of digestion buffer.
Centrifuge at 2,500× g for 5 min at 4 °C.
Remove supernatant and resuspend nuclei with 200 μL of digestion buffer.
Centrifuge at 2,500× g for 5 min at 4 °C.
Remove supernatant and resuspend nuclei with 100 μL of digestion buffer.
Add 5 μL of 25 U/μL MboI. Mix by pipetting.
Incubate at 37 °C overnight.
The next day, incubate at 65 °C for 20 min to inactive MboI.
Add 790 μL of water and resuspend nuclei by pipetting.
Take a 20 μL aliquot of the sample as the digested control.
Then, add 100 μL of 10× T4 DNA ligase buffer, 5 μL of 20 mg/mL BSA, and 20 μL of 1 U/μL T4 DNA ligase. Mix by pipetting.
Incubate at 16 °C for 4 h.
Take a 20 μL aliquot of the sample as the ligated control.
Single nuclei flow sorting (0.5–2 h, depending on the number of nuclei needed)
Filter approximately 1 mL of nucleus suspension through a 40 μm cell strainer.
Sort a single nucleus into each well of an empty 96-well plate with a BD FACSAriaTM cell sorter. Forward vs. side scatter (FSC vs. SSC) gating is used to identify single nucleus populations and exclude debris and clumps.
Pause point: Sorted nuclei in dry 96-well plates can be stored at -80 °C for up to six months.
Single nucleus lysis and preamplification (~8 h for each 96-well plate)
Prepare lysis buffer (420 μL for each 96-well plate, 4 μL for each well) and vortex to mix. Dilute 60 mg/mL Qiagen protease to a concentration of 6 mg/mL first.
2× SC lysis buffer 210 μL
6 mg/mL Qiagen protease 37.8 μL
50 μM GAT5 4.2 μL
H2O 168 μL
Add 4 μL of lysis buffer to each well of a 96-well plate containing sorted nuclei with a multichannel pipette (see General note 5).
Briefly spin down and lyse the nuclei by running the following PCR program in a thermocycler:
50 °C 3 h
70 °C 1 h
4 °C ∞
Prepare amplification mixture (3.6 mL for each 96-well plate, 36 μL for each well) and vortex to mix.
10× ThermoPol buffer 400 μL
10 mM dNTPs 120 μL
50 μM GAT5-7N 60 μL
50 μM GAT5 60 μL
100 mM MgSO4 30 μL
Deep Vent (exo-) 100 μL
H2O 2,830 μL
Take the 96-well plate from the thermocycler.
Add 36 μL of amplification mixture to each well of the above 96-well plate with a multichannel pipette.
Caution: Avoid touching the liquid in the plate to prevent template loss before amplification.
Briefly spin down and perform amplification by running the following PCR program in a thermocycler:
1) 95 °C 5 min
2) 4 °C 50 s
3) 10 °C 50 s
4) 20 °C 50 s
5) 30 °C 50 s
6) 40 °C 45 s
7) 50 °C 45 s
8) 65 °C 4 min
9) 95 °C 20 s
10) 58 °C 20 s
11) Go to step 2) for 9 cycles
12) 95 °C 1 min
13) 95 °C 20 s
14) 58 °C 30 s
15) 72 °C 3 min
16) Go to step 13) for 14 cycles
17) 72 °C 5 min
18) 4 °C ∞
Pause point: Amplified DNA can be stored at -20 °C for up to six months.
Measure the concentration of single-nucleus preamplification product with QubitTM 1× dsDNA HS Assay kit through random sampling to ensure the efficacy of the amplification reaction in each 96-well plate (see General note 6).
Library preparation (~4 h for each 96-well plate, ~2 h for library construction, ~2 h for post-PCR cleanup)
Prepare transposition mixture and vortex to mix (318 μL for each 96-well plate, 3 μL for each well):
5× TTBL 106 μL
TTE mix V50 26.5 μL
H2O 185.5 μL
Add 3 μL of transposition mixture to each well of an empty 96-well plate with a multichannel pipette.
Add 2 μL of diluted single-nucleus preamplification product to each well of the above 96-well plate with a multichannel pipette and mix thoroughly.
Briefly spin down and run the following PCR program in a thermocycler:
55 °C 10 min
4 °C ∞
Prepare 200 μL of 0.2 % SDS:
10% SDS 4 μL
H2O 196 μL
Take the 96-well plate from the thermocycler.
Add 1.25 μL of 0.2 % SDS to each well of the above 96-well plate with a multichannel pipette and mix thoroughly.
Briefly spin down and incubate at room temperature for 10 min.
Prepare PCR mixture (1198.5 μL for each 96-well plate, 11.75 μL for each well) and vortex to mix.
5× KAPA HiFi GC buffer 408 μL
10 mM dNTPs 61.2 μL
50 μM RNA-P7 20.4 μL
KAPA HiFi DNA polymerase 40.8 μL
H2O 668.1 μL
Add 2 μL of 5 μM i5 unique dual (UD) index to each well of the above 96-well plate with a multichannel pipette.
Add 11.75 μL of PCR mixture to each well of the above 96-well plate with a multichannel pipette.
Briefly spin down and run the following PCR program in a thermocycler for a slight enrichment of cDNA reads:
1) 72 °C 3 min
2) 98 °C 30 s
3) 98 °C 15 s
4) 60 °C 30 s
5) 72 °C 2 min
6) Go to step 3) for 5 cycles
7) 4 °C ∞
Take the 96-well plate from the thermocycler and add 2 μL of 5 μM i7 unique dual (UD) index with a multichannel pipette.
Briefly spin down and run the following PCR program in a thermocycler:
1) 4 °C 3 min
2) 98 °C 30 s
3) 98 °C 15 s
4) 60 °C 30 s
5) 72 °C 2 min
6) Go to step 3) for 5 cycles
7) 72 °C 5 min
8) 4 °C ∞
Pool the barcoded single-cell libraries and purify pooled libraries with 0.6× and 0.15× AMPure XP beads. The final libraries were sequenced with paired-end 150-bp reads on a NovaSeq 6000 (Illumina) platform.
Quality assessment (1–2 h)
Lyse the digested control and ligated control by adding 1 μL of 60 mg/mL Qiagen protease and run the following PCR program in a thermocycler:
50 °C 1 h
70 °C 15 min
4 °C ∞
Purify the digested control and ligated control with Zymo DNA Clean & Concentrator-5 columns.
To ensure that the crucial steps of the experiment were effectively carried out, we measure size distribution of the samples after digestion, ligation, single-nucleus preamplification, library construction after PCR, and size selection (as shown in Figure 1).
Figure 1. Representative Bioanalyzer traces for the quality control of digestion (A), ligation (B), preamplification (C–D), and library preparation steps (E–F). All samples were run with the High Sensitivity NGS Fragment Analysis Kit on Agilent 5200 Fragment Analyzer.
Data analysis
Preprocessing of HiRES data to derive single-cell RNA expression matrices, single-cell chromatin contact matrices, and three-dimensional reconstruction structures from the original FASTQ sequencing data includes the following steps:
We recommend processing HiRES raw data using our HiRES pre-processing Snakemake workflow. The code can be downloaded from https://github.com/skelviper/HiRES. Please ensure that the workflow code and the folder containing the fastq data are placed within the same working directory.
Set up the environment by executing the following command:
conda create -n hires -c conda-forge -c bioconda python=3.8 snakemake=5.20.1.
Modify the paths to the dependent files in config.yaml and select a reference genome that is appropriate for your experiment.
Execute the workflow by navigating to the HiRES_preprocess_pipeline directory and running the script with the following commands: cd HiRES_preprocess_pipeline; ./runHiRES_preprocess.sh. By default, the workflow utilizes the Slurm scheduler, but you can modify runHiRES_preprocess.sh to use a scheduler that better suits your preference.
Generate experimental statistical data by executing the Jupyter Notebook found at analysis/stat.ipynb.
Use output files for other downstream data analysis.
With HiRES data, the gene expression level can be mapped onto the reconstructed 3D genome structure for the same single cell (as shown in Figure 2).
Figure 2. Representative example of a single-cell 3D genome structure and its corresponding gene expression. Compartmentalization of euchromatin (green) and heterochromatin (magenta) was visualized by CpG frequency as a proxy (left). The size of yellow balls was proportional to the gene expression level (right). Example single-cell data and figures are from reference [8].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Liu et al. (2023). Linking genome structures to functions by simultaneous single-cell Hi-C and RNA-seq. Science (Figure 1, panel B–D; Figure 2, panel A–D; Figure S2, panel C; Figure S3, panel B).
General notes and troubleshooting
General notes
Before dissection, keep the workstation and hood RNase-free by thoroughly cleaning with RNase Zap.
For steps B2 and B3, the digestion time for embryos at different stages could be slightly adjusted according to the homogeneity of the cell suspension. Pipetting should be performed softly to maintain cell integrity as much as possible.
To reduce heat caused by friction, Dounce homogenization should be performed on ice with gentle strokes, and care should be taken to avoid foaming. The mortar should be immersed in ice. The precooled homogenization buffer is an important aid in heat reduction during homogenization.
Increasing the volume of SuperScript IV Reverse Transcriptase to 10 μL led to more transcripts detected but was accompanied by a decrease in the detected number of contacts.
The lysis buffer should be carefully added along the tube wall, while ensuring that the pipette tip does not reach the bottom of the tube to prevent loss of single nuclei. If subsequent experiments involving cell lysis can be conducted immediately after flow sorting, the single nuclei can be directly sorted into a 96-well plate containing lysis buffer.
The concentration of a single-nucleus preamplification product is generally between 30 and 50 ng/μL. For convenience, we generally dilute the product by a factor of 10 using water, resulting in a diluted product concentration of approximately 3–5 ng/μL. Each diluted sample was used for library preparation.
If the assay is intended to be applied to other biological samples, it might be necessary to modify the procedures for obtaining single-cell suspensions according to the specific characteristics of different tissues.
Troubleshooting
Insufficient single nuclei available for flow cytometry sorting.
Potential solution: if a substantial loss of nuclei is observed after each centrifugation, consider extending the centrifugation duration as needed.
Cell loss or cell aggregation after SDS treatment.
Potential solution: if the transparent cell pellet cannot be found after SDS treatment, aspirate the supernatant slowly from the opposite side of the pellet to avoid loss and resuspend the pellet by pipetting at least 20 times to avoid cell aggregation.
No or low-yield single-nucleus amplification product (i.e., less than 10 ng/μL) after preamplification.
Potential solution: check reagents that are crucial for lysis and preamplification. For example, Qiagen protease must be present in lysis buffer and the final concentration must be appropriate.
Acknowledgments
This protocol is based on the original research paper of Liu et al., 2023. This work was supported by the National Natural Science Foundation of China (grant 92049110) and the funding from Beijing Advanced Innovation Center for Genomics (ICG).
Competing interests
D.X., Y.C., and Z.L. are investors on a patent that covers HiRES. H.X. declares no competing interests.
Ethical considerations
All animal maintenance and experimental procedures were carried out in accordance with the guidelines of the Institutional Animal Care and Use Committee (IACUC) of Peking University.
References
Angermueller, C., Clark, S. J., Lee, H. J., Macaulay, I. C., Teng, M. J., Hu, T. X., Krueger, F., Smallwood, S. A., Ponting, C. P., Voet, T., et al. (2016). Parallel single-cell sequencing links transcriptional and epigenetic heterogeneity. Nat. Methods 13(3): 229–232. doi: 10.1038/nmeth.3728
Stoeckius, M., Hafemeister, C., Stephenson, W., Houck-Loomis, B., Chattopadhyay, P. K., Swerdlow, H., Satija, R. and Smibert, P. (2017). Simultaneous epitope and transcriptome measurement in single cells. Nat. Methods 14(9): 865–868. doi: 10.1038/nmeth.4380
Ma, S., Zhang, B., LaFave, L. M., Earl, A. S., Chiang, Z., Hu, Y., Ding, J., Brack, A., Kartha, V. K., Tay, T., et al. (2020). Chromatin Potential Identified by Shared Single-Cell Profiling of RNA and Chromatin. Cell 183(4): 1103–1116.e20. doi: 10.1016/j.cell.2020.09.056
Zhu, C., Zhang, Y., Li, Y. E., Lucero, J., Behrens, M. M. and Ren, B. (2021). Joint profiling of histone modifications and transcriptome in single cells from mouse brain. Nat. Methods 18(3): 283–292. doi: 10.1038/s41592-021-01060-3
Zheng, H. and Xie, W. (2019). The role of 3D genome organization in development and cell differentiation. Nat. Rev. Mol. Cell Biol. 20(9): 535–550. doi: 10.1038/s41580-019-0132-4
Hafner, A. and Boettiger, A. (2022). The spatial organization of transcriptional control. Nat. Rev. Genet. 24(1): 53–68. doi: 10.1038/s41576-022-00526-0
Tan, L., Ma, W., Wu, H., Zheng, Y., Xing, D., Chen, R., Li, X., Daley, N., Deisseroth, K., Xie, X. S., et al. (2021). Changes in genome architecture and transcriptional dynamics progress independently of sensory experience during post-natal brain development. Cell 184(3): 741–758.e17. doi: 10.1016/j.cell.2020.12.032
Liu, Z., Chen, Y., Xia, Q., Liu, M., Xu, H., Chi, Y., Deng, Y. and Xing, D. (2023). Linking genome structures to functions by simultaneous single-cell Hi-C and RNA-seq. Science 380(6649): 1070–1076. doi: 10.1126/science.adg3797
Mölder, F., Jablonski, K. P., Letcher, B., Hall, M. B., Tomkins-Tinch, C. H., Sochat, V., Forster, J., Lee, S., Twardziok, S. O., Kanitz, A., et al. (2021). Sustainable data analysis with Snakemake. F1000Research 10: 33. doi: 10.12688/f1000research.29032.2
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Systems Biology > Genomics > Sequencing
Molecular Biology > DNA > DNA sequencing
Cell Biology > Single cell analysis
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Improve Research Reproducibility
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Peer-reviewed
Phospholipid Preparations to Characterize Protein–Lipid Interactions In Vitro
LM Lisa Merklinger
JM J. Preben Morth
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4887 Views: 475
Reviewed by: Julie WeidnerJohn P PhelanPhilipp A.M. Schmidpeter
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Original Research Article:
The authors used this protocol in Life Science Alliance Apr 2022
Abstract
The lipid bilayers of the cell are composed of various lipid classes and species. These engage in cell signaling and regulation by recruiting cytosolic proteins to the membrane and interacting with membrane-embedded proteins to alternate their activity and stability. Like lipids, membrane proteins are amphipathic and are stabilized by the hydrophobic forces of the lipid bilayer. Membrane protein–lipid interactions are difficult to investigate since membrane proteins need to be reconstituted in a lipid-mimicking environment. A common and well-established approach is the detergent-based solubilization of the membrane proteins in detergent micelles. Nowadays, nanodiscs and liposomes are used to mimic the lipid bilayer and enable the work with membrane proteins in a more natural environment. However, these protocols need optimization and are labor intensive. The present protocol describes straightforward instructions on how the preparation of lipids is performed and how the lipid detergent mixture is integrated with the membrane protein MARCH5. The lipidation protocol was performed prior to an activity assay specific to membrane-bound E3 ubiquitin ligases and a stability assay that could be used for any membrane protein of choice.
Keywords: Membrane protein–lipid interaction Detergent Phospholipids Lipid-membrane strip Lipid preparation
Background
Membrane proteins interact with their lipid environment with the hydrophobic and hydrophilic patches of their transmembrane domains via a specific binding site (non-annular lipids) or the physiochemical properties of the lipid bilayer (annular lipids) (Contreras et al., 2011; Stangl and Schneider, 2015). These protein–lipid interactions can alter the activity, stability, folding, or localization of the target protein, leading to various cellular signaling events (Engelman, 2005; van Meer et al., 2008; Corradi et al., 2019a;). The characterization of these signaling mechanisms and the corresponding lipid–membrane protein interaction is a constant challenge due to the inherent phase difference imposed on the system from the presence of a mixture of detergent/lipid, lipid/protein, and detergent/lipid/protein (Sych et al., 2022). Therefore, research into the mechanisms used by lipids to regulate protein function and structure is restricted to a small number of membrane protein systems, often because of the technical difficulties as well as complexity associated with handling these systems in a laboratory setting (Fahy et al., 2009). Hence, understanding the mechanism of how lipids can modulate the activity of a specific membrane protein and the identification of the lipids that interact with the membrane protein of interest is often still an open and fundamental research question (Corradi et al., 2019b).
Phospholipids are the most abundant lipids in mammalian membranes and are characterized by their hydrophilic head group and their hydrophobic tail, which is built by two fatty acid chains. These fatty acid chains can vary in length and saturation, creating an immense variety (Horvath and Daum, 2013; Cockcroft, 2021). The chemically diverse hydrophilic head groups define the lipid classes (Table 1), whereas the different lengths and saturation subdivide the class in different lipid species (Liebisch et al., 2020). The lipid composition of the lipid bilayer is specific to the tissue and the cellular compartment, which comprises a distinct set of phospholipids (Table 1) (Ernst et al., 2016).
Table 1. Overview of phospholipids, their chemical characteristic, and the abundance of subcellular fractions [plasma membrane, mitochondria, and endoplasmic reticulum (ER)) of rat liver (Horvath and Daum, 2013)]
Phospholipid class Characteristic Abundance in cellular compartment (%)
Plasma membrane Mitochondria ER
Phosphatidylethanolamine (PE) zwitterionic 40 34 60
Cardiolipin (CL) anionic 1 14 1
Phosphatidic acid (PA) anionic 1 < 1 1
Phosphatidylcholine (PC) zwitterionic 24 44 23
Phosphatidylglycerol (PG) anionic - - -
Phosphatidylinositol phosphate (PI) anionic 8 5 10
Nowadays, different membrane-mimicking systems are available to solvate membrane proteins. The most common method for working with membrane proteins in vitro is to solubilize them in detergent. The detergent forms a micelle layer around the hydrophobic surface patches of the protein, leading to the stabilization of the membrane (or membrane-associated) protein outside the natural lipid bilayer. Over the last years, alternatives to detergent-based methods were developed, such as nanodiscs (Pettersen et al., 2023), styrene–maleic acid copolymers (SMAs) (Dörr et al., 2016), or liposomes (Verchère et al., 2017). However, these methods often need a detergent-solubilized membrane protein as the initial step and optimization is often labor intensive. Here, we describe two easy-to-establish and straightforward protocols to examine the interaction and effect of different lipid classes on the activity and stability of purified and detergent-solubilized MARCH5 in vitro. The described protocols were initially used to test lipid stability in connection with the human integral membrane protein SERINC5 (Pye et al., 2020) and, since then, tested against a broader spectrum of membrane proteins (Cecchetti et al., 2021).
Materials and reagents
Biological materials
Protein of interest (a membrane protein solubilized in detergent); in this protocol, using MARCH5 as benchmarking example (see Merklinger et al., 2022).
Reagents
Lipid preparation for activity and stability assays
Egg PE (Avanti Polar Lipids, catalog number: 840021)
Heart CL (Avanti Polar Lipids, catalog number: 840012)
Egg PA (Avanti Polar Lipids, catalog number: 840101)
Egg PC (Avanti Polar Lipids, catalog number: 840051)
Egg PG (Avanti Polar Lipids, catalog number: 841138)
Brain PI(4)P (Avanti Polar Lipids, catalog number: 840045)
Lauryl maltose neopentyl glycol (LMNG) (Anatrace, catalog number: NG310)
Argon gas
Lipid-binding assay
LMNG (Anatrace, catalog number: NG310)
Trizma base (Sigma-Aldrich, catalog number: 77-86-1)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S7653)
BSA (Sigma-Aldrich, catalog number: A9418)
PierceTM ECL western blotting substrate (Thermo Fischer Scientific, catalog number 32132)
Amersham HydondTM-LFP, Hybond LFP polyvinylidene fluoride or polyvinylidene difluoride (PVDF) transfer membrane (GE Healthcare, catalog number: RPN303LFP)
Egg PE (Avanti Polar Lipids, catalog number: 840021)
Heart CL (Avanti Polar Lipids, catalog number: 840012)
Egg PA (Avanti Polar Lipids, catalog number: 840101)
Egg PC (Avanti Polar Lipids catalog number: 840051)
Egg PG (Avanti Polar Lipids catalog number: 841138)
Brain Phosphatidylinositol-4-phosphate (PI(4)P) (Avanti Polar Lipids, catalog number: 840045)
Anti-MARCH5 antibody N-terminal (Abcam, catalog number: ab185054)
Anti-rabbit IgG, HRP-linked Antibody (Cell Signalling, catalog number: 7074)
Solutions
0.25% LMNG (see Recipes)
Tris-buffered saline (see Recipes)
Tris-buffered saline supplemented with 0.001% LMNG (see Recipes)
Blocking solution (TBS supplemented with 0.001% LMNG and 3% BSA) (see Recipes)
500× diluted Anti-MARCH5 antibody in TBS supplemented with 0.001% LMNG and 3% BSA (see Recipes)
2,000× diluted Anti-rabbit IgG antibody in TBS supplemented with 0.001% LMNG and 3% BSA (see Recipes)
Recipes
0.25% LMNG
Reagent Final concentration Quantity
LMNG 0.25% (w/v) 0.0025 g
H2O n/a 10 mL
Total 0.25% 10 mL
TBS (Tris-buffered saline)
Reagent Final concentration Quantity
Trizma base 50 mM 6.05 g
NaCl 150 mM 8.76 g
H2O 1,000 mL
Total 1,000 mL
pH is adjusted to 7.5 using HCl
TBS supplemented with 0.001% LMNG
Reagent Final concentration Quantity
TBS (see Recipe 2) 30 mL
0.25% LMNG (see Recipe 1) 0.001% 0.12 mL
Total 30 mL
Blocking solution
Reagent Final concentration Quantity
TBS (see Recipe 2) 30 mL
0.25% LMNG (see Recipe 1) 0.001% 0.12 mL
BSA 3% 0.9 g
Total 30 mL
500× diluted Anti-MARCH5 in TBS supplemented with 0.001% LMNG and 3% BSA
Reagent Final concentration Quantity
Blocking solution 5 mL
Anti-MARCH5 antibody 1:500 dilution 0.01 mL
Total 5 mL
2,000× diluted Anti-rabbit IgG antibody in TBS supplemented with 0.001% LMNG and 3% BSA
Reagent Final concentration Quantity
Blocking solution 5 mL
Anti-rabbit IgG antibody 1:2,000 dilution 0.0025 mL
Total 5 mL
Laboratory supplies
FisherbrandTM Borosilicate glass light-walled culture tubes (Fisher Scientific, catalog number: 11517403, diameter 12 mm, length 75 mm)
Eppendorf tubes (Eppendorf, catalog number: 0030120086)
Equipment
Pulsing Vortex mixer (VWR, catalog number: 10153-812)
Plate shaker (VWR, catalog number: 700-0246)
ChemiDocTM Touch Gel imaging system (Bio-Rad, catalog number: 1708370)
Procedure
Lipid preparation
Important: work in a laminar flow hood when working with chloroform and argon gas.
Fill round-bottom glass tubes with argon gas.
Transfer 20 μL of Egg PE, Heart CL, Egg PA, Egg PG, and Egg PC chloroform solutions and 50 μL of Brain PI(4)P chloroform solution (all stored at -20 °C under argon gas) to a separate argon gas–filled round-bottom glass tube. If lipid combinations are to be tested, it is possible to dry the lipids in the same round-bottom tube in the desired amounts. This reduces the detergent concentration in the protein–lipid mix.
Dry the lipids under an argon stream and shake on a pulsing vortex mixer for 15 min.
Re-suspend dry lipids to 5 μM under shaking, depending on their average molecular weight:
Egg PE (MW: 746 Da) in 54 μL of 0.25% LMNG.
Heart CL (MW: 1494 Da) 27 μL of 0.25% LMNG.
Egg PA (MW: 711 Da) 57 μL of 0.25% LMNG.
Egg PC (MW: 770 Da) 52 μL of 0.25% LMNG.
Egg PG (MW: 782) 51 μL of 0.25% LMNG.
Brain PI(4)P (MW: 974) 10 μL of 0.25% LMNG.
Add 1.8 μL of the 5 μM lipid solution to 50 μL of 32 μM purified MARCH5 (1 mg/mL) in 100 mM NaCl; 50 mM Tris pH 7.7; 5% glycerol; 0.001% LMNG; 0.1 mM TCEP (MARCH5 is incubated with the different lipids in a 1:10 molar ratio).
Incubate the lipid–protein solution on ice for 1 h.
The protein is now considered ready for activity and/or stability measurements.
Note: Generate lipid stocks that take the lipid:detergent ratio into consideration in the final assay conditions. Thus, the lipid stocks might have to be adjusted to keep the reaction mixture volumes equal, while still keeping the final critical micelle concentration (CMC) equivalent throughout the assay. This can be done by adjusting the detergent:lipid ratios already in the stock solution. However, this also limits the maximal detergent:lipid ratio that can be tested in solution.
Lipid-binding assay (Figure 1A)
Cut a rectangle (2 cm × 6 cm) of the PVDF membrane.
Spot 1 μL of the chloroform solutions of Egg PE, Heart CL, Egg PA, Egg PC, Egg PG, and Brain PI(4)P on the PVDF membrane at approximately 1 cm of distance.
Dry the spotted lipid for 1 h at room temperature (RT) in a laminar flow hood.
Spot positive controls on the dry membrane lipid strip:
Spot 1 μL of Anti-rabbit IgG, HRP-linked antibody on the membrane lipid strip.
Spot 1 μL of 0.35 mg/mL target protein.
Dry positive controls for 15 min.
Block the PVDF membrane with 30 mL of TBS supplemented with 3% BSA overnight at 4 °C with soft agitation on a plate shaker.
Remove the blocking solution.
Add 10 mL of 1 μg/mL of MARCH5 in TBS supplemented with 0.001% LMNG and 3% BSA (1 CMC of LMNG is added to ensure MARCH5 will not precipitate during incubation) to the PVDF membrane.
Incubate the PVDF membrane and MARCH5 for 4 h at 4 °C with soft agitation on a plate shaker.
Wash the PVDF membrane three times with 5 mL of TBS supplemented with 0.001% LMNG for 5 min at RT under soft agitation.
Add 5 mL of Anti-MARCH5 antibody N-terminal in TBS supplemented with 0.001% LMNG and 3% BSA to the PVDF membrane.
Incubate anti-MARCH5 antibody N-terminal for 4 h at 4 °C.
Remove the antibody solution and rinse the PVDF with 5 mL of TBS supplemented with 0.001% LMNG.
Add anti-rabbit IgG, HRP-linked Antibody in 5 mL of TBS supplemented with 0.001% LMNG and 3% BSA to the PVDF membrane.
Wash again the membrane lipid strip three times in 5 mL of TBS supplemented with 0.001% LMNG for 5 min at RT under soft agitation.
Prepare PierceTM ECL western blotting substrate, add onto the membrane lipid strip, and incubate for 1 min.
Measure chemiluminescence using the ChemiDocTM Touch Gel imaging system (Figure 1B and 1C).
Note: This assay will give you a yes/no answer on lipid membrane protein binding (see Figure 1).
Figure 1. Lipid binding assay. A. Schematic overview of lipid binding assay. B. Example of positive hits. Cardiolipin and PI4P interact with the protein of interest, indicated by the dark dots. The protein of interest spotted in the left-right corner is a positive control (+ cntl). C. Example of no lipid-protein interaction. The positive control (+ cntl) shows a positive signal indicating that the assay has worked properly.
Validation of protocol
Lipid preparation
The phospholipids prepared as described in this protocol were used in differential scanning fluorimetry as well as ubiquitination assays published in the article “Phospholipids alter activity and stability of mitochondrial membrane-bound ubiquitin ligase MARCH5”, Life Science Alliance Apr 2022, DOI: 10.26508/lsa.202101309 (see Figure 1D, Figures 2 and 4) (Merklinger et al., 2022). The experiments were performed in three independent experiments and showed consistent results. Similar experiments were performed by Cecchetti et al., investigating protein stability in different lipids (Cecchetti et al., 2021).
Lipid-binding assay
The lipid-binding assay described in this protocol was used to investigate the binding of MARCH5 to various lipids published in the article “Phospholipids alter activity and stability of mitochondrial membrane-bound ubiquitin ligase MARCH5”, Life Science Alliance Apr 2022, DOI: 10.26508/lsa.202101309 (see Figure 1B and S1). It was also used to investigate DRP1 interaction with PA (Adachi et al., 2016). Furthermore, commercially available lipid membrane strips are available from Echelon Biosciences Inc. (https://www.echelon-inc.com/product/membrane-lipid-strips).
General notes and troubleshooting
General notes
Lipid preparation
The purchased phospholipid classes are a mixture of different lipid species, meaning they vary in chain length and saturation (e.g., CL is a mix of 18:1 and 18:2 fatty acid chains) and are from natural origin. This allows us to study the whole lipid class and not only a specific lipid species. However, specific lipid species can be purchased as well. Purchasing lipids dissolved in chloroform makes the handling of the lipids easier (to avoid weighing out small quantities).
Changes in experimental output over time could be due to oxidized phospholipids. It is important to store the phospholipids under inert conditions at -20 °C (under argon or nitrogen).
Lipid binding assay
During the incubation and wash steps, the membrane should always be kept wet and never dry out.
Mark the membrane by cutting one edge to be aware of the orientation and the order of the spotted lipids.
Troubleshooting
Lipid binding assay
The best protein concentration for the lipid-binding assay has to be experimentally determined. A good starting concentration is 1 mg/mL.
The incubation time will need to be optimized for each protein of choice. If the incubation of the membrane protein is kept for too long, this might affect the stability of the protein, and this could lead to errors in the stability measurement and loss of reproducibility. At high membrane protein concentrations, aggregational effect might influence the thermal shift assay measurement; this could also lead to loss of reproducibility.
It is advisable to test a subset of different buffer conditions, i.e., testing the high and low concentrations of the desired salt in combination with variable pH ranges, to ensure that the average state of the membrane protein remain consistent during the time in which the assay is performed.
Acknowledgments
The protocol was first published in Life Science Alliance in April 2022 (DOI: 10.26508/lsa.202101309). We thank all authors for their contribution to this work and the Technical University of Denmark for supporting the project.
Competing interests
The authors declare that they have no conflicts of interest.
References
Adachi, Y., Itoh, K., Yamada, T., Cerveny, K. L., Suzuki, T. L., Macdonald, P., Frohman, M. A., Ramachandran, R., Iijima, M., Sesaki, H., et al. (2016). Coincident Phosphatidic Acid Interaction Restrains Drp1 in Mitochondrial Division. Mol. Cell 63(6): 1034–1043.
Cecchetti, C., Strauss, J., Stohrer, C., Naylor, C., Pryor, E., Hobbs, J., Tanley, S., Goldman, A. and Byrne, B. (2021). A novel high-throughput screen for identifying lipids that stabilise membrane proteins in detergent based solution. PLoS One 16(7): 1–20.
Cockcroft, S. (2021). Mammalian lipids: structure, synthesis and function. Essays Biochem. 65(5): 813–845.
Contreras, F. X., Ernst, A. M., Wieland, F. and Brugger, B. (2011). Specificity of Intramembrane Protein-Lipid Interactions. Cold Spring Harbor Perspect. Biol. 3(6): a004705–a004705.
Corradi, V., Sejdiu, B. I., Mesa-Galloso, H., Abdizadeh, H., Noskov, S. Y., Marrink, S. J. and Tieleman, D. P. (2019a). Emerging Diversity in Lipid–Protein Interactions. Chem. Rev. 119(9): 5775–5848.
Corradi, V., Sejdiu, B. I., Mesa-Galloso, H., Abdizadeh, H., Noskov, S. Y., Marrink, S. J. and Tieleman, D. P. (2019b). Emerging Diversity in Lipid–Protein Interactions. Chem. Rev. 119(9): 5775–5848.
Dörr, J. M., Scheidelaar, S., Koorengevel, M. C., Dominguez, J. J., Schäfer, M., van Walree, C. A. and Killian, J. A. (2016). The styrene–maleic acid copolymer: a versatile tool in membrane research. Eur. Biophys. J. 45(1): 3–21.
Engelman, D. M. (2005). Membranes are more mosaic than fluid. Nature 438(7068): 578–580.
Ernst, R., Ejsing, C. S. and Antonny, B. (2016). Homeoviscous Adaptation and the Regulation of Membrane Lipids. J. Mol. Biol. 428(24): 4776–4791.
Fahy, E., Subramaniam, S., Murphy, R. C., Nishijima, M., Raetz, C. R., Shimizu, T., Spener, F., van Meer, G., Wakelam, M. J., Dennis, E. A., et al. (2009). Update of the LIPID MAPS comprehensive classification system for lipids. J. Lipid Res. 50: S9–S14.
Horvath, S. E. and Daum, G. (2013). Lipids of mitochondria. Prog. Lipid Res. 52(4): 590-614.
Liebisch, G., Fahy, E., Aoki, J., Dennis, E. A., Durand, T., Ejsing, C. S., Fedorova, M., Feussner, I., Griffiths, W. J., Köfeler, H., et al. (2020). Update on LIPID MAPS classification, nomenclature, and shorthand notation for MS-derived lipid structures. J. Lipid Res. 61(12): 1539–1555.
Merklinger, L., Bauer, J., Pedersen, P. A., Damgaard, R. B. and Morth, J. P. (2022). Phospholipids alter activity and stability of mitochondrial membrane-bound ubiquitin ligase MARCH5. Life Sci. Alliance 5(8): 1–13.
Pettersen, J. M., Yang, Y. and Robinson, A. S. (2023). Advances in nanodisc platforms for membrane protein purification. Trends Biotechnol. 41(8): 1041–1054.
Pye, V. E., Rosa, A., Bertelli, C., Struwe, W. B., Maslen, S. L., Corey, R., Liko, I., Hassall, M., Mattiuzzo, G., Ballandras-Colas, A., et al. (2020). A bipartite structural organization defines the SERINC family of HIV-1 restriction factors. Nat. Struct. Mol. Biol. 27(1): 78–83.
Stangl, M. and Schneider, D. (2015). Functional competition within a membrane: Lipid recognition vs. transmembrane helix oligomerization. Biochim. Biophys. Acta Biomembr. 1848(9): 1886–1896.
Sych, T., Levental, K. R. and Sezgin, E. (2022). Lipid–Protein Interactions in Plasma Membrane Organization and Function. Annu. Rev. Biophys. 51(1): 135–156.
van Meer, G., Voelker, D. R. and Feigenson, G. W. (2008). Membrane lipids: where they are and how they behave. Nat. Rev. Mol. Cell Biol. 9(2): 112–124.
Verchère, A., Broutin, I. and Picard, M. (2017). Reconstitution of Membrane Proteins in Liposomes. Methods Mol. Biol. 259–282.
Article Information
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biochemistry > Lipid > Lipid-protein interaction
Molecular Biology > Protein > Stability
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4,888 | https://bio-protocol.org/en/bpdetail?id=4888&type=0 | # Bio-Protocol Content
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Peer-reviewed
Microinjection of β-glucan Into Larval Zebrafish (Danio rerio) for the Assessment of a Trained-Like Immunity Phenotype
HD Hannah Darroch
JA Jonathan W. Astin
CH Christopher J. Hall
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4888 Views: 576
Reviewed by: Alba BlesaWei FanAlberto Rissone
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Original Research Article:
The authors used this protocol in Developmental & Comparative Immunology Jul 2022
Abstract
The innate immune system can remember previous inflammatory insults, enabling long-term heightened responsiveness to secondary immune challenges in a process termed “trained immunity.” Trained innate immune cells undergo metabolic and epigenetic remodelling and, upon a secondary challenge, provide enhanced protection with therapeutic potential. Trained immunity has largely been studied in innate immune cells in vitro or following ex vivo re-stimulation where the primary insult is typically injected into a mouse, adult zebrafish, or human. While highly informative, there is an opportunity to investigate trained immunity entirely in vivo within an unperturbed, intact whole organism. The exclusively innate immune response of larval zebrafish offers an attractive system to model trained immunity. Larval zebrafish have a functional innate immune system by 2 days post fertilisation (dpf) and are amenable to high-resolution, high-throughput analysis. This, combined with their optical transparency, conserved antibacterial responses, and availability of transgenic reporter lines, makes them an attractive alternative model to study trained immunity in vivo. We have devised a protocol where β-glucan (one of the most widely used experimental triggers of trained immunity) is systemically delivered into larval zebrafish using microinjection to stimulate a trained-like phenotype. Following stimulation, larvae are assessed for changes in gene expression, which indicate the stimulatory effect of β-glucan. This protocol describes a robust delivery method of one of the gold standard stimulators of trained immunity into a model organism that is highly amenable to several non-invasive downstream analyses.
Key features
• This protocol outlines the delivery of one of the most common experimental stimulators of trained immunity into larval zebrafish.
• The protocol enables the assessment of a trained-like phenotype in vivo.
• This protocol can be applied to transgenic or mutant zebrafish lines to investigate cells or genes of interest in response to β-glucan stimulation.
Graphical overview
Keywords: Larval zebrafish Microinjection Trained immunity Innate immunity response β-glucan
Background
Trained immunity describes the phenomenon of a long-lived shift in responsiveness of innate immunity following an earlier immune challenge, whereby the innate immune system remembers previous infectious encounters. Training induces metabolic and epigenetic changes within innate immune cells (including macrophages, neutrophils, and natural killer cells) as well as non-immune cells (such as fibroblasts, intestinal stromal cells, and epithelial stem cells) that typically result in a heightened response to a secondary challenge (Hamada et al., 2019; Netea et al., 2020). Trained immunity has been induced by a range of stimuli including vaccines (Kleinnijenhuis et al., 2012; Arts et al., 2018; Kaufmann et al., 2018), metabolites (Arts et al., 2016; Ferreira et al., 2023), and fungal polysaccharides such as β-glucan (Arts et al., 2016; Mitroulis et al., 2018; Kalafati et al., 2020). Trained immunity has been largely studied in vitro or following ex vivo re-stimulation of cells extracted from trained mice, adult zebrafish, or humans (Kleinnijenhuis et al., 2012; Kaufmann et al., 2018; Rodríguez et al., 2009; Megías et al., 2016; Arts et al., 2018). While highly informative, in vitro and ex vivo re-stimulation of cells may not fully mimic the in vivo response (Silliman and Wang, 2006). A vertebrate model where the innate immune response can be analysed entirely in vivo, such as larval zebrafish, offers an attractive additional model to investigate trained immunity.
Larval zebrafish possess a functional innate immune system by 2 days post fertilisation (dpf) but only develop a functionally mature adaptive immune system by 4–6 weeks post fertilisation (Lam et al., 2004). This chronological separation enables the study of the innate immune response in isolation during larval stages. Additionally, larval zebrafish offer powerful downstream analysis techniques including single-cell resolution microscopy using transgenic reporter lines to directly observe innate immune cell (macrophage and neutrophil) responses. β-glucan is a conventional training stimulus with well-characterised immune-potentiating effects that have been demonstrated in both adult and larval zebrafish (Rodríguez et al., 2009; Medina-Gali et al., 2018; Darroch et al., 2022). We recently showed that systemic injection of β-glucan induces a trained-like phenotype in larval zebrafish that elevates survival to subsequent infectious challenges and increases the recruitment of neutrophils to infection (Darroch et al., 2022). In this protocol, we outline a method for microinjecting β-glucan into the circulation of larval zebrafish. A soluble form of β-glucan that does not trigger the activation of Dectin-1 (the main pattern recognition receptor involved in the detection of fungal polysaccharides) or downstream NF-kB signalling is used as a control (Roesner et al., 2019). Using this protocol, the expression of genes involved in the NF-kB pathway (ikbaa, nfkb1, nfkb2) as well as genes expressing inflammatory cytokines (il1b, tnfa, cxcl8a, and il10) are assessed following β-glucan stimulation (Rogers et al., 2005; Inoue and Shinohara, 2014; Megías et al., 2016). We found that two of these genes, il10 and tnfa, are significantly upregulated in larval zebrafish at 1 day post stimulation (Darroch et al., 2022). This protocol can facilitate the investigation of downstream β-glucan-induced changes in innate immune responses using different infection models and the large array of available zebrafish transgenic reporter lines.
Materials and reagents
Biological materials
Zebrafish (Danio rerio)
Reagents
TRIzolTM reagent (Invitrogen, catalog number: 15596026)
Chloroform (Sigma-Aldrich, catalog number: C2432)
Isopropanol (2-propanol) (Sigma-Aldrich, catalog number: I9516)
Ethanol absolute (Sigma-Aldrich, catalog number: 1.07017)
β-glucan peptide from Trametes versicolor (Invivogen, catalog code: tlrl-bgp)
Whole glucan particles (soluble, β-glucan control) (Invivogen, catalog number: tlrl-wgps)
Methylcellulose (Sigma-Aldrich, catalog number: M0512)
Phenol red (Sigma-Aldrich, catalog number: P3532)
Tricaine (Ethyl 3-aminobenzoate methanesulfonate salt) (Sigma-Aldrich, catalog number: A5040)
Phosphate buffered saline (PBS), pH 7.4 (Gibco, catalog number: 10010023)
Ultrapure water (Invitrogen, catalog number: 10977015)
MilliQ water
Mineral oil (Sigma-Aldrich, catalog number: M5904)
1-phenyl-2-thiourea (PTU) (Sigma-Aldrich, catalog number: P7629)
iScriptTM cDNA Synthesis kit (Bio-Rad, catalog number: 1708891)
PerfeCTa SYBR Green FastMix with ROX (Quanta Biosciences, catalog number: 95055-500)
1 M Tris (pH 9) (Thermo Scientific, catalog number: J62085-K2)
KCl (Sigma-Aldrich, catalog number: P9541)
CaCl2 (Sigma-Aldrich, catalog number: C4901)
MgSO4 (Sigma-Aldrich, catalog number: 208094)
NaCl (Sigma-Aldrich, catalog number: S3014)
RT qPCR primers—diluted 10 μM in ultrapure water (IDT) (Table 1)
Table 1. RT qPCR primer sequences
Gene Forward (5′ – 3′) Reverse (5′ – 3′)
ef1a TGCCTTCGTCCCAATTTCAG TACCCTCCTTGCGCTCAATC
il6 AGACGAGCAGTTTGAGAGAGATT GTTTGAGGAGAGGAGTGCTGAT
cxcl8a CTGCATTGAAACAGAAAGCC CTTGACTTCACAGGTGATCC
tnfa CCAGGGCAATCAACAAGATG TGGTCATCTCTCCAGTCTAAGG
il10 TTAAAGCACTCCACAACCC AGTACCTCTTGCATTTCACC
il1b ATCAAACCCCAATCCACAGAGT GGCACTGAATCCACCACGTT
ikbaa CCTGCGTTCCATTCTCACCT GGCCACTACACTGCTCCTTT
nfkb1 CGCAAGTCCTACCCACAAGT ACCAGACTGTGAGCGTGAAG
nfkb2 CATATGTCCCACACAATCAAGAC AGCCACCATAATGATCTGGAA
Solutions
E3 (embryo medium) (see Recipes)
Tricaine (see Recipes)
75% Ethanol (see Recipes)
β-glucan stock (see Recipes)
β-glucan control stock (see Recipes)
β-glucan injection mix (see Recipes)
0.5% Phenol red (see Recipes)
3% Methylcellulose (see Recipes)
iScript reaction mix (see Recipes)
PerfeCTa SYBR Green FastMix with ROX mix (see Recipes)
Recipes
E3 (embryo medium) (1×)
Reagent Final concentration Quantity
CaCl2 0.33 mM 36.6 mg
MgSO4 0.33 mM 39.7 mg
KCl 0.17 mM 12.7 mg
NaCl 5 mM 292.2 mg
MilliQ water n/a Up to 1 L
Total n/a 1 L
Tricaine
Reagent Final concentration Quantity
Tricaine n/a 400 mg
1 M Tris (pH 9) 20 mM 2 mL
MilliQ water n/a 98 mL
Total n/a 100 mL
Note: Adjust to neutral pH by adding acid or base as appropriate until the solution is pH 7. Dilute in embryo medium to 4.2% (v/v).
75% Ethanol
Reagent Final concentration Quantity
Ethanol (absolute) 75% 37.5 mL
Ultrapure water n/a 12.5 mL
Total n/a 50 mL
β-glucan peptide stock
Reagent Final concentration Quantity
β-glucan peptide 5 mg/mL 5 mg
PBS n/a 1 mL
Total n/a 1 mL
β-glucan control stock
Reagent Final concentration Quantity
Control β-glucan (whole glucan peptides, soluble) 5 mg/mL 5 mg
PBS n/a 1 mL
Total n/a 1 mL
β-glucan injection mix (one experiment)
Reagent Final concentration Quantity
5 mg/mL β-glucan peptide (or control) 4 mg/mL 8 μL
0.5% Phenol red 0.1% 2 μL
Total n/a 10 μL
Note: Make this injection mix fresh for every experimental day. Discard unused mix.
0.5% Phenol red
Reagent Final concentration Quantity
Phenol red 0.5% 0.5 g
PBS n/a 100 mL
Total n/a 100 mL
Note: Adjust to neutral pH by adding acid or base as appropriate until the solution is pH 7.
3% methylcellulose
Reagent Final concentration Quantity
Methylcellulose 3% 30 g
E3 n/a Up to 1 L
Total n/a 1 L
Note: Shake on an orbital shaker at room temperature for up to one week to dissolve.
iScript cDNA synthesis reaction mix (1 reaction)
Reagent Final concentration Quantity
RNA n/a 500 ng (1–15 μL)
iScript 5× reaction buffer 1× 4 μL
iScript reverse transcriptase n/a 1 μL
Ultrapure water n/a To 20 μL
Total n/a 20 μL
PerfeCTa SYBR Green FastMix with ROX mix (one reaction)
Reagent Final concentration Quantity
cDNA (1:5 diluted) n/a 1 μL
Forward primer (10 μM) 300 nM 0.3 μL
Reverse primer (10 μM) 300 nM 0.3 μL
2× FastMix solution 1× 5 μL
Ultrapure water n/a 2.4 μL
Total n/a 10 μL
Laboratory supplies
Thin wall borosilicate capillary tubes (Warner Instruments, catalog number: 64-0778)
Small Petri dish (60 mm × 15 mm) (Corning, catalog number: 430166)
Petri dish (150 mm × 25 mm) (Corning, catalog number: 430599)
1.5 mL tube (Axygen, catalog number: MCT-175-C)
0.2 mL PCR tube (Axygen, catalog number: PCR-0208-CP-C)
Dumont #55 forceps (Fine Science Tools, catalog number: 11295-51)
3 mL transfer pipettes (Biologix, catalog number: 30-0138)
384-well plate (Applied Biosystems, catalog number: 4309849)
Microloader pipette tips (0.5–20 μL) (Eppendorf, catalog number: EP5242956003)
1 mL Luer-Lok syringe (BD Biosciences, catalog number: 309628)
27 G needle (0.5 inch) (BD Biosciences, catalog number: 301801)
Equipment
Centrifuge, 2 mL tube rotor (Eppendorf, catalog number: EPP2232000060)
Incubator (set to 28 °C) (Sanyo, catalog number: MIR-162)
Microscope stage micrometre (0.01 mm) (ProSciTech, catalog number: S81K)
Stereomicroscope (Nikon, model: SMZ1500)
Micropipette puller (Sutter Instruments Co., flaming/brown puller set to: heat 680, pull 75, velocity 40, time 55, pressure 530, to produce tapered needles)
Micromanipulator (World Precision Instruments, catalog number: M3301R)
Quantstudio 6 Flex Real-Time PCR System (Thermo Fisher Scientific)
Mastercycler Nexus Thermal Cycler Eco (Eppendorf)
Nanodrop (Thermo Scientific, catalog number: 13-400-518)
Software and datasets
Excel (Microsoft)
Prism v9 (GraphPad)
Procedure
Microinjection of zebrafish larvae
Obtain appropriately staged zebrafish.
Collect eggs by natural spawning from 3–5 pairs of zebrafish. Keep clutches from different breeding pairs separate to enable the collection of data from biological replicates. Collect 30–50 eggs per breeding pair.
Place freshly laid eggs into a clean 150 mm × 25 mm Petri dish with E3 medium.
Observe freshly fertilised eggs under a stereomicroscope. Remove dead eggs (black coagulated material within a chorion) and any other debris (fish waste, fish scales) from the Petri dish.
Place eggs into a 28 °C incubator and leave overnight. Zebrafish were raised in a dark incubator throughout this study.
At 24 hours post fertilisation (hpf), observe embryos under a stereomicroscope and remove dead or underdeveloped embryos. Refer to a zebrafish development chart to assess the developmental stage, such as Kimmel et al. (1995). If transparency is required, supplement embryo medium with 0.003% PTU.
Place embryos in a 28 °C incubator and raise until they are 2.25 dpf/54 hpf.
If unhatched, remove the chorion. Use sharp forceps to puncture and split the chorion open and remove the embryo.
Deliver β-glucan into embryos.
Note: This protocol describes a “double injection” method where embryos/larvae are injected twice with β-glucan, once at 2.25 dpf (54 hpf) and again at 3 dpf (72 hpf), as two injections induce a stronger response than a single injection, as described in Darroch et al. (2022).
Anaesthetise 15–20 embryos at 2.25 dpf (54 hpf) per biological replicate by supplementing embryo medium with 4.2% (v/v) tricaine. Wait for 20–30 s.
Gently touch embryos with a transfer pipette. If they do not twitch or swim away, they are sufficiently anaesthetised.
Pour a small volume (roughly 1 mL) of 3% methylcellulose into a 60 mm × 15 mm Petri dish. Using a microloader pipette tip that has been shortened to 2 cm total length, move the methylcellulose around the Petri dish until it covers the dish in a thin, even layer.
Pool embryos into the middle of the Petri dish by swirling the plate. Pick up embryos using a transfer pipette. Hold the transfer pipette vertical and wait for the embryos to settle at the tip. In one or two drops, place the embryos onto one side of the methylcellulose-coated Petri dish.
Using the shortened microloader tip, array the embryos one by one in a straight line within the Petri dish. Array the embryos laterally with the hindbrain facing right (Figure 1A, 1B, and 1D).
Figure 1. Microinjection of β-glucan into the circulation of zebrafish embryos. (A) Photo showing injection setup. (B) Photo showing embryos arrayed in a Petri dish with microinjection needle in position. (C) Image of 0.01 mm microscope stage slide micrometre. The black circle indicates the size of injection bolus that will deliver 1 nL of solution. (D) Image of 2.25 dpf (54 hpf) embryos arrayed in 3% methylcellulose. The loaded microinjection needle approaches from the right to inject embryos sequentially. (E) Image shows individual embryo with needle positioned for microinjection. The white arrowhead indicates the tip of the needle in the embryo at the correct injection position. Black dashed box is magnified view. In magnified view, the black asterisk indicates the position where the needle enters the yolk sac prior to the injection site. The white arrowhead indicates where the needle can be seen to press against the pericardial wall, which means the needle is deep enough to inject into the circulation. (E′) Image shows the same embryo in (E) immediately following microinjection. The white arrowhead indicates the tip of the needle in the embryo. The black asterisk highlights the red colour of the phenol red dye, indicating successful injection. Black dashed box is magnified view. Scale bars 500 μm in (D), 50 μm in (E, E′), and 40 μm in (E, E′ magnified views).
To inject β-glucan, load the microinjection needle (hereafter referred to as needle) with 3 μL of 4 mg/mL β-glucan peptide solution. If injecting the control, load the needle with 3 μL of 4 mg/mL β-glucan control solution.
Under the stereomicroscope, cut the tip of the needle with sharp forceps. The injection mix will move to the tip of the needle after it has been cut.
Calibrate the injection bolus to 1 nL. Add one drop of mineral oil to a 0.01 mm microscope stage slide micrometre and, under a stereomicroscope, focus on the ruler. Adjust the pressure and pulse duration of the injection box until the injection bolus measures 1.2 units (Figure 1C). This is 1 nL.
Inject 1 nL (4 ng) of β-glucan into the circulation of embryos (Figure 1D and 1E). Approaching from the right, pierce through the yolk sac of each embryo and move the needle towards the pericardial cavity until you see the needle touch the pericardial wall. Once in position, inject. The red dye in the injection mix indicates successful injection, as the dye gets taken up into circulation as the heart beats (Figure 1E′).
Note: Do not pierce through the epidermal wall.
See Troubleshooting.
Pour enough E3 medium into the injection Petri dish to completely cover the embryos. Using a transfer pipette, suck up and eject E3 repeatedly over the embryos until they dislodge from the methylcellulose. Gently pick up the injected embryos with a transfer pipette and place into a new 150 mm × 25 mm Petri dish with fresh E3 medium.
Place the Petri dish into a 28 °C incubator and leave overnight.
At 3 dpf (72 hpf), repeat steps 2a–2k to deliver a second dose of β-glucan into the larvae. This second dose is 8 ng, so inject 2 nL into the larvae by injecting 1 nL twice in rapid succession.
Analysis of gene expression post injection
Extract total RNA from injected larvae.
At the desired time post injection, anaesthetise pools of 15–20 larvae per biological replicate by supplementing the embryo medium with 4.2% (v/v) tricaine. We investigated gene expression at 1, 4, and 11 days post injection (dpi) with three biological replicates per time point (Figure 2A).
Transfer pooled larvae into 1.5 mL microcentrifuge tubes.
Remove as much E3 as possible using a transfer pipette. A small volume (< 100 μL) of residual E3 will not disrupt downstream processes.
Pipette 1 mL of TRIzol reagent into the 1.5 mL tubes.
Caution: TRIzol is hazardous and should be handled in a fume hood.
Connect a 27-gauge needle to a sterile 1 mL syringe. Carefully aspirate the larvae into the syringe and dispense back into the 1.5 mL tube. Repeat until the larval tissue is homogenised.
Incubate for 5 min at room temperature.
Pause point: Store samples at -80 °C or continue immediately.
Add 200 μL of chloroform and vortex twice for 5 s.
Incubate for 3 min at room temperature.
Centrifuge at 12,000× g for 15 min at 4 °C.
Observe how the sample has split into three layers with a clear upper layer (phase), middle white interphase, and lower pink phase. Without touching the white or pink phases, carefully remove the upper aqueous phase and transfer into a new 1.5 mL microcentrifuge tube.
Add 500 μL of 100% isopropanol and vortex for 1 min.
Incubate for 10 min at room temperature.
Centrifuge at 12,000× g for 10 min at 4 °C.
Remove the supernatant by pouring and observe the white pellet.
Add 1 mL of 75% ethanol onto the pellet.
Spin at 7,500× g for 5 min at 4 °C.
Remove all ethanol by pipetting and allow the pellet to air dry for 5 min at room temperature.
Resuspend the pellet in 30 μL of ultrapure water. Keep on ice.
Quantify RNA concentration by nanodrop.
Pause point: Store RNA at -80 °C or continue immediately.
Real-time quantitative PCR (RT qPCR) expression analysis of genes of interest.
Reverse transcribe cDNA using the iScript cDNA Synthesis kit. In a 0.2 mL tube, prepare the iScript cDNA synthesis reaction mix (Recipe 9).
Place samples into an Eppendorf Mastercycler Nexus Thermal Cycler Eco and run the following protocol: 25 °C for 5 min, 46 °C for 20 min, 95 °C for 1 min.
Remove from the thermal cycler and place on ice.
Pause point: Store cDNA at -80 °C or continue immediately.
Add 80 μL of ultrapure water to the 20 μL cDNA reaction to dilute the cDNA 1:5.
For the qPCR reaction, prepare the PerfeCTa SYBR Green FastMix with ROX mix (Recipe 10).
Pipette into a 384-well plate with three technical replicates for each sample per gene.
Place the reaction into a Quantstudio 6 Flex Real-Time PCR System and run the following cycling parameters: initial denaturation 95 °C for 2 min, followed by 40 cycles at 95 °C for 15 s, 60 °C for 30 s.
Record the Ct values of each gene and proceed to data analysis (Figure 2B).
Figure 2. Gene expression analysis following β-glucan injection. (A) Schematic illustrating the injection protocol followed by time points of sample collection for subsequent RT qPCR. (B) Fold change of relevant immune genes at 1, 4, and 11 days post injection (dpi) with β-glucan relative to control. Data shown is from averaged ΔCt values from three biological replicates, n = 15 larvae per biological replicate. Endogenous control gene was ef1a.
Data analysis
Microsoft Excel was used to analyse the data with ef1a as the endogenous control (reference) gene. Briefly, the ΔCt value was calculated by subtracting the average Ct value of the reference gene from the average Ct value of the gene of interest (Ctaverage gene of interest – Ctaverage ef1a) (Livak and Schmittgen, 2001). The ΔΔCt value was calculated by subtracting the ΔCt value of the average ΔCt of the control samples from the average ΔCt of the experimental samples (i.e., β-glucan injected) (ΔCtaverage β-glucan injected – ΔCtaverage control). Gene-expression fold change was determined by the 2-ΔΔCT method (Livak and Schmittgen, 2001). Data was visualised using GraphPad Prism version 8.
Validation of protocol
Previous studies showed that the NF-kB pathway is involved in the response to β-glucan, and the expression of specific inflammatory cytokines such as il1b, tnfa, cxcl8, and il10 are upregulated following stimulation (Rogers et al., 2005; Inoue and Shinohara, 2014; Megías et al., 2016). The RT qPCR data presented in this current study shows that two predicted genes of interest were upregulated following β-glucan injection when compared to control, and this analysis was further characterised in the recent publication (Darroch et al., 2022). Darroch et al. (2022) utilised this method to stimulate a trained-like phenotype in larval zebrafish that protected larvae from subsequent bacterial challenge and increased neutrophil recruitment to infection.
General notes and troubleshooting
General notes
Larval zebrafish offer a unique opportunity to investigate trained innate responses entirely in vivo. On a whole animal scale, post-training phenotypes such as larval survival and pathogen clearance can be assessed following live infection as demonstrated in Darroch et al. (2022). Such analyses are enhanced with the use of fluorescent pathogens that can be visualised within transparent larvae using fluorescent microscopy. A wide range of pathogens or sterile insults can be used to challenge larvae post training to assess changes in innate immune function (Linnerz and Hall, 2020). In addition, there are numerous transgenic reporter lines available with fluorescently marked innate immune cells that can be directly observed in real time, at single-cell resolution (Astin et al., 2017; Linnerz and Hall, 2020). Analysis of phagocyte functions such as bacterial killing capacity, recruitment to infection/inflammation, or production of anti-microbial molecules (e.g., reactive intermediate species, as detected by fluorescent probes) can be assessed post training (Astin et al., 2017, Darroch et al., 2022).
Troubleshooting
Needle blocking: as the β-glucan is insoluble, it can block the needle. If the needle is blocked, increase the air pressure of the injector, change the mode of the injector to continuous airflow, and attempt to flush the β-glucan clumps out of the needle. If this does not work, you can cut the needle slightly larger and try to flush it again. You should not make the needle larger indefinitely; when the bore of the needle gets to a size that damages the larvae, you will need to load and cut a new needle.
Acknowledgments
This work was supported by grants awarded to CJH from Marsden Fund, Royal Society of New Zealand (grant # 3706135) and Health Research Council of New Zealand (grant # 17/294). This protocol was initially described and validated in Darroch et al. (2022).
Competing interests
The authors declare that they have no competing interests.
Ethical considerations
All zebrafish research was conducted with the approval of the University of Auckland Animal Ethics Committee (approval numbers AEC001911 and AEC22563).
References
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Detailed Protocol to Perform Direct PCR Using Filamentous Fungal Biomass—Tips and Considerations
HJ Hosung Jeon
HS Hokyoung Son
KM Kyunghun Min
Published: Vol 13, Iss 21, Nov 5, 2023
DOI: 10.21769/BioProtoc.4889 Views: 1281
Reviewed by: Shweta PanchalXiaofei LiangYueqiang Leng
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Original Research Article:
The authors used this protocol in Frontiers in Plant Science Jan 2023
Abstract
The precise and rapid detection of fungi is important in various fields, including clinics, industry, and agriculture. While sequencing universal DNA barcodes remains the standard method for species identification and phylogenetic analysis, a significant bottleneck has been the labor-intensive and time-consuming sample preparation for genomic DNA extraction. To address this, we developed a direct PCR method that bypasses the DNA extraction steps, facilitating efficient target DNA amplification. Instead of extracting genomic DNA from fungal mycelium, our method involves adding a small quantity of mycelium directly to the PCR mixture, followed by a heat shock and vortexing. We found these simple adjustments to be sufficient to lyse many filamentous fungal cells, enabling target DNA amplification. This paper presents a comprehensive protocol for executing direct PCR in filamentous fungi. Beyond species identification, this direct PCR approach holds promise for diverse applications, such as diagnostic PCR for genotype screening without fungal DNA extraction. We anticipate that direct PCR will expedite research on filamentous fungi and diagnosis of fungal diseases.
Key features
• Eliminates the time-consuming genomic DNA extraction step for PCR, enhancing the speed of molecular identification.
• Adds a small quantity of mycelium directly into the PCR mix.
• Emphasizes the crucial role of heat shock and vortexing in achieving efficient target DNA amplification.
• Accelerates the molecular identification of filamentous fungi and rapid diagnosis of fungal diseases.
Graphical overview
Direct PCR using filamentous fungal biomass
Keywords: Direct PCR Filamentous fungi Molecular identification ITS region
Background
Polymerase chain reaction (PCR) is a powerful tool widely used to amplify specific target regions of DNA. Since its invention in the mid-1980s [1, 2], PCR has become a fundamental technique employed in various fields. A notable application of PCR is in the diagnosis of pathogenic diseases. It allows for the amplification of universal DNA barcodes, facilitating the identification of causative pathogens [3, 4]. Given that multiple copies of DNA are generated from the DNA template, the preparation of this template is essential for performing PCR. For filamentous fungi, the steps for DNA extraction and purification have been prerequisites for PCR amplification. Most fungi possess a rigid cell wall, predominantly constituted of polysaccharides like chitin and glucans [5, 6]. This results in inefficient fungal DNA extraction. The procedure for extracting DNA from fungal sources requires laborious and time-consuming steps involving the grinding of freeze-dried mycelia [7]. Additionally, the fungal DNA extraction process involves multiple stages using toxic chemicals like phenol and chloroform. A technique known as colony PCR involves directly adding cells into the PCR mixture, and it has found widespread application in bacterial and yeast research [8]. In the case of filamentous fungi, previous studies have suggested various PCR methods to minimize fungal DNA extraction steps. The methods proposed by Alshahni et al. and Walch et al. eliminated the need for fungal genomic DNA extraction [9, 10]. However, these methods incorporated additional steps, such as using lysis buffer or bovine serum albumin (BSA), to lyse fungal cell walls. Despite these advancements, the adoption of these methods for filamentous fungi has been limited due to their reduced PCR efficiency and the need for a cell lysis process.
Here, we introduce a direct PCR procedure tailored for filamentous fungi. This approach eliminates the need for the fungal DNA extraction process and the use of additional reagents such as BSA or proteinase. Our method involves adding a small amount of mycelium directly into the PCR mixture, followed by heat shock and vortexing. These heat shock and vortexing steps are key in breaking the fungal cell walls and membrane, releasing the genomic DNA for PCR amplification. This method will expedite the molecular identification of filamentous fungi and facilitate rapid diagnosis of fungal diseases.
Materials and reagents
Potato dextrose broth (Difco, catalog number: 254920)
Agar powder (Duksan, catalog number: 601)
AccuPower® Taq PCR Premix (Bioneer, catalog number: 20-K-2602)
Nuclease-free water (Sigma-Aldrich, catalog number: 7732-18-5)
Primers:
ITS4: 5′-TCCTCCGCTTATTGATATGC-3′
ITS5: 5′- GGAAGTAAAAGTCGTAACAAGG-3′
EF1T: 5′-ATGGGTAAGGAGGACAAGAC-3′
EF2T: 5′-GGAAGTACCAGTGATCATGTT-3′
SeaKem® LE agarose (Lonza, catalog number: 50004)
RedSafe nucleic acid staining solution (iNtRON, catalog number: 21141)
Tris ultrapure (Duchefa Biochemie, catalog number: T1501.1000)
Ethylenediaminetetraacetic acid (EDTA) disodium salt dihydrate (Sigma-Aldrich, catalog number: E5134-50G)
Glacial acetic acid (Sigma-Aldrich, catalog number: PHR1748)
6× Loading buffer (Takara, catalog number: 9156)
100 bp Plus DNA ladder (Bioneer, catalog number: D-1035)
Ethanol EMSURE ACS, ISO, Reag. PH Eur (1 L) (Merck Millipore, catalog number: 1.0098.1011)
Solutions
Culture medium (PDA) (see Recipes)
50× TAE buffer (1 L) (see Recipes)
Recipes
Culture medium (PDA)
Reagent Final concentration Quantity
Potato dextrose broth 24 g/L 24 g
Agar powder 10 g/L 10 g
H2O n/a up to 1 L
50× TAE buffer (1 L)
*Note: Add Tris ultrapure and EDTA disodium salt dihydrate to approximately 700 mL of H2O and stir until dissolved. Carefully add the acetic acid and adjust the volume to 1 L.
Reagent Final concentration Quantity
Tris ultrapure 2 M 242 g
Glacial acetic acid 1 M 57.1 mL
EDTA disodium salt dihydrate 50 mM 18.61 g
H2O n/a up to 1 L
Laboratory supplies
Pipette tips (1,000, 200, 10 μL), DNase, RNase, DNA, & endotoxin free (Neptune)
Petri dish (90 mm × 15 mm) (SPL, catalog number 10090)
1.5 mL microtubes (Axygen, catalog number: MCT-15-C)
MEGAquick-spinTM Plus Total Fragment DNA Purification kit (iNtRON, catalog number: 17290)
Equipment
Pipettes (0.5–10 μL, 2–20 μL, 20–200 μL, 100–1,000 μL) (Eppendorf, model: Research Plus®)
Microcentrifuges mini (Labogene, catalog number: LZ-1312)
Vortex Genie 2 (Scientific Industries, catalog number: SI-0256)
MiniAmpTM thermal cycler (Applied Biosystems, catalog number: A37834)
Gel tray L (Takara, catalog number: AD210)
PowerPacTM basic power supply (Bio-Rad, catalog number: 1645050)
Gel imaging system MaXidoc G2 (DAIHAN Scientific, catalog number: DH.WGD00300)
NanoDrop 2000 (Thermo Scientific, catalog number: ND-2000)
Procedure
Preparation of fungal culture
Transfer an agar plug containing fungal mycelia onto a PDA plate.
Incubate the agar plate for five days at 25 °C.
Direct PCR amplification
For a 20 μL reaction volume, add 0.5 μL of each primer (20 μM) to AccuPower® Taq PCR Premix tubes.
To dissolve the vacuum-dried premixture, add 19 μL of nuclease-free water into the PCR tubes.
Gently scratch the fungal culture using a pipette tip to collect a small amount of mycelium (Figure 1).
Figure 1. Mycelium collection for direct PCR. Aerial hyphae of Fusarium graminearum were collected using a pipette and 10 μL pipette tips. The distal part of the pipette tip is magnified for clarity. Scale bar = 500 μm.
Transfer the collected mycelium to the reaction mixture. Vigorously vortex the PCR tubes and then briefly centrifuge to bring down the contents.
For the heat shock reaction, place the PCR tubes in a thermal cycler and set the program (3 min at 95 °C, then cool down to 25 °C).
After the program, immediately transfer the PCR tubes to an ice bath for 1 min.
Vortex the samples vigorously for 15 s and then chill them on ice for an additional 15 s.
Repeat step B7 two more times.
Centrifuge the reaction mixture briefly to collect all contents at the bottom and then return the PCR tubes to the thermal cycler.
Set the thermal cycler with the following program:
Initial denaturation at 95 °C for 1 min.
Denaturation at 95 °C for 30 s.
Annealing at 56 °C (adjust temperature based on primer specifications) for 1 min.
Extension at 72 °C (adjust time based on the expected amplicon size) for 1 min.
Repeat steps b–d for 40 cycles (based on our results, 40 cycles were more optimal than 30).
Final extension at 72 °C for 3 min.
Hold at 4 °C for 5 min.
Gel electrophoresis
Gel preparation:
Weigh 2.25 g of agarose powder.
Dissolve agarose powder in 150 mL of 1× TAE buffer (see Recipes) completely using a microwave (avoid overboiling) to make a 1.5 % agarose gel (w/v).
Add 7.5 μL of RedSafe nucleic acid staining solution and mix thoroughly.
Pour the dissolved agarose solution into a gel tray with a well comb.
Allow the poured gel to solidify fully at room temperature for 20–30 min.
Loading samples and running:
Add 6× loading buffer to each PCR product.
Place the solidified agarose gel into the electrophoresis unit and submerge it entirely in 1× TAE buffer.
Carefully load your samples and 100 bp Plus DNA ladder into the gel wells.
Operate PowerPacTM basic power supply at 100–120 V for 30–40 min (adjust the voltage and running time depending on gel concentration and PCR product size).
After gel running, carefully remove the gel from the unit and place it under a UV light device to visualize DNA fragments.
Purification and sequencing
To purify the DNA samples, use MEGAquick-spinTM Plus Total Fragment DNA Purification kit and follow the protocol provided with the kit.
Transfer 50 μL of your PCR product into a 1.5 mL microcentrifuge tube and add 250 μL of agarose gel lysis buffer (included in the DNA purification kit). Mix well by vortexing.
Insert a spin column in a collection tube.
Transfer the sample mixture to the spin column and centrifuge at 11,000× g for 30 s.
Discard the flowthrough.
Add 750 μL of the ethanol-added wash buffer solution (included in the kit) to the spin column and then centrifuge at 11,000× g for 30 s.
Discard the flowthrough.
Centrifuge again at 13,500× g for 3 min to dry the column matrix.
Place the spin column in a new 1.5 mL microcentrifuge tube.
Add 20–40 μL of elution buffer (included in the kit) to the membrane center of the spin column and wait for 3 min.
Centrifuge at 13,500× g for 1 min to elute the DNA.
Measure DNA concentration using a NanoDrop 2000 and submit the samples to a sequencing service center.
Data analysis
DNA sequences were used to conduct a BLAST search against the NCBI GenBank database for species identification.
This protocol or parts of it has been used and validated in the following research article:
Jeon et al. (2023) [4]. Application of direct PCR for phylogenetic analysis of Fusarium fujikuroi species complex isolated from rice seeds. Frontiers in Plant Science (Figures 1, 2, and 3).
General notes and troubleshooting
The amount of mycelium added can have a significant effect on the PCR outcome. Always ensure that only a small quantity of mycelium is added to the PCR mixture. Introducing too much mycelium may lead to reaction blockages and potential failure (Figure 2).
Figure 2. Influence of mycelium quantity on direct PCR efficiency. F. graminearum strain Z-3639 was subjected to amplification of the TEF-1α region using a direct PCR approach. The different quantities of mycelium affected PCR efficiency as follows: PC: positive control (using extracted genomic DNA), NC: negative control (no template control), S: small amount of mycelium (as shown in Figure 1), M: medium amount (twice that of S), and L: large amount (twice that of M).
While direct PCR can be performed using various PCR enzymes, we strongly recommend using the Taq PCR Premix. This premix contains all necessary components for the reaction (including polymerase, dNTP, reaction buffer, and loading dye), except for the template DNA and primers. Our observations show that Taq PCR Premix offers a notably higher success rate compared to other PCR enzymes. Furthermore, the presence of a loading dye in the Taq PCR Premix improves visualization of the added mycelium, allowing for easier verification of mycelium quantity.
In the previous study [4], direct PCR amplification of the ITS region was tested on samples from major fungal lineages including Ascomycota, Basidiomycota, and Mucoromycota. Our method displayed a remarkable PCR success rate of 92% (34 out of 37 fungal species). Nonetheless, the ITS region of three species—Mucor mucedo, Aspergillus niger, and Aspergillus brasiliensis—remained unamplified by direct PCR. Notably, these fungi exhibited highly hydrophobic mycelium and spores, which likely inhibited cell lysis within the PCR mixture.
In addition to the ITS, the TEF-1α region and other genotype markers were successfully amplified using the direct PCR method to identify strains within the Fusarium fujikuroi species complex.
If the PCR efficiency is low, consider increasing the duration of vortexing after the heat shock step.
It is advisable to include a reaction containing isolated genomic DNA as a positive control in each experiment. Also, always have a no-template control. This approach can provide valuable insights when troubleshooting is necessary.
Acknowledgments
This work was supported by the National Research Foundation of Korea (2022R1I1A1A01065138) and the Strategic Initiative for Microbiomes in Agriculture and Food and Crop Viruses and Pests Response Industry Technology Development Program funded by the Ministry of Agriculture, Food and Rural Affairs of Korea (MAFRA) (No. 321101-03). Cartoons in Graphical overview were created with BioRender.com. The protocol was adapted from the previous study by Jeon et al. (2023) [4].
Competing interests
The authors declare no competing interests.
References
Saiki, R. K., Scharf, S., Faloona, F., Mullis, K. B., Horn, G. T., Erlich, H. A. and Arnheim, N. (1985). Enzymatic Amplification of β-Globin Genomic Sequences and Restriction Site Analysis for Diagnosis of Sickle Cell Anemia. Science 230(4732): 1350–1354. doi: 10.1126/science.2999980
Saiki, R. K., Gelfand, D. H., Stoffel, S., Scharf, S. J., Higuchi, R., Horn, G. T., Mullis, K. B. and Erlich, H. A. (1988). Primer-Directed Enzymatic Amplification of DNA with a Thermostable DNA Polymerase. Science 239(4839): 487–491. doi: 10.1126/science.2448875
Schoch, C. L., Seifert, K. A., Huhndorf, S., Robert, V., Spouge, J. L., Levesque, C. A., Chen, W., Bolchacova, E., Voigt, K., Crous, P. W., et al. (2012). Nuclear ribosomal internal transcribed spacer (ITS) region as a universal DNA barcode marker for Fungi. Proc. Natl. Acad. Sci. U. S. A. 109(16): 6241–6246. doi: 10.1073/pnas.1117018109
Jeon, H., Kim, J. E., Yang, J. W., Son, H. and Min, K. (2023). Application of direct PCR for phylogenetic analysis of Fusarium fujikuroi species complex isolated from rice seeds. Front. Plant Sci. 13: e1093688. doi: 10.3389/fpls.2022.1093688
Bowman, S. M. and Free, S. J. (2006). The structure and synthesis of the fungal cell wall. BioEssays 28(8): 799–808. doi: 10.1002/bies.20441
Gow, N. A. R., Latge, J. P. and Munro, C. A. (2017). The Fungal Cell Wall: Structure, Biosynthesis, and Function. Microbiol. Spectrum 5(3): efunk-0035-2016. doi: 10.1128/microbiolspec.funk-0035-2016
Leslie, J. F. and Summerell, B. A. The Fusarium Laboratory Manual. Ames, IW, USA: Blackwell Publishing. doi: 10.1002/9780470278376
Ling, M., Merante, F. and Robinson, B. H. (1995). A rapid and reliable DNA preparation method for screening a large number of yeast clones by polymerase chain reaction. Nucleic Acids Res. 23(23): 4924–4925. doi: 10.1093/nar/23.23.4924
AlShahni, M. M., Makimura, K., Yamada, T., Satoh, K., Ishihara, Y., Takatori, K. and Sawada, T. (2009). Direct Colony PCR of Several Medically Important Fungi using Ampdirect® Plus. Jpn. J. Infect. Dis. 62(2): 164–167. doi: 10.7883/yoken.jjid.2009.164
Walch, G., Knapp, M., Rainer, G. and Peintner, U. (2016). Colony-PCR Is a Rapid Method for DNA Amplification of Hyphomycetes. J. Fungi 2(2): 12. doi: 10.3390/jof2020012
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Molecular Biology > DNA > PCR
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Peer-reviewed
Biochemical Reconstitution of Ca2+-Dependent Exosome Secretion in Permeabilized Mammalian Cells
JN Jordan M. Ngo
JW Justin K. Williams
IL Isabelle M. Lehman
RS Randy Schekman
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4890 Views: 887
Reviewed by: Willy R Carrasquel-UrsulaezJulie A Dougherty Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in eLIFE May 2023
Abstract
Exosomes are a subpopulation of the heterogenous pool of extracellular vesicles that are secreted to the extracellular space. Exosomes have been purported to play a role in intercellular communication and have demonstrated utility as biomarkers for a variety of diseases. Despite broad interest in exosome biology, the conditions that regulate their secretion are incompletely understood. The goal of this procedure is to biochemically reconstitute exosome secretion in Streptolysin O (SLO)-permeabilized mammalian cells. This protocol describes the reconstitution of lyophilized SLO, preparation of cytosol and SLO-permeabilized cells, assembly of the biochemical reconstitution reaction, and quantification of exosome secretion using a sensitive luminescence-based assay. This biochemical reconstitution reaction can be utilized to characterize the molecular mechanisms by which different gene products regulate exosome secretion.
Key features
• This protocol establishes a functional in vitro system to reconstitute exosome secretion in permeabilized mammalian cells upon addition of cytosol, ATP, GTP, and calcium (Ca2+).
Graphical overview
Schematic overview of the exosome secretion biochemical reconstitution protocol. Streptolysin O (SLO) is prepared as described in Procedure A. Cytosol is isolated from HCT116 WT cells as described in Procedure B. HCT116 CD63-Nluc cells are permeabilized by SLO as detailed in Procedure C. The assembly of the exosome secretion reactions are described in Procedure D. Quantification of CD63-Nluc secretion is detailed in Procedure E (Modified from Williams et al., 2023).
Keywords: Extracellular vesicle Exosome Reconstitution Streptolysin O Luciferase
Background
Extracellular vesicles (EVs) encompass a diverse pool of membrane-enclosed compartments released by cells to the extracellular space (Colombo et al., 2014). Exosomes are an EV subpopulation that is secreted upon fusion of multivesicular bodies (MVBs) at the cell surface and they have been suggested to play a role in intercellular communication in both physiological and disease states (Harding et al., 1983; Fong et al., 2015; Hsu et al., 2017). Of particular interest is the selective and likely tissue-specific protein and small RNA composition of exosomes, which offers the prospect of their use as biomarkers for disease progression (Shurtleff et al., 2017; Driedonks and Nolte-’t Hoen, 2019; Upton et al., 2021). Despite an accumulating interest in exosomes, the molecular mechanisms that regulate and execute their secretion are not well understood. A recent study inserted nanoluciferase (Nluc) into the endogenous locus of the exosome marker protein CD63 to allow simple quantification of exosome secretion (Hikita et al., 2018). We modified this assay by the addition of a membrane-impermeable Nluc inhibitor to allow a distinction between cellular debris and bona fide CD63-positive EVs (exosomes) (Walker et al., 2017; Williams et al., 2023). Using this assay, we demonstrated that MVBs participate in Ca2+-dependent plasma membrane repair. We then leveraged this modified CD63-Nluc assay to develop a biochemical reaction to reconstitute exosome secretion using cells that have been permeabilized by the bacterial pore-forming toxin, Streptolysin O (SLO). Using this biochemical reconstitution assay, we demonstrated that Annexin A6 is required for Ca2+-dependent exosome secretion during plasma membrane repair.
Materials and reagents
Biological materials
HCT116 WT cells
HCT116 CD63-Nluc cells
Note: Other cell lines expressing a functional and traffic-competent Nluc-CD63 fusion protein may be compatible for this reconstitution assay.
Reagents
DMEM, high glucose, GlutaMAXTM supplement (Thermo Fisher Scientific, GibcoTM, catalog number: 10566016)
Fetal bovine serum (FBS) (VWR, catalog number: 89510-194)
Phosphate-buffered saline (PBS), pH 7.4 (Thermo Fisher Scientific, catalog number: 10010023)
Liquid nitrogen
Protein assay dye reagent concentrate (Bio-Rad, catalog number: 5000006)
D-sorbitol (Sigma-Aldrich, catalog number: S1876-5KG)
HEPES (Sigma-Aldrich, catalog number: RDD002-1KG)
Potassium chloride (KCl) (Fisher Scientific, catalog number: P217-500)
Sodium chloride (NaCl) (Fisher Scientific, catalog number: S271-3)
Magnesium chloride hexahydrate [MgCl2·(H2O)6] (Fisher Scientific, catalog number: BP214-500)
Potassium phosphate monobasic (KH2PO4) (Fisher Scientific, catalog number: P285-500)
Potassium acetate (CH3COOK) (Fisher Scientific, catalog number: BP364-500)
Dithiothreitol (DTT) (GoldBio, catalog number: DTT25)
[Ethylenebis-(oxyethylenenitrilo)]-tetraacetic acid (EGTA) (Fisher Scientific, catalog number: O2783-100)
Calcium chloride dihydrate [CaCl2·(H2O)2] (EMD Millipore, catalog number: CX0130-1)
Adenosine-5′-triphophate (ATP) (GE Healthcare, catalog number: 27-1006-01)
Creatine phosphate disodium salt (Sigma-Aldrich, catalog number: 2380-25GM)
Creatine phosphokinase (Roche Diagnostics, catalog number: 10127566001)
GTP 100 mM lithium salt (Roche Diagnostics, catalog number: 11140957001)
Triton® X-100 (TX-100) (Sigma-Aldrich, catalog number: X100-500ML)
Streptolysin O (SLO) (Sigma-Aldrich, catalog number: SAE0089-100KU)
Intracellular TE Nano-Glo® substrate/inhibitor (Promega, catalog number: N2160)
Extracellular NanoLuc® inhibitor
NanoBRETTM Nano-Glo® substrate
Nano-Glo® Luciferase Assay System (Promega, catalog number: N1120)
Nano-Glo® Luciferase assay substrate
Nano-Glo® Luciferase assay buffer
Solutions
1 M HEPES, pH 7.4 (see Recipes)
1 M KCl (see Recipes)
1 M NaCl (see Recipes)
1 M MgCl2 (see Recipes)
1 M KH2PO4 (see Recipes)
1 M EGTA, pH 7.4 (see Recipes)
1 M DTT (see Recipes)
100 mM CaCl2 (see Recipes)
10% TX-100 (see Recipes)
10× ATP regeneration system (ATPr) (see Recipes)
10 mM GTP (see Recipes)
1,000× protease inhibitor cocktail (see Recipes)
PBS + 10 mM DTT (see Recipes)
PBS + protease inhibitors (see Recipes)
Hypotonic lysis buffer (see Recipes)
PBS + 1 mM EGTA (see Recipes)
Transport buffer (see Recipes)
SLO binding buffer (see Recipes)
Permeabilization buffer (see Recipes)
High-salt transport buffer (see Recipes)
PBS + 2% TX-100 (see Recipes)
Nluc substrate/inhibitor master mix (see Recipes)
Nluc lytic master mix (see Recipes)
Recipes
1 M HEPES, pH 7.4 (250 mL)
Note: First add 200 mL of ddH2O, adjust pH to 7.4 with 10 N NaOH, then add ddH2O up to 250 mL. Store at 4 °C.
Reagent Final concentration Quantity
HEPES 1 M 59.6 g
ddH2O n/a up to 250 mL
Total n/a 250 mL
1 M KCl (250 mL)
Note: Store at room temperature.
Reagent Final concentration Quantity
KCl 1 M 18.64 g
ddH2O n/a up to 250 mL
Total n/a 250 mL
1 M NaCl (250 mL)
Note: Store at room temperature.
Reagent Final concentration Quantity
NaCl 1 M 14.61 g
ddH2O n/a up to 250 mL
Total n/a 250 mL
1 M MgCl2 (250 mL)
Note: Store at room temperature.
Reagent Final concentration Quantity
MgCl2·(H2O)6 1 M 50.83 g
ddH2O n/a up to 250 mL
Total n/a 250 mL
1 M KH2PO4 (250 mL)
Note: Store at room temperature.
Reagent Final concentration Quantity
KH2PO4 1 M 34.02 g
ddH2O n/a up to 250 mL
Total n/a 250 mL
1 M EGTA, pH 7.4 (50 mL)
Note: First add 35 mL of ddH2O, adjust pH to 7.4 with solid NaOH, and then add ddH2O up to 50 mL.
Reagent Final concentration Quantity
EGTA 1 M 19.02 g
ddH2O n/a up to 50 mL
Total n/a 50 mL
1 M DTT (10 mL)
Note: Make 1 mL aliquots and store at -20 °C.
Reagent Final concentration Quantity
DTT 1 M 1.54 g
ddH2O n/a up to 10 mL
Total n/a 10 mL
100 mM CaCl2 (50 mL)
Note: Store at room temperature.
Reagent Final concentration Quantity
CaCl2·(H2O)2 100 mM 0.735 g
ddH2O n/a up to 50 mL
Total n/a 50 mL
10% TX-100 (50 mL)
Note: Mix end-over-end to resuspend thoroughly. Store at room temperature.
Reagent Final concentration Quantity
TX-100 10% 5 mL
ddH2O n/a up to 50 mL
Total n/a 50 mL
10× ATP regeneration system (ATPr) (20 mL)
Note: Make 100 μL aliquots, snap freeze in liquid nitrogen, and store at -80 °C.
Reagent Final concentration Quantity
Creatine phosphate disodium salt 400 mM 2.04 g
Creatine phosphokinase 2 mg/mL 40 mg
ATP 10 mM 101.44 mg
Transport buffer n/a up to 20 mL
Total n/a 20 mL
10 mM GTP (4 mL)
Note: Make 25 μL aliquots, snap freeze in liquid nitrogen, and store at -80 °C.
Reagent Final concentration Quantity
100 mM GTP 10 mM 400 μL
Transport buffer n/a 3.6 mL
Total n/a 4 mL
1,000× protease inhibitor cocktail (10 mL)
Note: Make 100 μL aliquots and store at -20 °C.
Reagent Final concentration Quantity
4-aminobenzamidine dihydrochloride 1 M 2.08 g
Antipain dihydrochloride 1 mg/mL 10 mg
Aprotinin 1 mg/mL 10 mg
Leupeptin 1 mg/mL 10 mg
ddH2O n/a up to 10 mL
Total n/a 10 mL
PBS + 10 mM DTT (1 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
PBS n/a 990 μL
1 M DTT 10 mM 10 μL
Total n/a 1 mL
PBS + protease inhibitors (25 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
PBS n/a 25 mL
1,000× protease inhibitor cocktail 1× 25 μL
Total n/a 25 mL
Hypotonic lysis buffer (10 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
1 M HEPES, pH 7.4 20 mM 200 μL
1 M KCl 10 mM 100 μL
1 M EGTA, pH 7.4 1 mM 10 μL
1 M DTT 1 mM 10 μL
1,000× protease inhibitor cocktail 1× 10 μL
ddH2O n/a 9.67 mL
Total n/a 10 mL
PBS + 1 mM EGTA (10 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
PBS n/a 10 mL
1 M EGTA, pH 7.4 1 mM 10 μL
Total n/a 1 mL
Transport buffer (50 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
1 M HEPES, pH 7.4 20 mM 1 mL
D-sorbitol 250 mM 2.28 g
1 M KCl 120 mM 6 mL
1 M NaCl 10 mM 500 μL
1 M MgCl2 2 mM 100 μL
1 M KH2PO4 1.2 mM 60 μL
1 M EGTA, pH 7.4 1 mM 50 μL
1,000× protease inhibitor cocktail 1× 50 μL
ddH2O n/a up to 50 mL
Total n/a 50 mL
SLO binding buffer (1.5 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
Transport buffer n/a 1.5 mL
SLO 0.6 μg/mL 0.9 μg
Total n/a 1.5 mL
Permeabilization buffer (5 mL)
Note: Make fresh and store at 37 °C until use.
Reagent Final concentration Quantity
Transport buffer n/a 4.99 mL
1 M DTT 2 mM 10 μL
Total n/a 5 mL
High-salt transport buffer (5 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
Transport buffer n/a 5 mL
CH3COOK 1 M 0.49 g
Total n/a 5 mL
Transport buffer + 2% TX-100 (1 mL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
Transport Buffer n/a 800 μL
10% TX-100 2% 200 μL
1,000× protease inhibitor cocktail 1× 1 μL
Total n/a 1 mL
Nluc substrate/inhibitor master mix (750 μL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
PBS n/a 747 μL
Extracellular NanoLuc® inhibitor 1:1,000 dilution 0.75 μL
NanoBRETTM Nano-Glo® substrate 1:333 dilution 2.25 μL
Total n/a 750 μL
Nluc lytic master mix (300 μL)
Note: Make fresh and store at 4 °C until use.
Reagent Final concentration Quantity
Nano-Glo® Luciferase assay buffer n/a 294 μL
Nano-Glo® Luciferase assay substrate 1:50 dilution 6 μL
Total n/a 300 μL
Laboratory supplies
150 mm TC-treated cell culture dish (Corning, Falcon, catalog number: 08-772-6)
24-well BioCoatTM Poly-D-Lysine (PDL) coated plates (Corning, catalog number: 356414)
Amicon® Ultra 15 mL centrifugal filter unit with Ultracel-3k membrane (Merck, catalog number: UFC800324)
Ultra-ClearTM Tube (5 mL) 13 mm × 51 mm (Beckman Coulter, catalog number: 344057)
Axygen® 1.5 mL Maxymum Recovery® tube (Corning, catalog number: MCT-150-L-C)
Posi-click 1.7 mL microcentrifuge tube [Danville Scientific, catalog number: C2170(1001002)]
Cell scraper 25 cm (Sarstedt, catalog number: 83.1830) or equivalent
7 mL dounce homogenizer or equivalent
AcroPrep Advance 96-well 0.4 μm filter plate (Pall Corporation, catalog number: 8029)
96-well plate, non-treated surface, non-sterile (Fisher Scientific, catalog number: 12-565-226)
Equipment
SorvallTM ST16R centrifuge, TX-200 swinging bucket rotor, 400 mL round buckets, 4 × 50 mL, 9 × 15 mL conical adapters (Thermo Fisher Scientific, catalog number: 75818382)
Optima XE-90 ultracentrifuge (Beckman Coulter, catalog number: A94471)
SW 55 Ti swinging bucket rotor and bucket set (Beckman Coulter, catalog number: 342196)
Eppendorf 5430 R refrigerated centrifuge (Eppendorf, catalog number: 5428000015)
Eppendorf FA-45-30-11 fixed angle rotor (Eppendorf, catalog number: 05-401-503)
Eppendorf 5810 R refrigerated centrifuge with A-4-62 swinging bucket rotor and plate adaptor (Eppendorf, catalog number: 022627040)
Phase contrast microscope with 10× objective (any equivalent microscope works well)
Promega Glowmax 20/20 luminometer (any single-tube luminometer works well)
Software and datasets
GraphPad Prism
Procedure
Pre-activation and storage of SLO aliquots
Carefully open the bottle containing the lyophilized SLO.
Add 500 μL of PBS + 10 mM DTT to the bottle.
Resuspend the SLO by inverting the bottle multiple times and briefly vortexing.
Pre-activate the SLO by placing the bottle within a 37 °C incubator for 2 h.
Measure the protein concentration of the reconstituted SLO using the protein assay dye reagent.
Snap-freeze 10 μL aliquots of pre-activated SLO in liquid nitrogen.
Store at -80 °C until use in Procedure C.
Note: Avoid repeated freeze-thaw cycles of the pre-activated SLO aliquots.
Isolation of cytosol from cultured HCT116 WT cells
Culture 20 mm × 150 mm plates of HCT116 WT cells to 95% confluence in 30 mL of cell culture growth medium at 37 °C in 5% CO2 (DMEM, GlutaMAXTM + 10% FBS).
Place the cells on ice and aspirate the conditioned medium.
Wash the cells with 10 mL of cold PBS per 150 mm plate.
Aspirate the PBS wash buffer.
Harvest the cells in cold PBS (containing protease inhibitors) using a cell scraper (1 mL of PBS per 150 mm plate) and transfer to a pre-chilled 50 mL conical tube.
Note: This is done five plates at a time. Add 5 mL of cold PBS to one 150 mm plate, harvest the cells, and use this buffer to collect the cells with the next four plates.
Centrifuge the cells at 200× g for 5 min at 4 °C to sediment the cells. Discard the supernatant.
Resuspend the cell pellet in 3 mL of cold hypotonic lysis buffer and incubate on ice for 15 min.
Transfer the cell suspension to a pre-chilled 7 mL dounce homogenizer.
Homogenize the cells using approximately 80 strokes with a tight-fitting dounce.
Centrifuge at 1,000× g for 15 min at 4 °C in a SorvallTM ST16R centrifuge using a TX-200 swinging bucket rotor with a 15 mL conical tube adapter to sediment unruptured cells.
Collect the supernatant conservatively and transfer to a 5 mL ultra-clear tube.
Note: The volume of the post-nuclear supernatant should be ~3–4 mL. Be careful not to disturb the pellet when collecting the supernatant.
Centrifuge at ~128,000× g (32,500 rpm) for 30 min at 4 °C in an Optima XE-90 ultracentrifuge using a SW 55 Ti rotor to sediment cellular membranes.
Collect the supernatant (cytosol fraction) conservatively and transfer to a 4 mL Amicon 3k concentrator.
Centrifuge at 4,000× g for 6 × 10 min at 4 °C in a SorvallTM ST16R centrifuge using a TX-200 swinging bucket rotor with a 15 mL conical tube adapter to concentrate the cytosol.
Note: Resuspend the cytosol after each 10-min centrifugation step to prevent protein precipitation. Monitor the retentate and flowthrough using the protein assay dye reagent to ensure minimal protein loss.
Collect the cytosol and measure the protein concentration using the Bio-Rad protein assay dye reagent.
Note: The cytosol concentration should be ~40 mg/mL. This may vary depending on the cell line utilized.
Snap-freeze aliquots of cytosol in liquid nitrogen.
Store at -80 °C until use in Procedure D.
Note: Avoid repeated freeze-thaw cycles of the concentrated cytosol aliquots.
Preparation of SLO-permeabilized HCT116 CD63-Nluc cells
Culture HCT116 CD63-Nluc cells to ~80% confluence in the desired number of wells within a 24-well PDL-coated plate. We typically utilize three technical replicates per experimental condition.
Note: Do not allow cells to become overconfluent. We have observed variability in SLO permeabilization in overconfluent cultures.
Place the cells on ice and aspirate the conditioned medium.
Wash each well with 500 μL of cold PBS containing 1 mM EGTA.
Aspirate the PBS wash buffer and replace with 200 μL of SLO binding buffer.
Gently shake at 4 °C for 15 min on a lateral shaker to allow SLO to bind to the surface of the cells.
Aspirate the SLO binding buffer and wash with 500 μL of cold transport buffer to remove excess unbound SLO.
Aspirate the wash buffer and add 500 μL of pre-warmed permeabilization buffer.
Incubate the 24-well plate at 37 °C for 10 min to initiate cell permeabilization.
Assess cell permeabilization (%) using the 10× objective of a phase contrast microscope.
Note: The nucleoli of SLO-permeabilized cells should appear very distinct. Approximately 95%–100% of cells should be permeabilized at this stage (Figure 1).
Figure 1. Morphology of unpermeabilized and Streptolysin O (SLO)-permeabilized cells. HCT116 CD63-Nluc cells were processed as detailed in steps C1–C9 without (left) or with (right) SLO addition. Scale bars: 100 μm.
Return the 24-well plate to ice.
Aspirate the permeabilization buffer, replace with 500 μL of cold transport buffer, and gently shake the 24-well plate at 4 °C for 10 min on a lateral shaker.
Note: Start to assemble the reaction mixes for the reconstitution reactions at this time (protocol detailed in step C1 and Table 1).
Table 1. Sample calculation for biochemical exosome secretion reconstitution reactions. Each column represents a single condition for the reconstitution reaction. The total volume for each reaction mix is 250 μL for one reaction replicate and pipetting error. Each row represents the component to be added to each reaction, and the volumes indicated are in microliter. Each reaction mix can be scaled for the desired number of reaction replicates.
(1) (2) (3) (4) (5)
Reagent\condition Membranes alone Membranes + ATPr/GTP Membranes + ATPr/GTP + cytosol Membranes + ATPr/GTP + CaCl2 Membranes + ATPr/GTP + cytosol + CaCl2
Transport buffer 250 221.2 196.2 216.2 191.2
10× ATPr - 25 25 25 25
10 mM GTP - 3.8 3.8 3.8 3.8
Cytosol (40 mg/mL) - - 25 - 25
100 mM CaCl2 - - - 5 5
Total 250 250 250 250 250
Aspirate the transport buffer, replace with 500 μL of cold high-salt transport buffer, and gently shake the plate at 4 °C for 10 min on a lateral shaker.
Aspirate the high-salt transport buffer, replace with 500 μL of cold transport buffer, and gently shake the plate at 4 °C for 10 min on a lateral shaker.
Proceed immediately to Procedure D.
Biochemical reconstitution of Ca2+-dependent exosome secretion
In low retention tubes, sequentially assemble the reconstitution reaction mixes by adding the reaction components (from top to bottom). A complete 200 μL reaction contains an ATP regeneration system (1 mM ATP, 40 mM creatine phosphate, 0.2 mg/mL creatine phosphokinase), 0.15 mM GTP, 4 mg/mL cytosol, and 2 mM CaCl2.
Aspirate the wash buffer from step C13 and replace with 200 μL of the reaction mix for each condition.
Incubate the 24-well plate on ice for 5 min.
Place the entire 24-well plate in a 30 °C water bath for 2 min to stimulate exosome secretion.
Place the 24-well plate back on ice and immediately load 100 μL of each reaction supernatant into an AcroPrep 96-well 0.4 μm filter plate placed on top of a 96-well collection plate.
Centrifuge the AcroPrep 96-well 0.4 μm filter plate at 1,500× g for 1 min at 4 °C in an Eppendorf 5810 R centrifuge using a A-4-62 swinging bucket rotor with a plate adaptor.
While waiting on the centrifuge run, add 100 μL of cold transport buffer + 2% TX-100 (containing protease inhibitors) to each well of cells within the 24-well plate (for a final TX-100 concentration of 1%) to lyse the cells.
The filtrate collected from step D6 is used to measure exosome secretion, and the lysate from step D7 is used to normalize exosome secretion between sample conditions.
Luminescence measurements
For the luminescence measures to quantify exosome secretion:
Add 50 μL of the filtrate from step D6 to a microcentrifuge tube.
Add 100 μL of Nluc substrate/inhibitor master mix to the filtrate.
Vortex the sample briefly (~1 s) and measure luminescence in a Promega Glowmax 20/20 luminometer.
Note: This luminescence reading (Measurement A) represents CD63-Nluc luminescence obtained from intact exosomes.
Remove the sample tube from the luminometer and add 1.5 μL of 10% TX-100 (for a final TX-100 concentration of 0.1%) to solubilize membranes.
Note: This allows the membrane-impermeable Nluc inhibitor to quench any luminescence derived from membrane-protected compartments.
Vortex the sample briefly (~1 s) and measure luminescence in a Promega Glowmax 20/20 luminometer.
Note: This luminescence reading (Measurement B) represents background Nluc luminescence for each sample.
For the luminescence measurements to normalize between samples:
Add 50 μL of the lysate from step D7 to a microcentrifuge tube.
Add 50 μL of Nluc lytic master mix to the lysate.
Vortex the sample briefly (~1 s) and measure luminescence in a Promega Glowmax 20/20 luminometer.
Note: This luminescence reading (Measurement C) represents total CD63-Nluc luminescence from cells and is used to normalize exosome secretion between samples.
Data analysis
The formula to calculate the exosome production index (EPI) is as follows:
EPI = [(Measurement A) - (Measurement B)]/Measurement C
The EPI can then be normalized to the desired control condition. Example data are presented in Figure 2.
Figure 2. Biochemical reconstitution of Ca2+-dependent exosome secretion. (A, B). Exosome secretion from Streptolysin O (SLO)-permeabilized CD63-Nluc cells was assessed under different reaction conditions. The experimental conditions are indicated below each column and refer to the conditions described in Table 1 (“B” indicates baseline). Data plotted represent the means from three independent experiments, and error bars represent each standard deviation. Statistical significance was evaluated in GraphPad Prism using an ANOVA (*p < 0.05, ***p < 0.001, ****p < 0.0001, and ns = not significant) (Modified from Williams et al., 2023).
General notes and troubleshooting
We have observed variability in the activity of commercial SLO preparations. We recommend titrating each batch of SLO to identify the optimal concentration required to permeabilize a majority of cells as depicted in Figure 1. If needed, trypan blue exclusion can be utilized to confirm cell permeabilization.
During optimization of this reconstitution assay, we observed a high level of background due to insufficient cytosol depletion. The identity and concentration of the salt utilized for the high-salt wash (step C12) may need to be optimized through empirical testing. We recommend conducting immunoblot analysis for a cytosolic marker (e.g., GAPDH) before and after the high-salt wash to ensure efficient cytosol depletion.
Acknowledgments
We thank Dr. Chitose Oneyama (Aichi Cancer Center Research Institute, Chikusa-ku, Nagoya, Japan) for kindly providing the HCT116 CD63-Nluc cell line. We also thank current and past members of the Schekman lab for helpful discussions on optimizing this protocol. JMN is supported by a National Science Foundation Graduate Research Fellowship and a Ruth L. Kirschstein NRSA Predoctoral Fellowship (F31CA284881). RS is an Investigator of the Howard Hughes Medical Institute, a Senior Fellow of the UC Berkeley Miller Institute of Science, and Chair of the Scientific Advisory Board of Aligning Science Across Parkinson’s Disease (ASAP). This protocol has been adapted from Williams et al. (2023).
Competing interests
The authors declare that no competing interests exist.
References
Colombo, M., Raposo, G. and Théry, C. (2014). Biogenesis, Secretion, and Intercellular Interactions of Exosomes and Other Extracellular Vesicles. Annu. Rev. Cell Dev. Biol. 30(1): 255–289.
Driedonks, T. A. P. and Nolte-’t Hoen, E. N. M. (2019). Circulating Y-RNAs in Extracellular Vesicles and Ribonucleoprotein Complexes; Implications for the Immune System. Front. Immunol. 9: e03164.
Fong, M. Y., Zhou, W., Liu, L., Alontaga, A. Y., Chandra, M., Ashby, J., Chow, A., O’Connor, S. T. F., Li, S., Chin, A. R., et al. (2015). Breast-cancer-secreted miR-122 reprograms glucose metabolism in premetastatic niche to promote metastasis. Nat. Cell Biol. 17(2): 183–194.
Harding, C., Heuser, J. and Stahl, P. (1983). Receptor-mediated endocytosis of transferrin and recycling of the transferrin receptor in rat reticulocytes. J. Cell Biol. 97(2): 329–339.
Hikita, T., Miyata, M., Watanabe, R. and Oneyama, C. (2018). Sensitive and rapid quantification of exosomes by fusing luciferase to exosome marker proteins. Sci. Rep. 8(1): e1038/s41598-018-32535-7.
Hsu, Y. L., Hung, J. Y., Chang, W. A., Lin, Y. S., Pan, Y. C., Tsai, P. H., Wu, C. Y. and Kuo, P. L. (2017). Hypoxic lung cancer-secreted exosomal miR-23a increased angiogenesis and vascular permeability by targeting prolyl hydroxylase and tight junction protein ZO-1. Oncogene 36(34): 4929–4942.
Shurtleff, M. J., Yao, J., Qin, Y., Nottingham, R. M., Temoche-Diaz, M. M., Schekman, R. and Lambowitz, A. M. (2017). Broad role for YBX1 in defining the small noncoding RNA composition of exosomes. Proc. Natl. Acad. Sci. U. S. A. 114(43): e1712108114.
Upton, H. E., Ferguson, L., Temoche-Diaz, M. M., Liu, X. M., Pimentel, S. C., Ingolia, N. T., Schekman, R. and Collins, K. (2021). Low-bias ncRNA libraries using ordered two-template relay: Serial template jumping by a modified retroelement reverse transcriptase. Proc. Natl. Acad. Sci. U.S.A. 118(42): e2107900118.
Walker, J. R., Hall, M. P., Zimprich, C. A., Robers, M. B., Duellman, S. J., Machleidt, T., Rodriguez, J. and Zhou, W. (2017). Highly Potent Cell-Permeable and Impermeable NanoLuc Luciferase Inhibitors. ACS Chem. Biol. 12(4): 1028–1037.
Williams, J. K., Ngo, J. M., Lehman, I. M. and Schekman, R. (2023). Annexin A6 mediates calcium-dependent exosome secretion during plasma membrane repair. eLife 12: e86556.
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Habituation of Sugar-Induced Proboscis Extension Reflex and Yeast-Induced Habituation Override in Drosophila melanogaster
ST Swati Trisal
KV K. VijayRaghavan
MR Mani Ramaswami
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4891 Views: 424
Reviewed by: Nafisa M. JadavjiSurya Jyoti Banerjee Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Neuroscience April 2022
Abstract
Habituation, the process by which animals learn to ignore insignificant stimuli, facilitates engagement with salient features of the environment. However, neural mechanisms underlying habituation also allow responses to familiar stimuli to be reinstated when such stimuli become potentially significant. Thus, the habituated state must allow a mechanism for habituation override. The remarkably precise knowledge of cell identity, connectivity, and information coding in Drosophila sensory circuits, as well as the availability of tools to genetically target these cells, makes Drosophila a valuable and important organism for analysis of habituation and habituation-override mechanisms. Studies of olfactory and gustatory habituation in Drosophila suggest that potentiation of GABAergic neurons underlies certain timescales of habituation and have specified some elements of a gustatory habituation-override pathway. More detailed understanding of gustatory habituation and habituation-override mechanisms will benefit from access to robust behavioral assays for (a) the proboscis extension reflex (PER) elicited by a sweet stimulus, (b) exposure paradigms that result in PER habituation, and, most critically, (c) manipulations that result in PER-habituation override. Here, we describe simple protocols for persistent sucrose exposure of tarsal hairs that lead to habituation of proboscis extension and for presentation of a novel appetitive stimuli that reinstate robust PER to habituated flies. This detailed protocol of gustatory habituation provides (a) a simple method to induce habituation by continuous exposure of the flies to sucrose for 10 min without leading to ingestion and (b) a novel method to override habituation by presenting yeast to the proboscis.
Key features
• A protocol for stimulation of Drosophila’s taste (sugar) sensory neurons that induces gustatory habituation without satiation due to ingestion.
• A chemical (yeast) stimulation protocol that rapidly induces habituation override/dishabituation in sugar-habituated Drosophila.
Keywords: Proboscis extension reflex Drosophila melanogaster Dishabituation Override Gustatory
Background
Habituation is a fundamental form of learning that is conserved across species. In habituation, the response to a persistent inconsequential stimulus is reduced, leading to an enhanced ability to attend to more pertinent stimuli. A defining feature of the habituated state is the presence of a latent ability to respond to the familiar stimulus (Thompson and Spencer, 1966; Rankin et al., 2009; Ramaswami, 2014). This latent ability to respond is made evident by the effect of a novel external stimulus, which results in override of habituation, i.e., restoration of a robust response in the habituated animal. This can also be observed when an animal is motivated to attend to the familiar stimulus (Kato et al., 2015). The neural mechanism of habituation has been extensively studied across different species and different modalities, particularly in Aplysia, Drosophila, and mouse, using a variety of different behavioral protocols (Castellucci et al., 1970; Krasne and Edwards, 2002; Engel and Wu, 2009; Ogg et al., 2015). In Aplysia, both homosynaptic depression at the sensory-motor synapse (Castellucci et al., 1970) and potentiation of an inhibitory interneuron synapse (Bristol and Carew, 2005) have been suggested as underlying mechanisms of habituation of the gill-induced siphon withdrawal reflex. Later studies in Drosophila and vertebrates indicate that heterosynaptic potentiation of inhibition can lead to olfactory, auditory, or gustatory habituation (Das et al., 2011; Paranjpe et al., 2012; Ramaswami, 2014; Kato et al., 2015). However, the neural correlates of habituation override in these types of habituation are only just beginning to be investigated (Trisal et al., 2022). Here, we first describe the protocol for the proboscis extension reflex (PER), which has been used previously to study gustatory habituation in Drosophila (Duerr and Quinn, 1982; Le Bourg, 1983; Fois et al., 1991; Engel and Wu, 2009; Paranjpe et al., 2012), and then a protocol to induce override of PER habituation following exposure to a novel sensory stimulus (Paranjpe et al., 2012; Trisal et al., 2022). The novel stimulus used is a non-invasive gustatory stimulus (10% yeast), which the flies have not encountered before, that is presented to the proboscis. Compared to previous publications on PER (Shiraiwa and Carlson, 2007; Li et al., 2022), we provide a simpler and potentially less traumatic method to immobilize the flies. In addition, our protocol (also used by Paranjpe et al., 2012) is unique in that it exposes the flies continuously to 10% sucrose (while preventing ingestion at the same time) to induce more robust PER habituation than in the original studies. Our approach allows a more dynamic range and time to study habituation and dishabituation. While our method requires approximately 15 min for analysis of an individual fly, it requires far fewer flies to reach statistical significance.
Materials and reagents
Biological material
Drosophila Canton-Special females, aged 3–4 days
Reagents
Sucrose (Fischer Scientific, catalog number: 15925)
Yeast (Sigma-Aldrich, catalog number: YSC2)
Corn flour (Avenue Supermarts Ltd)
Sugar (Avenue Supermarts Ltd)
D-Glucose (Qualigens, catalog number: Q15405)
Yeast extract (Himedia, catalog number: RM027)
Propionic acid (Qualigens, catalog number: Q26955)
Orthophosphoric acid (Qualigens, catalog number: Q29245)
Ethanol (Merck Millipore, catalog number: 100983)
Agar (Qualigens, catalog number: Q21185)
Tego-sept (Sigma-Aldrich, catalog number: M1650000)
Water (double distilled)
Solutions
10% sucrose solution (see Recipes)
2% sucrose solution (see Recipes)
10% yeast solution (see Recipes)
Corn meal media (1 L) (see Recipes)
Recipes
10% sucrose (weight/volume)
Reagent Quantity
Sucrose 1 g
Water (autoclaved double distilled) 10 mL
2% sucrose (weight/volume)
Reagent Quantity
Sucrose 20 mg
Water (autoclaved double distilled) 1 mL
10% yeast (weight/volume)
Reagent Quantity
Yeast 500 mg
Water (autoclaved double distilled) 5 mL
Corn meal media (1 L)
Reagent Quantity
Corn flour 80 g
D-Glucose 20 g
Sugar 40 g
Agar 8 g
Yeast extract 15 g
Propionic acid 4 mL
Orthophosphoric acid 0.6 mL
Tego-sept 1 g (dissolved in 5 mL ethanol)
Laboratory supplies
Coverslips 18 mm × 18 mm (Polar Industrial Corporation, Blue Star)
1 mL syringes with attached needle of size 0.30 mm × 8 mm, gauge 30 G × 5/16 (Hindustan Syringes and Medical Devices Ltd, model: Dispovan U-40)
Synthetic hairbrush (Camlin, catalog number: 2066774)
Transparent nail polish (Lakme Truewear, model: CG012)
Styrofoam box (20 cm × 14 cm × 8.5 cm)
Crushed ice
Styrofoam tray (19 cm × 16 cm × 3 cm)
Glass vial for cold anesthesia (8.5 cm high × 2 cm diameter)
Metal (aluminum) plate (10 cm diameter)
Timer (West Bend, catalog number: 40005X)
Filter paper No. 1 (Indica, Indicators, catalog number: 74039)
Plastic box of dimension 13 cm × 11 cm × 8 cm
Plastic bottle for Drosophila culture 250 mL volume (Bharat Plastic Manufacturing Company, catalog number: FLBT-20)
Absorbent cotton wool (Prabhat Surgical Cotton Private Limited)
Equipment
Stereomicroscope magnification 20× (10× eyepiece, 2× objective), numerical aperture 0.142 (Olympus, model: SZ51)
Micromanipulator (Narishige Japan, model: MM3)
Behavior room or chamber with controlled temperature (21 °C) and humidity (60%)
Software
GraphPad Prism v8.4.2
Procedure
Fly preparation
Raise flies in standard cornmeal media in plastic 250 mL bottles containing 15–20 mL of media at 25 °C temperature, 60% relative humidity, and 12:12 h light/dark cycle. Transfer adult flies every 2–3 days to prevent crowding and sogging of media.
Collect freshly eclosed flies of either sex, which may be 3–4 days old, in fresh media bottles such that each bottle has 20–30 flies. Age flies for 3–4 days at 25 °C, 60% humidity, and 12:12 h light/dark cycle.
Note: The media is not considered fresh if it is dry and starts separating from the walls of the bottle or has low levels of bacterial or fungal growth.
Starve flies in an empty glass vial containing wet filter paper for 20–24 h at 25 °C, 60% humidity, and 12:12 h light/dark cycle (Figure 1A).
Figure 1. Preparation of flies for proboscis extension reflex assay. (A) Flies are starved in a vial overnight with a wet filter paper. (B) Flies are anesthetized on ice for not more than 1 min. (C) Anesthetized flies in the vial. (D) Anesthetized flies are transferred on a metal plate kept on ice. (E) Single anesthetized fly on the metal plate turned dorsal side up in order to stick to the coverslip using nail polish. (F), (G) Fly stuck on the coverslip with ventral side up. (H) Size of the drop of nail polish on the syringe used to immobilize flies on the coverslip. (I) Flies are kept in a humidified chamber for 1–2 h for recovery. (J) Syringe containing 10% sucrose for habituation, mounted on a micromanipulator.
To anesthetize the starved flies, first cool an empty glass vial by placing it in a Styrofoam box filled with ice. After 5 min, transfer the starved flies into the now cold vial. To transfer the flies, tap the vial containing the flies such that they come to the bottom of the vial and quickly invert this vial over the cold empty vial. Tap both the vials together such that all flies are transferred to the cold vial.
Gently tap the cold vial on a rubber pad to bring the flies down to the bottom of the vial and place the vial back in ice for one more minute until all flies are anesthetized (Figure 1B, 1C).
Place a metal plate on a Styrofoam tray filled with ice and wipe it with filter paper to remove condensed water. Gently transfer the anesthetized flies onto the metal plate kept on ice. Make sure the plate is dry by wiping it repeatedly before or after transferring flies onto it, whenever water condenses over it. Keep only the female flies and discard all male flies. Male and female flies are distinguished based on the standard sexually dimorphic traits like male leg sex combs, male genitalia, and female abdomen pigmentation. Using a fine synthetic hairbrush, turn the flies’ dorsal side up (Figure 1D, 1E). It is important to note that the total amount of time flies spend on ice, including both in the vial and on the metal plate, should not be more than 5 min, as it can affect their response.
Take a small amount of transparent nail polish (~1 μL) using the needle of a 1 mL syringe (Figure 1H) and place the drop on the coverslip. Gently place the coverslip on the fly such that the nail polish touches the posterior part of the head and anterior part of the thorax. This is done so that the head does not move while performing PER. It is also important to use only a small drop of nail polish on the coverslip so that it does not reach the proboscis or the legs while fixing the fly (Figure 1F, 1G). Small bubbles may appear occasionally in the nail polish but that does not affect the immobilization of the fly on the coverslip. Note that all these steps requiring flies to be anesthetized are performed on the metal plate.
Place the fly fixed on the coverslip (in a ventral side up direction so that the proboscis and legs are exposed for stimulation) in a box for 1–2 h for recovery at 25 °C. The box is lined with wet filter paper as the coverslips with flies are at the base, to maintain humidity inside (Figure 1I). Flies are transferred manually by gently placing the coverslip with the immobilized flies in the box. The number of flies placed in the box depends on its size. A box with the dimensions mentioned in the current study allows a maximum of 15–20 flies so that crowding and therefore overlap can be avoided. The minimum number of flies can be determined by the user.
Habituation of the sucrose-induced PER
Fill three 1 mL syringes: one with 2% sucrose solution, one with 10% sucrose solution, and the third with double-distilled water. Fix the 1 mL syringe with 10% sucrose solution on a micromanipulator in the slot provided (Figure 1I).
Gently pick the coverslip with the immobilized fly and place it under the stereoscope. Using the syringe containing double-distilled water, touch the proboscis or tarsus of the fly with water, as shown in Video 1. Allow the fly to drink water through proboscis until it stops and is sated. Flies are sated after being presented with water 1–2 times. If the fly does not respond to water, consider the fly for the experiment and test the naïve response to sucrose as explained in step B3.
Note: The occasional fly may not stop drinking water even after being presented with water three or more times; discard such flies.
Video 1. Water satiation. Initially, the fly is presented with water. If the fly extends its proboscis, water is offered to satiate it until there is no further response to water.
Immediately after feeding water to the fly, carefully touch the distal tips of the foreleg tarsi with a sucrose drop with the syringe containing 2% sucrose, while preventing any contact with the proboscis. Record whether there is a proboscis extension in response or not. Repeat this step five times with a gap of 10 s between each two trials (Video 2).
Video 2. Proboscis extension reflex (PER) in response to 2% sucrose. Video in real time showing the tarsi of the forelegs being stimulated with 2% sucrose five times, with 10 s of inter-trial interval, to quantify the PER response before and after exposure to habituating stimulus.
Using the knobs attached to the micromanipulator to move the syringe in XY and Z plane, adjust the angle and height of the syringe containing 10% sucrose fixed on the micromanipulator, such that the sucrose drop hanging from the syringe is out of reach from the extended proboscis and touches only the distal tips of the forelegs (Video 3). Keep the syringe in this position, touching the tarsi, for 10 min, set on a timer. If the size of the sucrose drop becomes smaller due to evaporation, carefully press the plunger of the syringe and readjust the size. Flies are exposed to 10% sucrose immediately after step B3.
Video 3. Exposure to habituating stimulus. The tarsi of the fly are exposed to 10% sucrose for 10 min in order to habituate the PER response. The video is representative, showing 12 s from the 10 min exposure. The syringe containing 10% sucrose is fixed on a micromanipulator such that only tarsi are stimulated whereas the proboscis cannot reach the stimulus.
After 10 min, move aside the 10% sucrose syringe and wash the tarsal tips with double-distilled water immediately after sucrose exposure (Video 4).
Video 4. Leg wash after exposure to 10% sucrose. The tarsi are washed with double-distilled water to clear 10% sucrose that may be sticking to the legs.
Repeat step B3 immediately after leg wash to record the habituated response. Record all data. In a habituated fly, the number of times the proboscis is extended in response to 2% sucrose should decrease (Video 5).
Video 5. Response after sucrose exposure. After exposure to 10% sucrose for 10 min, tarsal stimulation with 2% sucrose does not lead to PER response, thus exhibiting habituation.
Dishabituation using yeast
Touch the proboscis of the habituated fly with 10% yeast solution using a 1 mL syringe and quickly remove the syringe in order to prevent or minimize potential consumption of yeast (Video 6). Repeat this over a period of 1 min, set on a timer, with a gap of approximately 10 s such that the proboscis is stimulated 5–6 times with yeast.
Video 6. Dishabituation/habituation override induced by 10% yeast. 10% yeast is presented five times to the proboscis in order to dishabituate the fly. Yeast is quickly removed in order to prevent ingestion.
Leave the fly for 1 min without any stimulus so that it gets time to groom itself in order to remove any sticking yeast.
Record the proboscis extension response to 2% sucrose as described previously, in yeast-exposed flies. The flies should show increased (dishabituated) response.
The procedure to induce habituation to sucrose and its override using 10% yeast is summarized in Figure 2.
Figure 2. Line diagram representing the protocol. Figure shows the various key steps in the protocol to habituate flies to sucrose and override it by presenting 10% yeast to proboscis. The figure also shows the time period of the time-bound steps.
Data analysis
For each fly, calculate the percentage of proboscis extensions in response to five tarsal stimulations using a simple formula as = [Total observed number of proboscis extension/5] × 100.
A total of 25–30 flies are required to achieve statistical significance. Therefore, 2–3 bottles containing 20–30 flies each may be required to get 25–30 female flies, which are tested in the experiment. Since data is paired, all the conditions have the same number of flies. PER response of the first five tarsal stimulations is considered. More stimulations are avoided to prevent any decrease in response due to habituation. Plot these PER response data observed during different conditions in all the flies tested as bar graphs representing the mean percentage PER with SEM error bars and individual fly data points overlaid on it using GraphPad Prism v8.4.2.
Note: Compare the %PER response observed with 2% sucrose during naïve, habituated, and dishabituated conditions only. PER response during 10 min exposure to 10% sucrose is not considered for data analysis.
The data is not normally distributed due to its discrete nature and therefore no normality test is required. The Friedman test, a non-parametric test, followed by Dunn’s post-hoc test is used to evaluate differences across naïve, habituated, and dishabituated responses. To perform Friedman test in GraphPad Prism software, carry out the following steps:
In the Result section, Select New Analysis.
Under Column Analysis section, select One way ANOVA (non-parametric or mixed) and tick the columns that need to be analyzed.
A new window opens. In the tab Under the Experimental Design, select Each row represents matched or repeated measures, data.
Select No. use nonparametric test under the option Assume Gaussian Distribution of residuals option.
Under the Multiple Comparison tab, select compare the mean of each column with every other column. In case of comparing PER responses between fed and starved flies (Figure 3A1) and between sucrose and water (Figure 3A2), Mann Whitney U test is used. Follow the steps from 3a to 3d, with the exception of under Column Analysis, where you should select the t tests (and nonparametric tests) option.
If the fly extends the proboscis to 2% sucrose less than three out of five trials while testing for the naïve response, discard the fly.
Raw data of naïve PER response to 2% sucrose, habituated response after 10 min of exposure to 10% sucrose, and PER response after presentation of 10% yeast solution, tested in 30 Canton S flies, is shown in Table 1; a representative bar plot of the data with individual data points is shown in Figure 3B. This protocol has previously been shown to mediate and override habituation (Figure 2B in Trisal et al., 2022). Analysis of PER response in fed and starved, age- and density-matched CS flies to 2% sucrose shows that PER is specifically induced by sucrose (Figure 3A1). In addition, water sated flies do not respond to water, whereas PER is observed when stimulated with sucrose, further confirming that PER is specific to sucrose (Figure 3A2).
Figure 3. Habituation override using a novel yeast stimulus. (A1) Control experiment shows proboscis extension reflex (PER) response in fed and starved flies. PER response is specific to 2% sucrose, as fed flies show significantly lower response compared to starved flies (Mann Whitney U stat = 159, ***p < 0.0001). (A2) Comparison of PER response to sucrose and water in water sated flies. Flies extend their proboscis to 2% sucrose, which is not observed when water is further presented, confirming that PER is specific to sucrose (Mann Whitney U stat = 194.5, ***p < 0.0001. (B) Presentation of 2% sucrose solution to the tarsal hairs of Canton S flies generates a proboscis extension response. PER response shows a significant decrease upon exposure to 10% sucrose for 10 min, which can be overridden significantly upon presentation of 10% yeast solution (Friedman statistic = 42.70, ***p < 0.0001, **p = 0.0072, Dunn’s multiple comparison post-hoc test). Triangles represent individual data point whereas bars represent mean ± SEM (error bars).
Table 1. Average %PER to 2% sucrose in 30 naïve, habituated, or dishabituated individual flies calculated from five stimulations
Naive response (% PER) Habituated response (% PER) Response after yeast exposure (% PER)
100 0 80
80 20 0
100 0 80
100 20 0
100 0 80
60 0 100
100 0 60
100 60 100
100 0 20
100 0 100
80 0 0
100 100 100
100 0 100
100 20 100
100 40 80
100 0 0
100 0 100
80 0 60
100 0 100
100 0 100
100 100 100
80 0 60
60 0 0
100 60 100
100 0 100
80 0 60
100 0 0
100 0 0
100 0 60
100 0 0
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Trisal et al. (2022). A Drosophila circuit for habituation override. Journal of Neuroscience (Figure 2, panel B).
General notes and troubleshooting
Mounted flies must be healthy and undamaged and capable of robust PER before they can be used for experiments. In protocol section B, step 3, we therefore discard any flies that respond less than three times to five sequential tarsal stimulations with 2% sucrose. If a large fraction of the flies in the batch do not respond to 2% sucrose or fail to habituate, make sure the flies are starved for 20–24 h, are not crowded, and are not kept on ice for more than 5 min. While dishabituating the flies using 10% yeast as in protocol section C, step 1, make sure flies do not consume significant amounts of yeast. In protocol section B, step 2, discard flies that do not stop consuming water, as such flies are likely damaged in some way.
Acknowledgments
We thank Marcia Aranha, Camilla Roselli, Ankita Chodankar and members of Ramaswami and VijayRaghavan labs for advice, support and useful discussions, particularly Pushkar Paranjpe for early help setting up the PER assay. We thank Spraha Bhandari for volunteering to test and confirm that this protocol was sufficient for a new researcher to successfully observe and record PER, its habituation to sucrose, and yeast-induced override. This protocol was previously described in Paranjpe et al. (2012) and Trisal et al. (2022). The work was supported by a Wellcome Trust Investigator Award and a Science Foundation Ireland Investigator Programme Grant to MR, by core support from the National Centre for Biological Sciences (Tata Institute of Fundamental Research) to KVR and by a CSIR postgraduate fellowship and a Biocon-Trinity Scholarship to ST.
Competing interests
The authors declare no competing financial interests.
References
Bristol, A. S. and Carew, T. J. (2005). Differential role of inhibition in habituation of two independent afferent pathways to a common motor output. Learn. Mem. 12(1): 52–60.
Castellucci, V., Pinsker, H., Kupfermann, I. and Kandel, E. R. (1970). Neuronal Mechanisms of Habituation and Dishabituation of the Gill-Withdrawal Reflex in Aplysia. Science 167(3926): 1745–1748.
Das, S., Sadanandappa, M. K., Dervan, A., Larkin, A., Lee, J. A., Sudhakaran, I. P., Priya, R., Heidari, R., Holohan, E. E., Pimentel, A., et al. (2011). Plasticity of local GABAergic interneurons drives olfactory habituation. Proc. Natl. Acad. Sci. U.S.A. 108(36): e1106411108.
Duerr, J. S. and Quinn, W. G. (1982). Three Drosophila mutations that block associative learning also affect habituation and sensitization. Proc. Natl. Acad. Sci. U. S. A. 79(11): 3646–3650.
Engel, J. E. and Wu, C. F. (2009). Neurogenetic approaches to habituation and dishabituation in Drosophila. Neurobiol. Learn. Mem. 92(2): 166–175.
Fois, C., Médioni, J. and Le Bourg, E. (1991). Habituation of the proboscis extension response as a function of age in Drosophila melanogaster. Gerontology 37(4): 187–192.
Kato, H. K., Gillet, S. N. and Isaacson, J. S. (2015). Flexible Sensory Representations in Auditory Cortex Driven by Behavioral Relevance. Neuron 88(5): 1027–1039.
Krasne, F. B. and Edwards, D. H. (2002). Modulation of the Crayfish Escape Reflex--Physiology and Neuroethology. Integr. Comp. Biol. 42(4): 705–715.
Le Bourg, E. (1983). Aging and habituation of the tarsal response in Drosophila melanogaster. Gerontology 29(6): 388–393.
Li, Q., Yang, L. and Montell, C. (2022). Drosophila proboscis extension response and GCaMP imaging for assaying food appeal based on grittiness. STAR protocols 3(4): 101806.
Ogg, M. C., Bendahamane, M. and Fletcher, M. L. (2015). Habituation of glomerular responses in the olfactory bulb following prolonged odor stimulation reflects reduced peripheral input. Front. Mol. Neurosci. 8: 53.
Paranjpe, P., Rodrigues, V., VijayRaghavan, K. and Ramaswami, M. (2012). Gustatory habituation in Drosophila relies on rutabaga (adenylate cyclase)-dependent plasticity of GABAergic inhibitory neurons. Learn. Mem. 19(12): 627–635.
Ramaswami, M. (2014). Network plasticity in adaptive filtering and behavioral habituation. Neuron 82(6): 1216–1229.
Rankin, C. H., Abrams, T., Barry, R. J., Bhatnagar, S., Clayton, D. F., Colombo, J., Coppola, G., Geyer, M. A., Glanzman, D. L., Marsland, S., et al. (2009). Habituation revisited: An updated and revised description of the behavioral characteristics of habituation. Neurobiol. Learn. Mem. 92(2): 135–138.
Shiraiwa, T. and Carlson, J. R. (2007). Proboscis Extension Response (PER) Assay in Drosophila. J. Vis. Exp. (3): 193.
Thompson, R. F. and Spencer, W. A. (1966). Habituation: A model phenomenon for the study of neuronal substrates of behavior. Psychol. Rev. 73(1): 16–43.
Trisal, S., Aranha, M., Chodankar, A., VijayRaghavan, K. and Ramaswami, M. (2022). A Drosophila Circuit for Habituation Override. J. Neurosci. 42(14): 2930–2941.
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4,892 | https://bio-protocol.org/en/bpdetail?id=4892&type=0 | # Bio-Protocol Content
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Peer-reviewed
Expression and Purification of Recombinant Human Mitochondrial RNA Polymerase (POLRMT) and the Initiation Factors TFAM and TFB2M
AH An H. Hsieh
SR Sean D. Reardon
JM Jubilee H. Munozvilla-Cabellon
JS Jiayu Shen
SP Smita S. Patel
TM Tatiana V. Mishanina
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4892 Views: 1054
Reviewed by: Abhilash PadavannilJeremy BirdHassan Rasouli
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Apr 2022
Abstract
Human mitochondrial DNA (mtDNA) encodes several components of oxidative phosphorylation responsible for the bulk of cellular energy production. The mtDNA is transcribed by a dedicated human mitochondrial RNA polymerase (POLRMT) that is structurally distinct from its nuclear counterparts, instead closely resembling the single-subunit viral RNA polymerases (e.g., T7 RNA polymerase). The initiation of transcription by POLRMT is aided by two initiation factors: transcription factor A, mitochondrial (TFAM), and transcription factor B2, mitochondrial (TFB2M). Although many details of human mitochondrial transcription initiation have been elucidated with in vitro biochemical and structural studies, much remains to be addressed relating to the mechanism and regulation of transcription. Studies of such mechanisms require reliable, high-yield, and high-purity methods for protein production, and this protocol provides the level of detail and troubleshooting tips that are necessary for a novice to generate meaningful amounts of proteins for experimental work. The current protocol describes how to purify recombinant POLRMT, TFAM, and TFB2M from Escherichia coli using techniques such as affinity column chromatography (Ni2+ and heparin), how to remove the solubility tags with TEV protease and recover untagged proteins of interest, and how to overcome commonly encountered challenges in obtaining high yield of each protein.
Key features
• This protocol builds upon purification methods developed by Patel lab (Ramachandran et al., 2017) and others with greater detail than previously published works.
• The protocol requires several days to complete as various steps are designed to be performed overnight.
• The recombinantly purified proteins have been successfully used for in vitro transcription experiments, allowing for finer control of experimental components in a minimalistic system.
Keywords: POLRMT TFAM TFB2M Protein purification Bacterial protein expression Ni-NTA Maltose binding protein (MBP) fusion protein purification TEV protease Heparin
Background
The mitochondrial RNA polymerase (POLRMT) is a DNA-dependent RNA polymerase (RNAP) that transcribes the human mitochondrial DNA (mtDNA), which encodes several protein components of the complexes that carry out oxidative phosphorylation. POLRMT is structurally similar to other single-subunit RNAPs such as T7 phage RNAP (Schwinghammer et al., 2013; Sultana et al., 2017), and current models suggest that POLRMT transcribes short RNA primers used in mtDNA replication initiation (Tan et al., 2022). Considering the central role of POLRMT in cellular metabolism and maintenance of mtDNA, it is not surprising that mutations associated with mitochondrial dysfunction map to POLRMT, such as respiratory chain defects, hypotonia, and global developmental delays (Oláhová et al., 2021).
To start transcription de novo from a promoter sequence, POLRMT requires two initiation factors: transcription factor A (TFAM) and transcription factor B2 (TFB2M), which aid the melting of the promoter sequence and the formation of a stable initiation complex (Hillen et al., 2017; Ramachandran et al., 2017). TFAM is also the mtDNA packaging protein that compacts the mtDNA into structures called nucleoids. While too much TFAM can prevent proteins involved in transcription and other processes from accessing the mtDNA, TFAM is required to bend the upstream promoter region to recruit POLRMT to the promoter site; the underlying mechanisms of this process are yet to be worked out. Published work suggests that post-translational modifications (PTMs) to TFAM, such as acetylation or phosphorylation, do not compromise its function as a transcription factor but make it easier for POLRMT to remove TFAM from DNA, thus increasing transcription processivity through TFAM-compacted DNA (Reardon and Mishanina, 2022).
This protocol describes in detail how to isolate recombinant POLRMT, TFAM, and TFB2M from Escherichia coli in high purity and how to overcome the challenges in achieving high protein yield. These protocols build upon the procedures developed in the Patel lab and others (Yakubovskaya et al., 2014; Ramachandran et al., 2017) and describe the steps in greater detail than in any previously published work to provide access to these resources to a wider community. The resulting proteins can be used to reconstitute human mitochondrial transcription on promoter sequences in vitro, as in prior biochemical and structural studies (Bird et al., 2018; Tan et al., 2022). These proteins can be post-translationally modified to probe the impact of PTMs on their function to fine-tune mitochondrial transcription output (Reardon and Mishanina, 2022). Finally, the protocols can also be applied to express and purify mutant proteins to study the underlying basis of mutations associated with mitochondrial dysfunction.
Materials and reagents
4× Laemmli sample buffer (Bio-Rad, catalog number: 1610747)
10× Tris/Glycine/SDS (Bio-Rad, catalog number: 1610732)
Dithiothreitol (DTT) (Fisher BioReagents, catalog number: BP172-5), see Recipes for preparation
Ethanol (EtOH) (Fisher Scientific, catalog number: 2818500)
Glycerol (Fisher Scientific, catalog number: BP229-1)
HiTrap heparin column (GE Healthcare, catalog number: 17-0407-03)
Hydrochloric acid (HCl) (Fisher Scientific, catalog number: LC149502)
Imidazole (molecular biology grade, > 99%) (Fisher Scientific, catalog number: O3196-500)
Isopropyl-β-D-1-thiogalactopyranoside (IPTG) (Fisher Scientific, catalog number: BP1755100), see Recipes for preparation
LB agar, Miller (Fisher Scientific, catalog number: BP1425-2)
LB broth (LB), Miller, granulated (Fisher Scientific, catalog number: BP9723-2)
Lysozyme (Fisher Scientific, catalog number: BP535-5)
Precision Plus ProteinTM All Blue Prestained Protein Standards (Bio-Rad, catalog number: 1610373)
Protease inhibitor cocktail (PIC) powder (used in this protocol as 1,000× stock and prepared following instructions online: https://www.sigmaaldrich.com/US/en/product/sigma/p2714) (Sigma-Aldrich, catalog number: P2714-1BTL)
Sodium hydroxide (NaOH) pellets (Fisher Scientific, catalog number: S318-500)
β-mercaptoethanol (BME) (Fisher Scientific, catalog number: AC125472500)
Part I: POLRMT purification materials and reagents
Amicon Ultra-15 centrifugal filter units (50K MWCO, 15 mL) (Millipore Sigma, catalog number: UFC905024)
Ammonium sulfate (AS) (Fisher Scientific, catalog number: A702-3)
Ampicillin (Fisher Scientific, catalog number: BP1760-25), see Recipes for preparation
Cytiva CaptoTM DEAE chromatography media (Fisher Scientific, catalog number: 45-002-960) or, if using an FPLC, HiTrap DEAE Fast Flow (Cytiva, catalog number: 17-5154-01)
HisTrap FPLC column (Cytiva, catalog number: 17-3712-06)
Polyethyleneimine (PEI), MN 60,000 50% (Fisher Scientific, catalog number: AC178571000), see Recipes for preparation
Poly-Prep® chromatography columns (Bio-Rad, catalog number: 7311550)
Sodium chloride (NaCl) (Fisher Scientific, catalog number: BP358-212, CAS number: 7647-14-5)
Tris base (Fisher BioReagents, catalog number BP152-1)
Tween 20 (Fisher BioReagents, catalog number: BP337-100)
Part II: TFAM purification materials and reagents
1 kb DNA ladder (NEB, catalog number: N3232S)
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Fisher Scientific, catalog number: BP310-1)
Amicon Ultra-15 centrifugal filter units (3K MWCO, 15 mL) (Millipore Sigma, catalog number: UFC900324)
Ethidium bromide solution, 10 mg/mL (Bio-Rad, catalog number: 1610433)
HisPur Ni-NTA resin (Thermo Scientific, catalog number: 88222)
Kanamycin (Fisher Scientific, catalog number: 611290050), see Recipes for preparation
Potassium chloride (KCl) (molecular biology grade) (Fisher Scientific, catalog number: P217-3)
Sodium chloride (Fisher Scientific, catalog number: BP358-212, CAS number: 7647-14-5)
Part III: TFB2M purification materials and reagents
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Fisher Scientific, catalog number: BP310-1)
Amicon Ultra-4 centrifugal filter units (10K MWCO, 4 mL) (Millipore Sigma, catalog number: UFC801008)
HisPur Ni-NTA resin (Thermo Scientific, catalog number: 88222)
Kanamycin (Fisher Scientific, catalog number: 611290050), see Recipes for preparation
Potassium chloride (KCl) (Fisher Scientific, catalog number: P217-3)
Potassium hydroxide (KOH) (Fisher Scientific, catalog number: P250 500)
Biological materials
Plasmids available upon request
Escherichia coli (E. coli) BL21-CodonPlus (DE3)-RIPL competent cells (Agilent, catalog number: 230280) or
E. coli BL21-CodonPlus (DE3)-RIL competent cells (Agilent, catalog number: 230245)
Optional: E. coli ArcticExpress (DE3) competent cells (Agilent, catalog number: 230192)
Tobacco Etch Virus (TEV) protease (Kapust et al., 2001; Raran-Kurussi et al., 2017)
POLRMT construct in pPROEXHTb. It contains residues 43–1230 [lacking its N-terminal mitochondrial targeting sequence (MTS)] with an N-terminal hexahistidine 6x-His-tag
TFAM construct in pPROEXHTb. It contains residues 43–246 (lacking its N-terminal MTS) with an N-terminal 6x-His-tag and a TEV protease site
TFB2M construct in pT7TEV-HMBP4. It contains residues 20–398 with an N-terminal 6x-His-tag, maltose binding protein (MBP) tag, and a TEV protease site
Solutions
See Recipes for all solutions.
Part I: POLRMT purification
5% Polyethylenimine (PEI) solution pH 7.0
POLRMT Lysis Buffer
POLRMT HisTrap Buffer A
POLRMT HisTrap Buffer B
POLRMT Heparin No Salt Buffer
POLRMT Heparin Buffer A
POLRMT Heparin Buffer B
Part II: TFAM purification and activity test
TFAM Lysis Buffer
TFAM Ni-NTA Buffer A
TFAM Ni-NTA Buffer B
TFAM TEV Cleavage Buffer
TFAM Heparin Buffer A
TFAM Heparin Buffer B
TFAM Storage Buffer (2×)
10× TFAM Binding Buffer
TFAM Electrophoretic Mobility Shift Assay (EMSA) Buffer
Part III: TFB2M Purification
TFB2M Lysis Buffer
TFB2M Ni-NTA Buffer A
TFB2M Ni-NTA Buffer B
TFB2M TEV Cleavage Buffer
TFB2M No Salt Heparin Buffer
TFB2M Heparin Buffer A
TFB2M Heparin Buffer B
Recipes
Common stock solutions
Note: Sterilize all stocks by filtering through a 0.22 μm syringe filter, aliquot into 1 mL portions, and store at -20 °C.
1 M IPTG Stock
2.38 g of IPTG
Dissolve in Milli-Q water and bring the final volume up to 10 mL.
*Add 200 μL of 1 M IPTG per 1 L of cell culture for a final concentration of 0.2 mM IPTG.
Amp stock (100 mg/mL)
1 g of ampicillin
Dissolve in Milli-Q water and bring the final volume up to 10 mL.
Kan stock (25 mg/mL)
0.25 g of kanamycin
Dissolve in Milli-Q water and bring the final volume up to 10 mL.
1 M DTT Stock
1.54 g of DTT
Dissolve in Milli-Q water and bring the final volume up to 10 mL.
Part I: POLRMT Purification
Note: All buffers (except for 5% PEI) are adjusted to pH 7.0 with HCl or NaOH, sterilized with a 0.22 μm filter, and stored at 4 °C. Add BME to the Lysis and HisTrap purification buffers and DTT to the HiTrap heparin purification buffers right before use. Prepare cold, filtered Milli-Q H2O and 20% EtOH to wash and store the FPLC columns after use.
5% PEI solution
Reagent (stock concentration) Final concentration Quantity
PEI (50%) 5% 10 mL
Milli-Q H2O n/a 90 mL
Total 100 mL
Adjust pH to 7.0 with HCl when the solution is cold or on ice. No need to filter.
POLRMT Lysis Buffer
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.9) 40 mM 10 mL
NaCl (3 M) 400 mM 33.3 mL
Glycerol (100%) 15% v/v 37.5 mL
Tween 20 (100%) 0.1% v/v 0.25 mL
EDTA (500 mM) 1 mM 0.5 mL
BME* (14.3 M) 5 mM See Note
Lysozyme* 1 mg/mL See Note
PIC (1,000×)* 1× See Note
Milli-Q H2O n/a 168 mL
Total 250 mL
*Add right before use. Pour 50 mL of the buffer into a conical tube and then add these components (5 mg of lysozyme, 17.5 μL of BME, and 50 μL of PIC) to the tube instead of the entire bottle of buffer. PIC was prepared following instructions online and is used as 1,000× stock: https://www.sigmaaldrich.com/US/en/product/sigma/p2714.
POLRMT HisTrap Buffer A
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.9) 40 mM 40 mL
NaCl (3 M) 300 mM 100 mL
Glycerol (100%) 15% v/v 150 mL
Tween 20 (100%) 0.1% v/v 1 mL
Imidazole 20 mM 1.36 g
BME* (14.3 M) 5 mM 350 μL
Milli-Q H2O n/a 707 mL
Total 1,000 mL
*Add BME right before use.
POLRMT HisTrap Buffer B
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.9) 40 mM 20 mL
NaCl (3 M) 300 mM 50 mL
Glycerol (100%) 15% v/v 75 mL
Tween 20 (100%) 0.1% v/v 0.5 mL
Imidazole 500 mM 17.02 g
BME* (14.3 M) 5 mM 175 μL
Milli-Q H2O n/a 353 mL
Total 500 mL
*Add BME right before use.
POLRMT Heparin No Salt Buffer
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.9) 40 mM 4 mL
NaCl (3 M) 0 mM n/a
Glycerol (100%) 15% v/v 15 mL
Tween 20 (100%) 0.1% v/v 0.1 mL
EDTA (500 mM) 1 mM 0.2 mL
DTT* (1 M) 1 mM 100 μL
Milli-Q H2O n/a 81.6 mL
Total 100 mL
This buffer is used to adjust the salt concentration of the elution from the HisTrap column. *Add DTT right before use.
POLRMT Heparin Buffer A
Reagent (Stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.9) 40 mM 40 mL
NaCl (3 M) 150 mM 50 mL
Glycerol (100%) 15% v/v 150 mL
Tween 20 (100%) 0.1% v/v 1 mL
EDTA (500 mM) 1 mM 2 mL
DTT* (1 M) 1 mM 1,000 μL
Milli-Q H2O n/a 756 mL
Total 1,000 mL
*Add DTT right before use.
POLRMT Heparin Buffer B
Reagent (Stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.9) 40 mM 20 mL
NaCl (3 M) 1,000 mM 167 mL
Glycerol (100%) 15% v/v 75 mL
Tween 20 (100%) 0.1% v/v 0.5 mL
EDTA (500 mM) 1 mM 1 mL
DTT* (1 M) 1 mM 500 μL
Milli-Q H2O n/a 236 mL
Total 500 mL
*Add DTT right before use.
Part II: TFAM purification
Note: All buffers are adjusted to pH 7.5 with HCl or NaOH, sterilized with a 0.22 μm filter, and stored at 4 °C. Prepare cold, filtered Milli-Q H2O and 20% EtOH to wash and store the FPLC columns after use.
TFAM Lysis Buffer
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 20 mM 4 mL
NaCl (3 M) 200 mM 13.33 mL
Glycerol (100%) 15% v/v 30 mL
Imidazole 10 mM 0.136 g
BME* (14.3 M) 5 mM See Note
Lysozyme* 1 mg/mL See Note
Protease inhibitor cocktail* (1,000×) 1× See Note
Milli-Q H2O n/a 152.47 mL
Total 200 mL
*Add right before use. We only need 50 mL of the buffer in a conical tube; then, add these components (5 mg of lysozyme, 17.5 μL of BME, and 50 μL of PIC) to the tube instead of the entire bottle of buffer.
TFAM Ni-NTA Buffer A*
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
NaCl (3 M) 200 mM 66.65 mL
Glycerol (100%) 15% v/v 150 mL
Imidazole 10 mM 0.681 g
BME* (14.3 M) 5 mM 350 μL
Milli-Q H2O n/a 762.35 mL
Total 1,000 mL
*This buffer is the same as the TFAM Lysis Buffer without lysozyme and PIC. Add BME right before use.
TFAM Ni-NTA Buffer B
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
NaCl (3 M) 200 mM 66.65 mL
Glycerol (100%) 15% v/v 150 mL
Imidazole 500 mM 34.04 g
BME* (14.3 M) 5 mM 350 μL
Milli-Q H2O n/a 762.35 mL
Total 1,000 mL
*Add BME right before use.
TFAM TEV Cleavage Buffer
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 20 mM 40 mL
NaCl (3 M) 300 mM 200 mL
Glycerol (100%) 15% v/v 300 mL
BME* (14.3 M) 5 mM 700 μL
Milli-Q H2O n/a 1,459 mL
Total 2,000 mL
*Add BME right before use.
TFAM Heparin Buffer A
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
NaCl (3 M) 200 mM 66.67 mL
Glycerol (100%) 15% v/v 150 mL
DTT* (1 M) 1 mM 1,000 μL
Milli-Q H2O n/a 762.33 mL
Total 1,000 mL
*Add DTT right before use.
TFAM Heparin Buffer B
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 30 mM 30 mL
NaCl (3 M) 1500 mM 500 mL
Glycerol (100%) 15% v/v 150 mL
DTT* (1 M) 1 mM 1,000 μL
Milli-Q H2O n/a 319 mL
Total 1,000 mL
*Add DTT right before use.
TFAM Storage Buffer (2×)
Reagent (stock concentration) Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 80 mM 16 mL
NaCl (3 M) 200 mM 13.33 mL
DTT* (1 M) 2 mM 400 μL
Milli-Q H2O n/a 170.27 mL
Total 200 mL
*Add DTT right before use.
10× TFAM Binding Buffer
500 mM Tris-HCl pH 8.0
1 M KCl
50 mM MgCl2
1 mM DTT (add right before use)
TFAM EMSA Running Buffer
50 mM Tris-acetate pH 8.0
2.5 mM EDTA
Part III: TFB2M purification
Note: All buffers are adjusted to pH 8.0 with HCl or KOH, sterilized with a 0.22 μm filter, and stored at 4 °C. Use BME for the Lysis and HisTrap buffers and DTT for the HiTrap Heparin column and Storage buffers. Prepare cold, filtered Milli-Q H2O and 20% EtOH to wash and store the FPLC columns after use.
TFB2M Lysis Buffer
Reagent (stock concentration) Final concentration Quantity
HEPES-KOH (1 M, pH 8.0) 20 mM 6 mL
KCl (3 M) 1 M 100 mL
Glycerol (100%) 5% v/v 15 mL
Imidazole 10 mM 0.204 g
BME* (14.3 M) 5 mM See Note
Lysozyme* 1 mg/mL See Note
Protease inhibitor cocktail* (PIC, 1,000×) 1× See Note
Milli-Q H2O n/a 179 mL
Total 300 mL
*Add right before use. We only need 50 mL of the buffer in a conical tube; then, add these components (5 mg of lysozyme, 17.5 μL of BME, and 50 μL of PIC) to the tube instead of the entire bottle of buffer.
TFB2M Ni-NTA Buffer A
Reagent (stock concentration) Final concentration Quantity
HEPES-KOH (1 M, pH 8.0) 20 mM 20 mL
KCl (3 M) 300 mM 100 mL
Glycerol (100%) 5% v/v 50 mL
Imidazole 20 mM 1.36 g
BME* (14.3 M) 5 mM 350 μL
Milli-Q H2O n/a 830 mL
Total 1,000 mL
*Add BME right before use.
TFB2M Ni-NTA Buffer B
Reagent (stock concentration) Final concentration Quantity
HEPES-KOH (1 M, pH 8.0) 20 mM 10 mL
KCl (3 M) 300 mM 50 mL
Glycerol (100%) 5% v/v 25 mL
Imidazole 500 mM 17.02 g
BME* (14.3 M) 5 mM 175 μL
Milli-Q H2O n/a 415 mL
Total 500 mL
*Add BME right before use.
TFB2M TEV Cleavage Buffer
Reagent (stock concentration) Final concentration Quantity
HEPES-KOH (1 M, pH 8.0) 20 mM 40 mL
KCl (3 M) 300 mM 200 mL
Glycerol (100%) 5% v/v 100 mL
BME (14.3 M) * 5 mM 699.3 μL
Milli-Q H2O n/a 1,659.3 mL
Total 2,000 mL
*Add BME right before use.
TFB2M No Salt Heparin Buffer
Reagent (stock concentration) Final concentration Quantity
HEPES-KOH (1 M, pH 8.0) 20 mM 4 mL
Glycerol (100%) 5% v/v 10 mL
EDTA (500 mM) 1 mM 0.4 mL
DTT (1 M) * 1 mM 0.2 mL
Milli-Q H2O n/a 185.4 mL
Total 200 mL
*Add DTT right before use.
TFB2M Heparin Buffer A
Reagent (stock concentration) Final concentration Quantity
HEPES-KOH (1 M, pH 8) 20 mM 20 mL
KCl (3 M) 200 mM 66.67 mL
Glycerol (100%) 5% v/v 50 mL
EDTA (500 mM) 1 mM 2 mL
DTT (1 M) * 1 mM 1 mL
Milli-Q H2O n/a 860.33 mL
Total 1,000 mL
*Add DTT right before use.
TFB2M Heparin Buffer B
Reagent (stock concentration) Final concentration Quantity
HEPES-KOH (1 M, pH 8.0) 20 mM 10 mL
KCl (3 M) 1.5 M 250 mL
Glycerol (100%) 5% v/v 25 mL
EDTA (500 mM) 1 mM 1 mL
DTT (1 M) * 1 mM 0.5 mL
Milli-Q H2O n/a 213.5 mL
Total 500 mL
*Add DTT right before use.
SDS-PAGE sample recipes
Note: These recipes make 20 μL of each SDS-PAGE sample. Boil samples for 5 min at 95 °C before running and load 9–12 μL per lane. All POLRMT samples can be run on 8%–10% SDS-PAGE. For TFAM and TFB2M, 10%–12% SDS-PAGE would be suitable. We run the gels in 1× Tris/Glycine/SDS buffer and stain with Coomassie.
Uninduced, induced, pellet, lysate, PEI pellet, PEI supernatant, AS pellet, Ni-NTA or HisTrap flowthrough, and wash samples
1.5 μL of sample or scrape a small amount of pellet directly into the microfuge tube
13.5 μL of Milli-Q water
5 μL of 4× Laemmli buffer
Ni-NTA Elution samples, AS supernatant
7.5 μL of sample or small piece of pellet
7.5 μL of Milli-Q water
5 μL of 4× Laemmli buffer
TEV-cleaved sample, HisTrap, and Heparin column fractions
Note: Load as much as possible (at least 12 μL per lane) to visualize very diluted samples
10 μL of sample or small piece of pellet
5 μL of Milli-Q water
5 μL of 4× Laemmli buffer
Laboratory supplies
17 mm × 100 mm culture tubes, 14 mL (VWR, catalog number: 60818-689)
BD disposable syringe, 50 mL (Fisher Scientific, catalog number: 13-689-8)
Biotix 10 μL tips (Fisher Scientific, catalog number: 12-111-021)
Fernsbach culture flasks, 2.8 L (Fisher Scientific, catalog number: 09-552-39)
Membrane filters, 0.22 μm pore size, 47 mm diameter (Fisher Scientific, catalog number: 09-719-2B)
Petri dishes (Fisher Scientific, catalog number: FB0875713)
Pipette tips, microfuge tubes, and low-retention tubes (Genesee Scientific, catalog numbers: 24-282, 23-150RL, 23-165RL, 28-103, 22-281LR)
Serological pipettes, 5, 10, and 25 mL (VWR, catalog numbers: 75816-094, 75816-100, 75816-090)
Syringe filters (0.22 μm pore size, 25 mm diameter) (VWR, catalog number: 76479-024)
Equipment
1 L centrifuge bottles with sealing closure (Thermo Scientific, catalog number: 31411006)
4 L capacity centrifuge (Thermo Scientific, Sorvall LYNX 4000 Superspeed Centrifuge, catalog number: 75006580)
Benchtop centrifuges (Eppendorf, 5910 R, 5425)
Benchtop incubator shaker (Eppendorf, model: New Brunswick Benchtop Incubator Shaker I24)
BioSpectrometer Basic (Eppendorf)
Dialysis tubing, 14,000 MWCO (Sigma-Aldrich, catalog number: D9777-100FT)
Fast protein liquid chromatography (FPLC) system (Bio-Rad, NGC Quest 10 Chromatography System, catalog number: 7880001)
Gel imager (Bio-Rad, model: Gel Doc EZ Imager)
Glass column for Ni-NTA resin (Bio-Rad, Econo-Column Chromatography Column, catalog number: 7371512)
Hot plate stirrer (VWR, catalog number: 97042-674)
Magnetic stir bar (Fisher Scientific, catalog number: 14-513-51)
Reusable bottle top filter unit (Thermo Scientific, Nalgene, catalog number: DS0320-2533)
SDS-PAGE running system (Bio-Rad, Mini-PROTEAN Tetra System)
Sonicator (Qsonica, CL-18 Sonicator Probe)
Software and datasets
Bio-Rad Image Lab (https://www.bio-rad.com/en-us/product/image-lab-software?ID=KRE6P5E8Z)
ImageJ version 1.53e (https://imagej.net/ij/index.html)
Procedure
Part I: POLRMT Purification
Overexpression of recombinant POLRMT in E. coli
Transform the plasmid into BL21-CodonPlus (DE3)-RIPL and plate on LB agar plates containing ampicillin (or the appropriate antibiotic). Incubate inverted overnight at 37 °C.
Note: This transformation can also be done with BL21-CodonPlus-RIL. This step can be skipped if glycerol stocks of cells carrying the POLRMT-coding plasmid have been made previously.
Get an autoclaved flask with 50 mL of autoclaved LB media and add 50 μL of Amp stock (100 mg/mL, see Recipes).
Pick five separate colonies from the plate with a sterile pipette tip or stick for inoculation (or add a small piece of glycerol stock).
Once inoculated, start incubating the cultures in the afternoon around 5 pm at 28 °C and shake overnight (14–17 h) at 200 rpm (see General note 2 for temperature and oversaturation of cells).
In four flasks (2.8 L Fernsbach culture flasks), prepare 1 L each of autoclaved LB media and add 1 mL of Amp stock per liter of media.
Add 10 mL of inoculum per liter of LB media with ampicillin (see General note 3 for how to make glycerol stocks).
Incubate the inoculated flasks at 37 °C with shaking at 200 rpm. Check cell density by measuring the OD600 after 3–4 h to see if it is between 0.4 and 0.6.
Once in the above OD range, lower the shaker temperature to 18 °C. Withdraw a 1 mL sample (label as “Uninduced”) into a microfuge tube and tape it on the side of the flask to let it continue shaking.
Cool down the flasks to 18 °C from 30 min to 1 h before adding IPTG (0.2 mM final concentration) to induce POLRMT expression. To do this, add 200 μL of 1 M IPTG stock (see Recipes) per 1 L of LB (see General note 4 for cooling flasks).
Continue growing overnight (16–20 h) at 18 °C at 200 rpm.
Withdraw a 1 mL sample at the end of the induction period (“Induced”) and remove the “Uninduced” sample tube off the side of the flask (see Recipes for SDS-PAGE samples). Run out samples on a 10% SDS-PAGE gel and Coomassie stain to visualize POLRMT overexpression.
Harvest the cells by centrifugation at 2,991× g for 30 min at 4 °C in 1 L centrifuge bottles.
Safely dispose of the supernatant with 10% bleach and scrape the pellets into a 50 mL conical tube. Store the combined pellet at -80 °C (flash freezing is not necessary) or proceed with the next step.
Lysis of POLRMT overexpression pellet by sonication
Note: All steps below should be done at 4 °C or on ice unless noted otherwise.
Thaw the pellet by putting the tube in tap water. Scrape out the cells with a spatula into a metal beaker and add 50 mL of cold Lysis Buffer with BME, lysozyme, and PIC. Optional: Stir on a stir plate with a stir bar at 4 °C for 1 h to let the lysozyme break down cells.
Put the beaker in an ice bucket and sonicate cell suspension for 30 min at 4 °C (30 s on and off, 60% amplitude). Check that the suspension has become less viscous and slightly darker at this point.
Centrifuge the suspension at 30,000× g for 1 h at 4 °C.
Pour supernatant (“Lysate”) into a cold beaker on ice and record the volume.
Take samples to run on an SDS-PAGE gel: Pellet and Lysate (see Recipes). There will be more POLRMT in the pellet than in the lysate, but we will use the soluble POLRMT in the lysate for purification (Figure 1B).
Figure 1. Mitochondrial RNA polymerase (POLRMT) purification flowchart and polyethyleneimine (PEI) and ammonium sulfate (AS) precipitation. (A) Expression construct containing the nucleotide sequence for POLRMT residues 43–1,230 (Uniprot: O00411) and an N-terminal 6x-His-tag. (B) 10% SDS-PAGE analysis of samples from the sonication step, PEI precipitation, and AS precipitation steps. (C) Schematic of POLRMT purification including sonication, PEI precipitation, AS precipitation, a DEAE column, a HisTrap column, a HiTrap heparin column, and concentration.
POLRMT PEI precipitation
With the lysate in a cold glass beaker on ice or at 4 °C, add a stir bar and check that it gently stirs the solution while on a stir plate.
Slowly add cold 5% PEI (see Recipes) dropwise into the lysate to get 0.3% PEI final concentration. To calculate how much 5% PEI we need, find 6% of the volume of the lysate:
______ mL of 5% PEI to be added = ______ mL of lysate × 0.06
Cover the top of the beaker with aluminum foil and keep stirring gently at 4 °C for an additional 30 min.
Centrifuge at 30,000× g for 10 min at 4 °C.
Take out samples for gel analysis [PEI pellet, PEI supernatant (see Recipes)] and run them out on a 10% SDS-PAGE gel. POLRMT should be in the supernatant (Figure 1B).
POLRMT AS precipitation
Add AS powder slowly (for a period of 30 min) to the supernatant to get 55% saturation.
For example, 55% saturation of 100 mL of lysate requires 35.1 g of AS.
______mL lysate × 0.351 g/mL = _______ grams AS to be added
Cover the top of the beaker with aluminum foil and gently stir at 4 °C for 1 h (or overnight) to dissolve AS and precipitate proteins.
Centrifuge at 30,000× g for 10 min at 4 °C. The POLRMT protein is in the pellet.
Take out samples for gel analysis: AS pellet, AS supernatant (see Recipes). On an SDS-PAGE gel, you should see POLRMT (~140 kDa) in the AS pellet and not in the AS supernatant (Figure 1B).
Record the weight of the pellet in grams and store it at -80 °C.
Pause point: The AS pellet can be stored for a few months before moving on to FPLC purification.
POLRMT DEAE gravity column and overnight HisTrap column
Note: One column volume (CV) corresponds to 5 mL for the HisTrap column (Cytiva). The DEAE gravity column can be replaced by using a HiTrap DEAE Fast Flow column (Cytiva) attached in tandem with the HisTrap column.
Start by attaching the 5 mL HisTrap column and loading POLRMT HisTrap buffers (see Recipes) onto the FPLC.
Wash and equilibrate the column (2 mL/min for five CV each of filtered Milli-Q water, HisTrap Buffer B, and then HisTrap Buffer A).
Resuspend the AS pellet in the HisTrap Buffer A by adding 4 mL of cold buffer per gram of pellet. Stir gently until dissolved.
Note: The following steps with the DEAE column can be done at room temperature as long as all the buffers are cold and the samples are kept on ice.
Prepare the DEAE by placing 10 mL of the DEAE resin suspension into a clean gravity column. Once the resin settles, you should get approximately 5 mL of bed resin.
Wash the DEAE with 5× bed volume of cold filtered water and cold HisTrap Buffer A.
Note: This DEAE column can be washed with water and stored in 20% EtOH for reuse.
With the stopper on the gravity column closed, pour the dissolved AS pellet into the column and gently stir with a spatula. Optional: Incubate for 15 min at 4 °C with gentle rocking.
Elute into a clean container on ice and then add 10 mL of HisTrap Buffer A to ensure all POLRMT is washed out.
Filter the eluent with a 0.22 μm syringe filter and prepare to load it on the HisTrap column on the FPLC. Use these settings for the FPLC run while monitoring eluent absorbance at 280 nm (A280) and collect all flowthrough/eluent for a later SDS-PAGE analysis (Figure 2B):
Sample application: 0.3 mL/min (for running overnight).
Column wash with Buffer A: 0.5 mL/min (for running overnight). Wash the column until the A280 trace returns to baseline (approximately five CV or 25 mL). After reaching baseline, continue to wash for at least 20 mL.
Protein elution: 1 mL/min, collect 5 mL fractions.
Elution program:
0–50% Buffer B gradient for 40 CV (200 mL). Optional: Wash the column with Buffer B. No need to collect samples from these steps.
70% Buffer B for five CV (25 mL)
100% Buffer B for five CV (25 mL)
Wash the column with five CV of water and five CV of 20% EtOH (use a lower flow rate of 1 mL/min) and store the columns.
Note: POLRMT elutes in a small broad peak and is very dilute. Do not let the fractions sit for longer than one day, as the more concentrated fractions of POLRMT will start precipitating (see General note 5).
Figure 2. Mitochondrial RNA polymerase (POLRMT) HisTrap chromatography. (A) Chromatogram for the elution phase of the HisTrap column run on the FPLC and (B) 8% SDS-PAGE analysis of the collected fractions.
Take out samples of the HisTrap fractions and run them on a 10% SDS-PAGE gel alongside a protein ladder. POLRMT appears around 140 kDa on an SDS-PAGE gel and is more concentrated roughly around fractions #4–22 (Figure 2B). (To save time, start preparing the filters and concentrate while the gel is running; see step below.)
Equilibrate two 50K MWCO Amicon filters at 4,000× g and 4 °C.
Concentrate POLRMT by centrifuging the Amicon filtration units at 1,300× g. Pipette solution gently every 5–10 min to avoid precipitation. Stop when the total volume becomes less than 25 mL (see General note 6 about this concentration step).
Add an equal volume of No Salt Heparin Buffer to the concentrate to bring the final NaCl concentration to 150 mM (see General note 7).
POLRMT overnight HiTrap heparin column
Wash and equilibrate the 5 mL HiTrap heparin column in Buffer A on the FPLC (2 mL/min for five CV each of filtered Milli-Q water, Heparin Buffer B, and then Heparin Buffer A).
Load your protein sample that has been diluted with the No Salt Heparin buffer.
Run the following FPLC program:
Sample application: 0.3 mL/min (for running overnight).
Column wash with Buffer A: 0.5 mL/min (for running overnight). Wash the column until the A280 trace returns to baseline (approximately five CV or 25 mL). After reaching baseline, continue to wash for at least 20 mL.
Protein elution: 2 mL/min, collect 3 mL fractions.
0–50% Buffer B gradient for 20 CV (100 mL).
55% Buffer B for five CV or 25 mL (POLRMT elutes here!).
70% Buffer B for five CV or 25 mL.
100% Buffer B for five CV or 25 mL.
Wash the column with five CV of water and five CV of 20% EtOH (use a lower flow rate of 1 mL/min) for storage of the columns.
Concentration and storage of POLRMT
Run a 10% SDS-PAGE gel of the fractions (see Recipes) and pool together the fractions with POLRMT (Figure 3B).
Figure 3. Mitochondrial RNA polymerase (POLRMT) HiTrap Heparin chromatography. (A) Chromatogram for the elution phase of the HiTrap Heparin column run on the FPLC and (B) 10% SDS-PAGE analysis of the collected fractions.
Equilibrate the membrane of a 50K MWCO Amicon filtration unit with 150 mM NaCl Heparin Buffer by spinning in the centrifuge for 5 min. Add pooled POLRMT fractions to the unit.
Start centrifuging at 2,500× g at 4 °C and gradually decrease the speed to 1,900× g, pipetting gently from the bottom of the membrane and checking concentration frequently until at least 7 μM. POLRMT will easily precipitate out, so if you notice precipitate forming or if the concentration in the solution is not increasing, stop concentrating (see General note 8).
Measure and calculate protein concentration using these parameters by the Protparam tool (https://web.expasy.org/protparam/):
6x-His-POLRMT: 137.7 kDa or 137,700 mg/mole
Molar extinction coefficient: 143,700 L (cm*M)-1 (assuming all cys reduced)
Abs 0.1% = 1.043 (assuming all cys reduced)
Add sterile glycerol to get a final concentration of 25% glycerol.
Aliquot 10 μL into low-retention tubes, flash freeze in liquid nitrogen, and store at -80 °C.
Part II: TFAM purification
Overexpression of recombinant TFAM in E. coli
Transform BL21-CodonPlus(DE3)-RIPL cells using ~200 ng of ProEX_Htb_TFAM plasmid and plate on LB agar plates containing ampicillin. Incubate inverted overnight at 37 °C.
Grow 5 × 5 mL starter cultures overnight in LB supplemented with ampicillin, one colony per tube.
In the morning, spin down the 5 mL starter cultures at 2,782× g for 6 min and pour off the media. Carefully resuspend the cells in a total of 1 mL of LB, transferring the 1 mL to resuspend all the cells from the five tubes.
Add ampicillin to LB media (1 mL of 100 mg/mL ampicillin stock per 1 L of LB). Evenly split the 1 mL cell suspension between the 1 L LB flasks.
Incubate at 37 °C at 200 rpm. Check the OD600 after 4–5 h to see if it is between 0.4 and 0.6.
Once in the above OD range, withdraw a 1 mL sample (“uninduced”) into a microfuge tube, tape the tube on the side of the flask, and lower the shaker temperature to 16 °C.
Cool down the flasks before inducing with IPTG (0.2 mM final concentration; add 200 μL of 1 M IPTG stock per 1 L of LB).
Continue shaking overnight at 16 °C (~16–20 h) at 200 rpm.
Take a sample at the end of the induction period (“Induced”) and run it alongside the “Uninduced” sample (see Recipes) on an SDS-PAGE gel to visualize protein expression.
Harvest the cells by centrifuging at 2,991× g for 15 min at 4 °C.
Dispose of the supernatant and scrape the pellets into a 50 mL conical tube. Store the combined pellet at -80 °C or proceed with the cell lysis.
Cell lysis: lysozyme treatment and sonication
Note: All steps below should be done at 4 °C or on ice unless noted otherwise.
Thaw cells if frozen. Add 40 mL of TFAM Lysis Buffer (with lysozyme, BME, and PIC; see Recipes) to the pellet (10 mL of lysis buffer per liter of expressed culture) and resuspend the pellet until homogenous.
Incubate for 1 h at 4 °C.
Sonicate cell suspension for 20 min at 4 °C (20 s on and off, 60% amplitude).
Centrifuge at 30,000× g for 1 h at 4 °C.
Save ~10 μL of “Cell Pellet” and “Lysate” samples (see Recipes) for SDS-PAGE gel analysis.
TFAM Ni-NTA resin #1
Note: This is Ni-NTA by centrifugation method at 4 °C.
Prepare a 5 mL Ni-NTA resin bed in a 50 mL conical tube according to the manufacturer’s instruction and wash three times with Ni-NTA Buffer A by resuspending the resin in the buffer, spinning it down at 363× g for 3 min at 4 °C, carefully decanting the buffer, and repeating the process.
Add the supernatant and incubate overnight at 4 °C with gentle agitation.
Spin down the resin, collect the supernatant as “Sample Application” into a conical tube, and begin resin washing steps by adding 5 mL of Ni-NTA Buffer A to the resin, gently mixing, and spinning the sample at 1,932× g at 4 °C.
Collect the supernatant as “Wash 1” and repeat washes to wash #10.
Beginning with wash #10, check the absorbance of the sample at 280 nm and continue the washes until the absorbance is close to 0.
Elute the proteins by adding 5 mL of Ni-NTA Buffer B and spinning as before. Collect the sample as “Elution 1” and repeat this 3–5 times, keeping “Elution” supernatants separate.
Run a 10% or 12% SDS-PAGE gel with collected samples from the cell lysis, sample application, washing, and elution (see Recipes). A TFAM band just above 25 kDa should be obvious but some contaminants will still be present (Figure 4B).
Figure 4. Transcription factor A, mitochondrial (TFAM) purification flowchart, Ni-NTA and HiTrap Heparin chromatography, and TFAM DNA-compaction test. (A) Schematic of TFAM purification including Ni-NTA Resin purification, cleavage by TEV protease, a second Ni-NTA resin step, a HiTrap Heparin column, and concentration. (B) 12% SDS-PAGE analysis of Ni-NTA #1 results including washes (W) and elution steps (E). (C) 12% SDS-PAGE analysis of TEV protease cleavage of TFAM and Ni-NTA resin step. (D) Chromatogram for the elution phase of the HiTrap Heparin column run on the FPLC and the SDS-PAGE analysis of the collected fractions. (E) Electrophoretic mobility shift assay (EMSA) to test TFAM ability to compact plasmid DNA.
Dialysis and TEV cleavage of the His tag on TFAM
Add 2 L of ice-cold TFAM TEV Cleavage Buffer to a large beaker.
Place the elution samples with significant TFAM content into dialysis tubing and add TEV protease at a 1:30 TEV:TFAM protein molar ratio. (Estimate TFAM content using the absorbance of the pooled elution samples at 280 nm. See step G for protein mass and extinction coefficient.)
Allow the sample to dialyze in Cleavage Buffer and cleave overnight at 4 °C with gentle stirring.
Check the success of TEV cleavage on a 12% SDS-PAGE gel by comparing the sample before and after dialysis. If cleavage is complete, continue to the next step. If not, add more TEV protease and continue dialysis (Figure 4C).
Note: The version of TEV protease used in our lab is ~26 kDa in size. This means it will look a lot like uncleaved TFAM on the SDS-PAGE gel.
TFAM Ni-NTA resin #2
Prepare a 5 mL resin bed of Ni-NTA resin as before (see step C1).
Add the dialyzed sample to the resin and incubate overnight at 4 °C as before with gentle agitation.
Spin down the resin as before (1,932× g at 4 °C) and collect the supernatant [“Sample application” (this will have the TEV-cleaved, now untagged TFAM)].
Add 5 mL of Ni-NTA Buffer B to the resin, mix, and spin to collect “Elution 1.” You want to collect this to be sure the TEV protease was bound by the Ni resin.
Repeat the elution 2–3 times.
Run a 10% or 12% SDS-PAGE gel to be sure that TFAM is in the “Sample application” (Figure 2C).
TFAM concentration and HiTrap heparin column
Note: An alternative to the first concentration step is to adjust the salt concentration by adding a version of the Ni-NTA Buffer A without any salt. Adjust the salt to <200 mM for use on the HiTrap heparin column.
Wash and equilibrate the 5 mL HiTrap heparin column (Cytiva) on the FPLC with five CV of heparin Buffer B and then Buffer A.
Add elution fractions from Ni-NTA resin #2 to a 50 mL conical tube and begin buffer exchanging the sample by adding it in 15 mL volumes to a 3K MWCO Amicon filter and spinning at 3,434× g at 4 °C. Stop concentrating when the buffer exchange sample is down to 2 mL.
When the entire protein sample has been added and spun down to approximately 2 mL, fill the 3K MWCO Amicon filter with Heparin Buffer A and continue to spin. Continue to add Heparin Buffer A and spin until ~30 mL has flown through the filter and you are left with 2–3 mL. This lowers the salt concentration from 300 mM to 200 mM to ensure binding to the HiTrap Heparin column.
Load the buffer-exchanged protein sample onto the prepared HiTrap Heparin column.
Run using these settings:
Sample application: 1.0 mL/min.
Column wash: 1.0 mL/min until A280 reaches ~0.
Elution: 2.5 mL/min with 2 mL fractions.
0–100% Buffer B gradient over 10 CV (50 mL).
Wash the column with five CV of water and five CV of 20% EtOH (1 mL/min) for storage.
Concentration and storage of TFAM
Run a 10% or 12% SDS-PAGE gel of the fractions and pool together the fractions with TFAM (Figure 4D)
Concentrate as before using a fresh 3K MWCO Amicon filter.
When all the pooled fractions have been concentrated to a small volume (0.5–1 mL), add 2× Storage Buffer to fill the filter and continue to centrifuge. Repeat this process to pass two full volumes of Storage Buffer through the filter to exchange the buffer. Stop when the second full pass of 2× Storage buffer has reached ~2 mL and begin measuring the absorbance at 280 nm for TFAM concentration while continuing to centrifuge.
Measure and calculate protein concentration using these parameters from the protein sequence by the Protparam tool (https://web.expasy.org/protparam/):
TFAM after TEV cleavage: 24.4 kDa or 24,400 mg/mole.
Molar extinction coefficient: 35,410 L (cm*M)-1 (assuming all cys reduced).
Abs 0.1% = 1.45 (assuming all cys reduced).
When the desired concentration has been reached (100–200 mM), move the sample from the Amicon filter to a conical tube and add an equal volume of filtered 50% glycerol for a final concentration of 25% glycerol.
Aliquot 20 mL per tube, flash freeze, and store at -80 °C.
Testing TFAM DNA-compaction activity with electrophoretic mobility shift assay (EMSA)
Note: To know if the purified TFAM is active, it is best to run an electrophoretic mobility shift assay. TFAM can bind DNA non-specifically, so any plasmid DNA can be used. If the bands shift upwards in the assay, the TFAM is active and will work with transcription assays. In our assays, we use the Puc19 plasmid.
Prepare a 1% agarose EMSA gel using TFAM EMSA Running Buffer
Prepare reactions with 1× TFAM Binding Buffer, add ~200 ng of plasmid DNA, and add TFAM in increasing amounts to the plasmid. These reactions are typically done in total volumes of 20–30 mL.
Incubate samples at 37 °C for 30 min.
Add filtered glycerol to 10% in the samples and load onto the EMSA gel along with a 1 kb DNA ladder.
Run the EMSA gel at 70 V for 65 min at room temperature in TFAM EMSA Running Buffer.
Stain the gel with gentle agitation using ethidium bromide for 30 min at room temperature and image using UV light on a Bio-Rad Gel Doc EZ Imager system or equivalent (Figure 4E).
Part III: TFB2M purification
Expression of TFB2M
Transform the plasmid containing TFB2M (Figure 5) into BL21-CodonPlus (DE3)-RIPL and spread on Kan-containing (25 μg/mL, see Recipes) plates. Incubate inverted overnight at 37 °C.
Note: The transformation step can be skipped if glycerol stocks have been previously made. E. coli strain ArcticExpress (DE3) could also be used for transformation.
Figure 5. Transcription factor B2, mitochondrial (TFB2M) purification flowchart. (A) Expression construct containing the nucleotide sequence for TFB2M, corresponding to residues 21–396 (Uniprot: Q9H5Q4). This plasmid has an N-terminal 6x-His-tagged Maltose-binding protein (MBP) solubility tag connected to TFB2M with a TEV-cleavage site. (B) Schematic of TFB2M purification including protein expression, Ni-NTA #1 Resin purification, cleavage by TEV protease, Ni-NTA #2 Resin step, a HiTrap Heparin column, and concentration.
Add five colonies or a small chunk of the frozen glycerol stock to 5–10 mL of LB culture with Kan (25 μg/mL). Grow overnight while shaking at 28–37 °C; avoid letting the starter culture become oversaturated (see General note 2 for POLRMT purification).
In the morning, add primary culture (2.5–5 mL) to 1 L of LB culture.
Grow at 37 °C at 200 rpm until OD600 reaches 0.7–0.8 (approximately 3–5 h).
Once in the OD range, cool down the shaker to 14 °C and withdraw a 1 mL sample (“uninduced”) before inducing the rest of the culture with 0.2 mM final IPTG (refer to Recipes). Continue to shake for 16–20 h at 14 °C.
Take a 1 mL sample (“Induced”) at the end of the induction period to run on an SDS-PAGE gel.
Harvest the cells by centrifuging at 2,991× g for 15 min at 4 °C.
Store cell pellet at -80 °C.
Sonication
Note: All steps below should be done at 4 °C or on ice unless noted otherwise.
Add 40 mL of Lysis Buffer (with lysozyme, BME, and PIC, see Recipes) to 4 L-worth of cell pellet (10 mL of lysis buffer per liter of expressed culture) and stir the pellet at 4 °C for 1 h.
In the cold room, sonicate for 20 min at 60% amplitude, 30 s on and off.
Centrifuge at 30,000× g for 1 h at 4 °C to remove cell debris.
Ni-NTA Resin #1
Set up a gravity column by clamping it onto a stand and pipette 10 mL of Ni-NTA resin solution into the column to get 5 mL bed volume.
Wash the resin with filtered, cold Milli-Q water three times and then wash once to equilibrate with your Ni Buffer A.
Note: The volume of each wash should be five times the bed volume of the resin.
Pour the cell lysate onto Ni-NTA resin and gently stir with a metal spatula to promote the binding of your His-tagged protein. Securely close the column/vial and rock gently for 1 h at 4 °C.
Clamp the column again on the stand and collect your sample application flowthrough. Label the sample as “Ni flowthrough” for SDS-PAGE.
Prepare the following mixtures of Buffer A and B to perform the wash and elution steps:
One CV = 5 mL bed resin.Wash 1, 2, and 3: 0% B, five CV: prepare three tubes containing 50 mL of Buffer A each.
Wash 4: 5% B, five CV: one tube containing 47.5 mL of Buffer A and 2.5 mL of Buffer B.
Elution 1: 40% B, two CV: one tube containing 6 mL of Buffer A and 4 mL of Buffer B.
Elution 2: 50% B, two CV: one tube containing 5 mL of Buffer A and 5 mL of Buffer B.
Elution 3: 100% B, two CV: one tube with 10 mL of Buffer B.
Keep them on ice or at 4 °C before use.
Note: The washes and elution can be done at room temperature if all the buffers are cold and samples are kept on ice. Avoid drying out the resin.
Wash the resin with Wash 1. Collect and label the flowthrough as “W1.”
Repeat with Wash 2 and Wash 3.
Wash a final time with Wash 4 (which contains 5% v/v of Buffer B in Buffer A). Save the flowthrough as “W4.”
Elute TFB2M with Elution 1. Collect and label the elution as “E1.”
Repeat with Elution 2 and 3.
Pool elution 1, 2, and 3 together and estimate the protein concentration with the spectrophotometer:
6x-His-MBP-TFB2M: 87.7 kDa or 87,691 mg/mole.
Molar extinction coefficient: 117,230 L (cm·M)-1 (assuming all cys reduced).
Abs 0.1% = 0.00134 (assuming all cys reduced).
Overnight TEV cleavage
Estimate the amount of TFB2M using the absorbance of the pooled elution samples at 280 nm:
Amount of total TFB2M (spectrometer reading) =_____mg/mL
Total volume = ____ mL
Total amount of protein = ____mg (assume it is all TFB2M)
We need a molar ratio of 1:25 TEV:TFB2M
Amount of TEV to be added =_____mg
Concentration of your TEV protease stock = _____mg/mL
Volume of TEV to be added = _____mL.
Add 2 L of ice-cold TEV Cleavage Dialysis Buffer to a large beaker with a large stir bar.
Add TEV to the same tube as your Ni-resin eluted TFB2M, gently mix, and then transfer into the dialysis tubing (see General note 9).
Allow the sample to dialyze in the TEV Cleavage Buffer for 19–20 h overnight at 4 °C with gentle stirring.
Check the completion of TEV cleavage of TFB2M on a 10% or 12% SDS-PAGE gel by comparing the TFB2M band size before (~88 kDa) and after dialysis (~43 kDa). If cleavage is complete, continue to the next step. If not, add more TEV protease and continue dialysis (Figure 6).
Note: The MBP tag (42 kDa) will be similar in size to TFB2M, and the band may be more intense. The version of purified TEV protease used in our lab is approximately 26 kDa in size.
Figure 6. Transcription factor B2, mitochondrial (TFB2M) Ni-NTA chromatography. (A) Flowchart of the Ni-NTA #1, TEV cleavage, and Ni-NTA #2 steps. (B) SDS-PAGE of TFB2M expression, the first Ni-NTA purification step, the TEV cleavage process, and TFB2M presence in Ni-NTA #2 wash.
Ni-NTA Resin #2
Note: This is to remove His-tagged TEV protease and other contaminants from the cleaved TFB2M, which is now missing its His-tag. TFB2M will be in the flowthrough instead of the elution (Figure 6A).
Using the same Ni-NTA column from the previous step, wash the column with 5× bed volume of Buffer B to remove all the bound proteins, and then wash with Buffer A to equilibrate the column.
Before applying your cleaved TFB2M sample onto the Ni-NTA resin, add Buffer B to the TFB2M solution to bring the imidazole concentration to 20 mM (matching that in Buffer A):
(20 mM imidazole needed) (____ mL of cleaved TFB2M sample) = (500 mM imidazole in Buffer B) (x mL Buffer B needed)
Note: Introducing a small (20 mM) amount of imidazole ensures that most of TFB2M does not bind to the resin. The other protein contaminants like the His-MBP-tag and His-tagged TEV protease will stay bound to the Ni-NTA.
Add the sample to the Ni resin and rock gently for 30 min at 4 °C.
Collect the flowthrough (or “wash”) containing TFB2M and run a 10% or 12% SDS-PAGE gel to be sure that TFB2M is present in the flowthrough (Figure 6B).
Note: What remains bound to the column are the His-MBP tag and His-tagged TEV protease. Be aware that the His-MBP tag is similar in size to untagged TFB2M (approximately 42 kDa), and its band is often more intense on a gel.
Overnight heparin column
Wash and equilibrate a 5 mL HiTrap heparin column (Cytiva) on the FPLC.
Dilute the TFB2M sample (currently in 300 mM KCl buffer) with the No Salt Heparin Buffer to get a final salt concentration between 75 and 150 mM KCl.
Note: The lower the salt, the more TFB2M binds to the heparin column. Do not concentrate TFB2M before loading it on the column or it will precipitate out.
Run using these settings:
Note: One column volume (CV) is 5 mL for the HiTrap Heparin column (Cytiva).
Sample application: 1 mL/min.
Column wash: 0.5 mL/min (for running overnight) for five CV.
Protein elution: 0.5 mL/min, collect 2 mL fractions. Elution program:
0–70% gradient for 10 CV (50 mL total)
100% for one CV
Wash the column with five CV of water and five CV of 20% EtOH (1 mL/min) for storage.
Collect fractions and run a sample on a 10%–12% SDS-PAGE to determine which fractions contain TFB2M (Figure 7B).
Figure 7. Transcription factor B2, mitochondrial (TFB2M) HiTrap Heparin chromatography. (A) Chromatogram for the elution phase of the HiTrap Heparin column run on the FPLC and (B) a 12% SDS-PAGE analysis of the collected fractions.
TFB2M concentration
Pool together the fractions containing TFB2M.
Equilibrate a 10K MWCO Amicon membrane in a centrifuge for 5 min with the Heparin Buffer A.
Start centrifuging the TFB2M sample at 1,300× g. Stop the centrifuge every 10 min and gently pipette TFB2M solution up and down with a pipette to mix. If you notice precipitate forming or if the concentration in the solution is not increasing, stop concentrating.
Measure and calculate the protein concentration using these parameters from the protein sequence by the Protparam tool (https://web.expasy.org/protparam/):
TFB2M after TEV cleavage: 43.405 kDa or 43,405 mg/mole.
Molar extinction coefficient: 49,390 L (cm·M)-1 (assuming all cys reduced).
Abs 0.1% = 1.138 (assuming all cys reduced).
Add sterile glycerol to get a final concentration of 25%, aliquot 15 μL into low-retention tubes, flash freeze in liquid nitrogen, and store at -80 °C (see General note 10 for estimating salt concentration in TFB2M samples).
Data analysis
To assess the purity of the proteins, run the purified samples on an SDS-PAGE gel (see Figure 8A below). Protein band intensity can be analyzed using Bio-Rad Image Lab and ImageJ software.
Figure 8. Validation of POLRMT, TFAM, and TFB2M purification. (A) SDS-PAGE analysis of purified POLRMT, TFAM, and TFB2M. (B) Validation of POLRMT activity on a nucleic acid scaffold. The size of a weakly radiolabeled RNA was increased by POLRMT-catalyzed incorporation of [a-32P]NTP to produce a 15-nt RNA. (C) TFB2M activity was validated in a promoter (LSP)-initiated transcription assay. The reaction was stopped at 19 nt with the incorporation of 3'-dCTP. The two lanes are the same sample loaded in increasing amounts.
To evaluate the expression of the proteins, a western blot can be run using an anti-His antibody. POLRMT can be identified this way after the purification as it still retains its His-tag. For TFAM and TFB2M, which will lack their His-tag after purification, performing the TFAM functional assay described above and performing an in vitro transcription assay will confirm the activity of the proteins.
Validation of protocol
The protocol was used to purify the POLRMT, TFAM, and TFB2M used for an in vitro transcription initiation assay in Figure 5D of our paper (Reardon and Mishanina, 2022).
POLRMT activity was validated by performing an in vitro transcription assay (Figure 8B) on a nucleic acid scaffold designed by Schwinghammer et al., 2013 (Supplementary Figure 1 in the paper). This assay indicated that POLRMT was able to add the incoming nucleotide to the starting RNA.
TFB2M activity was validated by performing an in vitro transcription initiation assay (Figure 8C) following the protocol by Ramachandran et al., 2017 (Figure 1 in the paper). The template contained the light strand promoter (LSP) sequence of mtDNA, which requires TFB2M and TFAM for transcription to occur.
General notes and troubleshooting
This protocol describes how to lyse cells with a sonicator, but a French press could be used instead. Similarly, a peristaltic pump could be used to generate buffer gradients instead of an FPLC.
For POLRMT expression: the goal is to prevent oversaturation of the cells. The lower the temperature, the longer the colonies can be left growing overnight, so any temperature between 25 °C and 37 °C will work. If colonies are oversaturated, the growth will be extremely slow, most likely because the leaky expression of POLRMT is toxic to E. coli cells.
Glycerol stocks could also be prepared at this time by storing 1 mL of the inoculum in 25% glycerol at -80 °C. We usually vortex 500 μL of inoculum with 500 μL of 50% sterile glycerol before freezing.
Lower temperatures will slow down translation and promote proper folding of the protein. To cool down the flasks, we leave them shaking on the incubator while they cool or put the flasks in large buckets of ice. The OD600 can be checked again but inducing in the range of 0.6–1.0 works fine.
We have tried to change the method to get more concentrated POLRMT in fewer fractions, but POLRMT ended up precipitating out very quickly, so it is not recommended.
We concentrate the HisTrap elution in the Amicon filter down to 25 mL (step 10) because 50 mL is a convenient volume that we can load onto our Bio-Rad NGC FPLC via a sample pump. If your system can handle a larger loading volume, you can add an equal volume of the No Salt Heparin buffer and skip the concentration step.
Because the HisTrap Buffer B has 300 mM NaCl, we need to bring the salt concentration down to the same as in the Heparin Buffer A to have POLRMT bind to the Heparin column. The Heparin column does not greatly increase purity but removes endogenous bound nucleic acids from the protein.
Previous attempts at buffer exchange resulted in increased POLRMT precipitation and are thus not recommended. We usually concentrate POLRMT to 6 or 7 μM before we start losing protein to precipitation.
The goal of the dialysis is to remove imidazole from the buffer, which inhibits TEV protease activity. It is also important to keep the salt concentration above 300 mM KCl to prevent TEV protease from crashing out. There is no EDTA in the TEV buffer because EDTA inhibits TEV activity.
Optional for downstream transcription experiments where salt concentration could be an issue: we can estimate the salt concentration of the purified TFB2M by the % Buffer B line at the time of TFB2M elution on the FPLC trace and calculate based on the known salt concentrations of Buffers A and B:
% of buffer B: _____
% of buffer A: 100% - % buffer B = ____
Volume of that fraction collected: ____mL
Volume of buffer B: Vol of fraction * % buffer B = _____mL
Volume of buffer A: Vol of fraction * % buffer A = _____mL
Moles of B = Vol Buffer B (1.5 mol)/(1,000 mL)=x mol KCl
Moles of A = Vol Buffer A (0.200 mol)/(1,00 0mL)= x mol KCl
Salt concentration of fraction = (Moles A+Moles B)/(Volume of fraction collected (mL)) × (1,000 mL)/(1 L) = x M salt concentration
Acknowledgments
We greatly appreciate Dr. Smita Patel (Rutgers University) and her lab members for providing the expression plasmids for POLRMT and TFAM and the protocols used in this work. We would like to thank Dr. Miguel Garcia-Diaz (Stony Brook University) for providing the expression plasmid for TFB2M. This work was supported by UCSD institutional support, Yinan Wang Memorial Chancellor’s Endowed Junior Faculty Fellowship, and R35GM142785 to T.V.M., R35GM118086 to S.S.P.; NIH Molecular Biophysics Training Grant (T32 GM008326) to A.H.H. and J.H.M-C.; NIH Multi-Scale Analysis of Biological Structure and Function Training Grant (T32 EB009380) to S.D.R. This protocol was adapted from and validated in previous works (Ramachandran et al., 2017; Reardon and Mishanina, 2022).
Competing interests
The authors declare no competing interests.
References
Bird, J. G., Basu, U., Kuster, D., Ramachandran, A., Grudzien-Nogalska, E., Towheed, A., Wallace, D. C., Kiledjian, M., Temiakov, D., Patel, S. S., et al. (2018). Highly efficient 5' capping of mitochondrial RNA with NAD+ and NADH by yeast and human mitochondrial RNA polymerase. eLife 7: e42179.
Hillen, H. S., Morozov, Y. I., Sarfallah, A., Temiakov, D. and Cramer, P. (2017). Structural Basis of Mitochondrial Transcription Initiation. Cell 171(5): 1072–1081.e10.
Kapust, R. B., Tözsér, J., Fox, J. D., Anderson, D., Cherry, S., Copeland, T. D. and Waugh, D. S. (2001). Tobacco etch virus protease: mechanism of autolysis and rational design of stable mutants with wild-type catalytic proficiency. Protein Eng. Des. Sel. 14(12): 993–1000.
Oláhová, M., Peter, B., Szilagyi, Z., Diaz-Maldonado, H., Singh, M., Sommerville, E. W., Blakely, E. L., Collier, J. J., Hoberg, E., Stránecký, V., et al. (2021). POLRMT mutations impair mitochondrial transcription causing neurological disease. Nat. Commun. 12(1): e1038/s41467-021-21279-0.
Ramachandran, A., Basu, U., Sultana, S., Nandakumar, D. and Patel, S. S. (2017). Human mitochondrial transcription factors TFAM and TFB2M work synergistically in promoter melting during transcription initiation. Nucleic Acids Res. 45(2): 861–874.
Raran-Kurussi, S., Cherry, S., Zhang, D., Waugh, D. S. (2017). Removal of Affinity Tags with TEV Protease. In: Burgess-Brown, N. (Ed.) Heterologous Gene Expression in E. coli (pp. 221–230). Methods in Molecular Biology. Humana Press, New York, NY.
Reardon, S. D. and Mishanina, T. V. (2022). Phosphorylation and acetylation of mitochondrial transcription factor A promote transcription processivity without compromising initiation or DNA compaction. J. Biol. Chem. 298(4): 101815.
Schwinghammer, K., Cheung, A. C. M., Morozov, Y. I., Agaronyan, K., Temiakov, D. and Cramer, P. (2013). Structure of human mitochondrial RNA polymerase elongation complex. Nat. Struct. Mol. Biol. 20(11): 1298–1303.
Sultana, S., Solotchi, M., Ramachandran, A. and Patel, S. S. (2017). Transcriptional fidelities of human mitochondrial POLRMT, yeast mitochondrial Rpo41, and phage T7 single-subunit RNA polymerases. J. Biol. Chem. 292(44): 18145–18160.
Tan, B. G., Mutti, C. D., Shi, Y., Xie, X., Zhu, X., Silva-Pinheiro, P., Menger, K. E., Díaz-Maldonado, H., Wei, W., Nicholls, T. J., et al. (2022). The human mitochondrial genome contains a second light strand promoter. Mol. Cell 82(19): 3646–3660.e9.
Yakubovskaya, E., Guja, K. E., Eng, E. T., Choi, W. S., Mejia, E., Beglov, D., Lukin, M., Kozakov, D. and Garcia-Diaz, M. (2014). Organization of the human mitochondrial transcription initiation complex. Nucleic Acids Res. 42(6): 4100–4112.
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Phylogenetic Inference of Homologous/Orthologous Genes among Distantly Related Plants
ZX Zilong Xu
WS Wenyan Sun
Ziqiang Zhu
BZ Bojian Zhong
ZZ Zhenhua Zhang
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4893 Views: 852
Reviewed by: Xin QiaoYao XiaoYe Xu
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Original Research Article:
The authors used this protocol in Communications Biology Apr 2023
Abstract
The recent surge in plant genomic and transcriptomic data has laid a foundation for reconstructing evolutionary scenarios and inferring potential functions of key genes related to plants’ development and stress responses. The classical scheme for identifying homologous genes is sequence similarity–based searching, under the crucial assumption that homologous sequences are more similar to each other than they are to any other non-homologous sequences. Advances in plant phylogenomics and computational algorithms have enabled us to systemically identify homologs/orthologs and reconstruct their evolutionary histories among distantly related lineages. Here, we present a comprehensive pipeline for homologous sequences identification, phylogenetic relationship inference, and potential functional profiling of genes in plants.
Key features
• Identification of orthologs using large-scale genomic and transcriptomic data.
• This protocol is generalized for analyzing the evolution of plant genes.
Keywords: Homolog Ortholog Similarity search Phylogenetic inference Functional profiling
Background
Evolution of plant genes is inextricably coupled with various evolutionary events, including endosymbiotic events, whole-genome duplication/triplication (WGD/T), gene loss, and horizontal gene transfer (Zhang et al., 2022). Archaeplastida, including green plants (Viridiplantae), glaucophytes (Glaucophyta), and red algae (Rhodophyta), originate anciently and most of them have experienced multiple WGD/T events, resulting in dramatic changes in copy numbers and complicated evolutionary trajectories of their homologous genes (Qiao et al., 2019). Homologs, orthologs, and paralogs are important concepts for the evolutionary classification of genes, being prevalent in recent comparative genomic studies. Homologs are genes sharing a common origin; orthologs and paralogs are two types of homologous genes, which separately evolved via speciation and gene duplication (Thornton and DeSalle, 2000; Koonin, 2005). Homologous genes generally have a relatively higher degree of sequence similarity than non-homologous genes. Sequence similarity–based searching and phylogenetic analyses are useful tools for identifying homologous sequences of genes and reconstructing their evolutionary routes.
Although the definition of homology/orthology has nothing to do with biological functions, there are major functional connotations (Koonin, 2005). Homologous/orthologous genes among different plants typically perform similar or equivalent functions, which is theoretically plausible and empirically supported. Thus, for a newly identified gene in non-model plants, identifying its homologs/orthologs in model plants or crops that have well-documented functional annotations is very useful to assign its possible functions. Phylogenetic analyses can reconstruct the evolutionary trajectories of homologs/orthologs among various species, which can facilitate the understanding of the molecular mechanisms underpinning its biological functions. Here, taking the acetyltransferase like protein HOOKLESS1 (HLS1) as an example (Lehman et al., 1996; Li et al., 2004), we provide a detailed procedure for homologs/orthologs identification using large-scale genomic and transcriptomic data of distantly related plants. This protocol includes generalized steps and parameters for evolutionary analyses of plant genes, and some of these steps and parameters can be customized based on the genes of interest.
Equipment
Server with a 64-bit Linux-based operating system (Ubuntu 18.04.6 LTS): 512 GB RAM and Intel Xeon (R) Gold 6238 CPU
Desktop with a Windows 10 operating system: Intel Core i5-8300H CPU and 8 GB RAM
Software and datasets
Software and databases used in this protocol are as follows:
Miniconda3-py39_4.12.0-Linux-x86_64 (https://mirrors.tuna.tsinghua.edu.cn/anaconda/miniconda/Miniconda3-py39_4.12.0-Linux-x86_64.sh)
TBtools v1.120 (Chen et al., 2020)
Diamond v2.1.7.161 (Buchfink et al., 2015)
MAFFT v7.453 (Katoh and Standley, 2013)
trimAL v1.4.rev15 (Capella-Gutiérrez et al., 2009)
IQ-TREE v2.2.2.6 (Minh et al., 2020)
InterProScan 5.63-95.0 (Jones et al., 2014)
1KP dataset (One Thousand Plant Transcriptomes Initiative, 2019)
MEME 5.5.3 (Bailey and Elkan, 1994)
iTOL (Interactive Tree Of Life) (Letunic and Bork, 2021)
Jalview v2.11.2.0 (Waterhouse et al., 2009)
Procedure
We show a detailed procedure for homologs/orthologs identification with large-scale genomic data (Figure 1).
Figure 1. Pipeline for homologs/orthologs identification with large-scale genomes and transcriptomes
Software installation
Miniconda3-py39_4.12.0-Linux-x86_64
Miniconda3 is a package manager for downloading and installing bioinformatics software. It can be downloaded and installed in the server by the following commands:
wget https://mirrors.tuna.tsinghua.edu.cn/anaconda/miniconda/Miniconda3-py39_4.12.0-Linux-x86_64.sh
bash Miniconda3-py39_4.12.0-Linux-x86_64.sh
Diamond v2.1.7.161
Diamond is a fast protein aligner for protein sequences that can be downloaded and installed using the following commands:
wget https://github.com/bbuchfink/diamond/releases/download/v2.1.7/diamond-linux64.tar.gz
tar -zxvf diamond-linux64.tar.gz
Add the path of Diamond to the environment variable.
MAFFT v7.453
MAFFT is a package for sequence alignment that can be installed by conda:
conda install mafft
trimAL v1.4.rev15
TrimAL is a tool for automated alignment trimming.
wget https://github.com/inab/trimal/archive/refs/tags/v1.4.1.tar.gzhttps://github.com/inab/trimal/archive/refs/tags/v1.4.1.tar.gz
tar -zxvf trimal-1.4.1.tar.gz
cd ./trimal-1.4.1/source/
make
Add the current directory to the environment variable after compilation.
IQ-TREE v2.2.2.6
Q-TREE 2 is a widely used tool for maximum-likelihood phylogeny inference.
wget https://github.com/iqtree/iqtree2/releases/download/v2.2.2.6/iqtree-2.2.2.6-Linux.tar.gz
tar -zxvf iqtree-2.2.2.6-Linux.tar.gz
Add the ./iqtree-2.2.2.6-Linux/bin to the environment variable.
InterProScan 5.63-95.0
InterProScan is a protein function annotation software. It can be download and installed following the instruction: InterProScan documentation-interproscan-docs documentation (https://interproscan-docs.readthedocs.io/en/latest/).
TBtools v1.120
TBtools is an integrated tool for bioinformatic analysis and may be downloaded from https://github.com/CJ-Chen/TBtools/releases/download/1.123/TBtools_windows-x64_1_123.exe.
Jalview v2.11.2.0
Jalview a free cross-platform program for multiple sequence alignment editing, visualization, and analysis. This software may be downloaded from https://www.jalview.org/.
Genome and transcriptome download and processing
Plant genomes download.
A total of 39 streptophytes (land plants and charophytes), 54 chlorophytes, 9 rhodophytes, and 1 glaucophyte were selected, covering all main clades of Archaeplastida (Table S1). The protein sequences or coding sequences (CDS) and GFF annotation files were downloaded. The used transcriptomes data of algae are available at the 1KP website (https://db.cngb.org/onekp/).
Removing redundant transcripts and short genes.
Based on the GFF annotation files of each genome, the redundant transcripts and short genes are removed by TBtools: (a) the longest transcript of each gene is retained to remove redundancy resulting from alternative splicing variations (detailed steps are briefly displayed in Video 1); (b) protein sequences length of genes is calculated by TBtools, and sequences shorter than 50 amino acids are manually filtered.
Video 1. Removing the redundant transcripts by TBtools
Identifying orthologous genes within large-scale genomic/transcriptomic data
Identifying candidate homologs from genomes.
In order to identify candidate homologs of Arabidopsis thaliana HLS1 (AtHLS1), we conducted similarity searches with relative stringency threshold (E value < 1 × 10-5) against the protein sequences of plant genomes. For genomic data, the relative stringency threshold (E value < 1 × 10-5) is used, and the reasons include: (1) gene annotation of genomic data is based on multiple evidences (such as gene database and transcriptomic data), which ensured the completeness and quality of gene sequences; (2) the genomic data in this study is mainly derived from the Phytozome (https://phytozome-next.jgi.doe.gov/) and has high qualities. We provide brief descriptions of files used in the command line in Table S2.
diamond makedb --in all_genome_sequences.fa -d all_genome_sequences
diamond blastp --db all_genome_sequences.dmnd --query AtHLS1.fa --out genome_out.result --outfmt 6 --sensitive -e 1e-5 --block-size 1.0 --index-chunks 1
Identifying candidate homologs from transcriptomes.
We used a relatively relaxed threshold to search candidate homologs from plant transcriptomes (E value < 1 × 10-2). The reason includes the intrinsic incompleteness of transcriptomes resulting from alternative splicing and premature termination. The detailed steps are briefly displayed in Video 2.
Video 2. BLASTp for AtHLS1 at the 1KP website
Filtering sequences with blast-hits.
Candidate homologs from both genomes and transcriptomes are integrated into a single fasta file (ID_sequences.fa), and InterProScan database is used to filter homologous sequences without a conserved functional domain: N-acetyltransferase (PF00583). Consolidate the homologous sequences with “N-acetyltransferase” domain into a new fasta file (PF00583_ID_sequences.fa) for further phylogenetic analyses.
Interproscan.sh -i ID_sequences.fa -f tsv -appl Pfam -o interproscan_ID_sequences.txt
Orthologs inference with phylogenetic analyses
Alignment and trimming of homologous sequences.
Homologous sequences are aligned by MAFFT using the following command:
mafft PF00583_ID_sequences.fa > mafft_out.fa
Note: An alignment of up to ~200 sequences × ~2,000 sites is suitable for an accurate option (L-INS-i), and an alignment of <~30,000 sequences is suitable for fast option (FFT-NS-2).
TrimAl is a widely used tool for automated alignment trimming in large-scale phylogenetic analyses. Before trimming, we perform manual inspection for sequence alignment using Jalview and exclude one sequence (in red rectangle) for its error alignment (Figure 2). We use a relatively relaxed threshold (‘-gt=0.13’: retaining columns that have at least 13% gap-free sites for keeping as much informative sites of conserved domains as possible) for trimming multiple sequence alignment (MSA). The trimmed MSA is further visually inspected to (1) filter the obvious ambiguously aligned regions and (2) retain the regions of functional domains, ensuring a greater proportion of reliably phylogenetically informative sites.
trimal -in mafft_out.fa -out mafftout_0.13.fas -gt 0.13
Figure 2. Multiple sequence alignment of AtHLS1 and its homologs. We empirically focused on the alignment region of conserved domains and found an obvious sequence that is poorly aligned (no amino acid residue was aligned).
Constructing a phylogenetic tree of homologous proteins.
The maximum likelihood phylogenetic tree of homologous proteins is inferred using IQ-TREE 2, and the best-fitting model is determined by ModelFinder. Branch supports are evaluated by the ultrafast bootstrap (UFBoot) approach and SH approximate likelihood ratio test (SH-aLRT test) with 1,000 replicates.
iqtree2 -s mafftout_0.13.fas -st AA -m MF
iqtree2 -s mafftout_0.13.fas -m Q.plant+I+R6 -alrt 1000 -bb 1000 –bnni -pre 0.13_result
Confirmation of orthologous sequences using function domains/motifs.
The conserved functional domains/motifs are analyzed by MEME suite. The parameter “number of motifs expected to be found” is set to ten (a common setting for this parameter) and other parameters are as default. Corresponding functional domains and motifs of each orthologous sequence are mapped to the phylogenetic tree using iTOL (Video 3). Two conserved residues (L327 and E346 in AtHLS1) are checked and highlighted in the MSA. Both the conserved functional domains/motifs and residues are used for the confirmation of orthologous sequence (Figure 3).
Video 3. Mapping the domain/motifs to the phylogenetic tree
Figure 3. Phylogenetic tree, two conserved residues, and conserved motifs of plant HLS1 homologs
Acknowledgments
This work is supported by the Natural Science Fund for Colleges and Universities in Jiangsu Province of China (21KJB180015). This protocol was adapted from our recent work (Wang et al., 2023). We thank the editor and anonymous reviewers for their helpful suggestions.
Competing interests
The authors declare that there are no conflicts of interest or competing interests.
References
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Chen, C., Chen, H., Zhang, Y., Thomas, H. R., Frank, M. H., He, Y. and Xia, R. (2020). TBtools: An Integrative Toolkit Developed for Interactive Analyses of Big Biological Data. Mol. Plant 13(8): 1194–1202.
Jones, P., Binns, D., Chang, H. Y., Fraser, M., Li, W., McAnulla, C., McWilliam, H., Maslen, J., Mitchell, A., Nuka, G., et al. (2014). InterProScan 5: genome-scale protein function classification. Bioinformatics 30(9): 1236–1240.
Katoh, K. and Standley, D. M. (2013). MAFFT Multiple Sequence Alignment Software Version 7: Improvements in Performance and Usability. Mol. Biol. Evol. 30(4): 772–780.
Koonin, E. V. (2005). Orthologs, Paralogs, and Evolutionary Genomics. Annu. Rev. Genet. 39(1): 309–338.
Lehman, A., Black, R. and Ecker, J. R. (1996). HOOKLESS1, an Ethylene Response Gene, Is Required for Differential Cell Elongation in the Arabidopsis Hypocotyl. Cell 85(2): 183–194.
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Minh, B. Q., Schmidt, H. A., Chernomor, O., Schrempf, D., Woodhams, M. D., von Haeseler, A. and Lanfear, R. (2020). IQ-TREE 2: New Models and Efficient Methods for Phylogenetic Inference in the Genomic Era. Mol. Biol. Evol. 37(5): 1530–1534.
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Qiao, X., Li, Q., Yin, H., Qi, K., Li, L., Wang, R., Zhang, S. and Paterson, A. H. (2019). Gene duplication and evolution in recurring polyploidization–diploidization cycles in plants. Genome Biol. 20(1): e1186/s13059-019-1650-2.
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Supplementary information
The following supporting information can be downloaded here:
Table S1. Sources of the genomic data used in this protocol
Table S2. Brief description of files used in the command line
Article Information
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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4,894 | https://bio-protocol.org/en/bpdetail?id=4894&type=0 | # Bio-Protocol Content
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Peer-reviewed
Purification of Human Cytoplasmic Actins From Saccharomyces cerevisiae
BH Brian K. Haarer
DA David C. Amberg
JH Jessica L. Henty-Ridilla
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4894 Views: 522
Reviewed by: William Jennings ValentineAlexandros C Kokotos Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal of Cell Science May 2023
Abstract
Eukaryotic cells rely on actin to support cellular structure, motility, transport, and a wide variety of other cytoplasmic functions and nuclear activities. Humans and other mammals express six closely related isoforms of actin, four of which are found primarily in skeletal, cardiac, and smooth muscle tissues. The final two isoforms, β and γ, are found in non-muscle cells. Due to the ease of purification, many biochemical studies surveying the functions of actin and its regulators have been carried out with protein purified from skeletal muscle. However, it has become increasingly clear that some activities are isoform specific, necessitating more accessible sources of non-muscle actin isoforms. Recent innovations permit the purification of non-muscle actins from human cell culture and heterologous systems, such as insect cell culture and the yeast Pichia pastoris. However, these systems generate mixtures of actin types or require additional steps to remove purification-related tags. We have developed strains of Saccharomyces cerevisiae (budding yeast) that express single untagged isoforms of either human non-muscle actin (β or γ) as their sole actin, allowing the purification of individual homogeneous actin isoforms by conventional purification techniques.
Key features
• Easy growth of yeast as a source of human cytoplasmic actin isoforms.
• Uses well-established actin purification methods.
• The tag-free system requires no post-purification processing.
Graphical overview
Isolating human cytoplasmic actins from yeast
Keywords: Actin purification Human actin Cytoplasmic actin β-actin γ-actin
Background
Actin and its many regulators are critical to a wide variety of eukaryotic cellular functions, ranging from determination of cell shape and motility to critical nuclear functions that include gene regulation and DNA repair (Belin et al., 2015; Schaks et al., 2019; Kadzik et al., 2020). In humans, the importance of actin regulation is further implicated in cancer metastasis, cardiomyopathies, neurological dysfunctions, and many other debilitating disorders (Rivière et al., 2012; Rubenstein and Wen, 2014; Bamburg and Bernstein, 2016; Hyrskyluoto and Vartiainen, 2020; Izdebska et al., 2020). However, mechanistic advances linking actin to these maladies have been confounded by the presence of six closely related yet distinct actin isoforms. Four actin isoforms are found predominantly in skeletal, cardiac, and smooth muscle tissues (α1, α2, α-cardiac, and γ2), and two are found primarily in non-muscle cells (β and γ1, commonly referred to as cytoplasmic actins). Further, much of what we know about the biochemical nature of critical actin regulators has been deciphered using the readily abundant and easily obtained 1 actin from animal muscle sources (e.g., rabbit, chicken, or cow). While muscle actin has been invaluable for the study of actin organization, regulation, and dynamics, it is becoming increasingly clear that some actin regulators display isoform specificity (De La Cruz, 2005; Perrin and Ervasti, 2010; Antoku et al., 2019). Thus, reliable sources of pure cytoplasmic actin isoforms are required to effectively explore these differing functions. However, unlike the muscle actin isoform α1, these isoforms are difficult to obtain in suitable concentrations and purity for widespread adoption and use.
Initially developed methods obtained mixtures of cytoplasmic actins from human cells (Schafer et al., 1998); similar mixtures of β and γ1 are available from commercial sources (Cytoskeleton, Inc., Denver, CO). Newer systems involving cleavable fusion proteins expressed in human cells or in the yeast Pichia pastoris improved purity to individual isoforms and extended expression to disease-relevant mutations in actin isoforms (Hatano et al., 2018; Hatano et al., 2020; Ceron et al., 2022). These purification systems have the further benefit of extending isoform analysis to important actin post-translational modifications, such as His73 methylation, and N-terminal processing. We produced a different, relatively inexpensive system that yields Human β or γ1 actin from the budding yeast Saccharomyces cerevisiae. Here, we combine engineered yeast strains with the classic purification of actin via DNase I affinity and elution with the denaturant formamide (Zechel et al., 1980; Kilmartin and Adams 1984; Kron et al., 1992; Goode 2002). This provides a relatively simple technique that does not require additional column chromatography. These and related sources of human cytoplasmic actins (Hatano et al., 2018; Hatano et al., 2020; Ceron et al., 2022) will play an important role in understanding the biochemical activities of non-muscle actin regulatory proteins, which may provide important new insights into actin-based disease development and progression.
Materials and reagents
Biological materials
BHY845 [MATa ura3∆0 leu2∆0 his3∆1 act1∆::NatR (pBH839 = 2μ LEU2 GPDpr-HsACTB)] S. cerevisiae strain that expresses only human β actin (Haarer et al., 2023); available with material transfer agreement (MTA) due to pending patent application (see conflict of interest statement).
BHY848 [MATα ura3∆0 leu2∆0 his3∆1 mfa1∆:: MFA1pr-SpHIS5 act1∆::NatR (pBH869 = 2μ LEU2 GPDpr-HsACTG1)] S. cerevisiae strain that expresses only human γ actin (Haarer et al., 2023); available on request, as above.
Reagents
Bacto peptone (BD-Gibco, catalog number: 211677)
Bacto yeast extract (BD-Gibco, catalog number: 212750)
Bacto agar (BD-Difco, catalog number: 214010)
Glucose (Sigma, catalog number: G-7021; Mallinckrodt, catalog number: 4908)
Glacial acetic acid (Fisher, catalog number: A38-212)
Hydrochloric acid (HCl) (Fisher, catalog number: A144S-212)
Tris base (Fisher, BP152-5) or Trizma base (Sigma, catalog number: T6066)
Sodium acetate (J.T. Baker, catalog number: 3470-01; Fisher, catalog number: BP333-500)
Sodium chloride (NaCl) (Sigma, catalog number: S7653)
Sodium bicarbonate (NaHCO3) (Sigma, catalog number: S5761)
Formamide, deionized (Sigma, catalog number: F9037)
Ammonium chloride (NH4Cl) (Sigma, catalog number: A9434)
Potassium chloride (KCl) (J.T. Baker, catalog number: 3040-01)
Calcium chloride dihydrate (CaCl2·2H2O) (Fisher, catalog number: BP510-500)
CNBr-activated Sepharose 4B (Cytiva, catalog number: 17-0430-01)
Deoxyribonuclease I (DNase I) (Worthington, catalog number: LS002007)
Bio-Rad protein assay dye reagent (Bio-Rad, catalog number: 5000006)
Calbiochem protease inhibitor cocktail IV (Millipore-Sigma, catalog number: 539136)
Phenylmethylsulfonyl fluoride (PMSF) (Sigma, catalog number: P7626)
Magnesium chloride, 6-hydrate (MgCl2·6H2O) (J.T. Baker, catalog number: 4003-01)
Ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′-tetraacetic acid (EGTA) (Sigma, catalog number: E4378)
Adenosine 5′-triphosphate, disodium trihydrate (ATP) (GoldBio, catalog number: A-081-25)
Dithiothreitol (DTT) (Bio-Rad, catalog number: 161-0611; GoldBio, catalog number: DTT25)
Potassium hydroxide (KOH) (Fisher, catalog number: P250-500)
Sodium hydroxide (NaOH) (Fisher, catalog number: BP359-500)
Sodium azide (Fisher, catalog number: BP922I-500)
Solutions
40% (w/v) glucose
1 mM HCl
1 M Tris pH 8.0
1 M Tris pH 7.5
3 M KCl
1 M MgCl2
0.5 M CaCl2
5 M NaCl
2% sodium azide
YPD medium (1 L), liquid (see Recipes)
YPD medium (1 L), plates (see Recipes)
40% glucose (1 L) (see Recipes)
Coupling buffer (250 mL) (see Recipes)
Low pH wash buffer (100 mL) (see Recipes)
High pH wash buffer (100 mL) (see Recipes)
0.5 M sodium acetate, pH 4 (500 mL) (see Recipes)
1 M tris, pH 7.5 or pH 8.0 (1 L) (see Recipes)
DNase I storage buffer (100 mL) (see Recipes)
G-buffer (50 mL) (see Recipes)
Lysis buffer (50 mL) (see Recipes)
Actin wash buffer 1 (50 mL) (see Recipes)
Actin wash buffer 2 (50 mL) (see Recipes)
Actin elution buffer (20 mL) (see Recipes)
20× F-buffer (1 mL) (see Recipes)
100 mM PMSF (100 mL) (see Recipes)
0.5 M ATP (20 mL) (see Recipes)
1 M DTT (50 mL) (see Recipes)
0.5 M EGTA (100 mL) (see Recipes)
Recipes
YPD medium (1 L), liquid
Reagent Final concentration Quantity
Bacto peptone 2% (w/v) 20 g
Bacto yeast extract 1% (w/v) 10 g
H2O 900 mL
Glucose (dextrose) 4% (w/v) 100 mL of 40% stock (see Note*, below)
*Note: Autoclave the peptone and yeast extract solution separately from the 40% glucose solution. This limits media caramelization and reaction with media components. Thus, to complete the YPD, add the sterile peptone-yeast solution to the sterile glucose aseptically (i.e., 900 mL of peptone-yeast solution and 100 mL of 40% glucose). Separate media components can be stored for several weeks at room temperature or at 4 °C for longer periods.
YPD medium (1 L), plates
Reagent Final concentration Quantity
Bacto peptone 2% (w/v) 20 g
Bacto yeast extract 1% (w/v) 10 g
Bacto agar 1.8% (w/v) 18 g (see note*)
H2O 900 mL
Glucose (dextrose) 4% (w/v) 100 mL of 40% stock (see Note*, below)
*Note: For plates, add agar to peptone and yeast extract solution, add magnetic stir bar, and autoclave. After autoclaving, add sterile 40% glucose solution and place on stir plate with slow stirring to avoid bubbles; let cool to ~55 °C. Carefully pour media plus agar solution into (35–40) sterile Petri dishes (10 cm) and let sit on bench for 1–2 days at room temperature; plates can be stored upside down in sealed Petri dish bags for several weeks at room temperature or at 4 °C for longer periods.
40% glucose (1 L)
Filter sterilize or autoclave
Reagent Quantity
Glucose (dextrose) 400 g
H2O to 1 L
Coupling buffer (250 mL)
Reagent Final concentration Stock solution Quantity
Sodium bicarbonate 0.1 M 2.1 g
Sodium chloride 0.5 M 5 M 25 mL
Calcium chloride 0.1 mM 0.5 M 50 μL
H2O Final volume to 250 mL
Low pH wash buffer (100 mL)
Reagent Final concentration Stock solution Quantity
Sodium acetate, pH 4 0.1 M 0.5 M, pH 4 20 mL
Sodium chloride 0.5 M 5 M 10 mL
H2O 70 mL
High pH wash buffer (100 mL)
Reagent Final concentration Stock solution Quantity
Tris, pH 8 0.1 M 1 M, pH 8 10 mL
Sodium chloride 0.5 M 5 M 10 mL
H2O 80 mL
0.5 M sodium acetate, pH 4 (500 mL)
Reagent Quantity
Sodium acetate 20.5 g
Glacial acetic acid to pH 4.0
H2O to 500 mL
1 M tris, pH 7.5 or pH 8.0 (1 L)
Reagent Quantity
Tris base 121.1 g
HCl to pH 7.5 or 8.0
H2O to 1 L
DNase I storage buffer (100 mL)
Reagent Final concentration Stock solution Quantity
Tris, pH 7.5 50 mM 1 M 5 mL
Sodium chloride 0.5 M 5 M 10 mL
Sodium azide 0.02% (w/v) 2% (w/v) 1 mL
H2O 84 mL
G-buffer (50 mL)
Reagent Final concentration Stock solution Quantity
Tris, pH 7.5 10 mM 1 M 0.5 mL
Calcium chloride 0.2 mM 0.5 M 20 μL
ATP 0.5 mM 0.5 M 50 μL (see Note*, below)
DTT 0.2 mM 1 M 10 μL*
H2O 49.4 mL*
*Note: Add ATP and DTT solutions the day of use; store G-buffer with ATP and DTT on ice or at 4 °C.
Lysis buffer (G-buffer and protease inhibitors; 50 mL)
Reagent Final concentration Stock solution Quantity
Tris, pH 7.5 10 mM 1 M 0.5 mL
Calcium chloride 0.2 mM 0.5 M 20 μL
ATP 0.5 mM 0.5 M 50 μL
DTT 0.2 mM 1 M 10 μL
PMSF 0.1 mM 0.1 M 50 μL
Calbiochem protease inhibitor cocktail IV 0.2% (v/v) 100 μL
H2O 49.27 mL
Actin wash buffer 1 (G-buffer and 10% formamide; 50 mL)
Reagent Final concentration Stock solution Quantity
Tris, pH 7.5 10 mM 1 M 0.5 mL
Calcium chloride 0.2 mM 0.5 M 20 μL
ATP 0.5 mM 0.5 M 50 μL
DTT 0.2 mM 1 M 10 μL
PMSF 0.1 mM 0.1 M 50 μL
Formamide 10% (v/v) 5 mL
H2O 44.37 mL
Actin wash buffer 2 (G-buffer and 0.2 M NH4Cl; 50 mL)
Reagent Final concentration Stock solution Quantity
Tris, pH 7.5 10 mM 1 M 0.5 mL
Calcium chloride 0.2 mM 0.5 M 20 μL
ATP 0.5 mM 0.5 M 50 μL
DTT 0.2 mM 1 M 10 μL
Ammonium chloride 0.2 M 0.5 M 20 mL
H2O 29.42 mL
Actin elution buffer (G-buffer and 50% formamide; 20 mL)
Reagent Final concentration Stock solution Quantity
Tris, pH 7.5 10 mM 1 M 0.2 mL
Calcium chloride 0.2 mM 0.5 M 8 μL
ATP 0.5 mM 0.5 M 20 μL
DTT 0.2 mM 1 M 4 μL
Formamide 50% (v/v) 10 mL
H2O 9.77 mL
20× F-buffer (1 mL)
10 mM tris pH 7.5, 500 mM KCl, 80 mM MgCl2, 20 mM EGTA, 10 mM ATP
Reagent Final concentration Stock solution Quantity
Tris, pH 7.5 10 mM 1 M 10 μL
Potassium chloride 0.5 M 3 M 167 μL
Magnesium chloride 80 mM 1 M 80 μL
EGTA 20 mM 0.5 M 40 μL
ATP 10 mM 0.5 M 20 μL
H2O 683 μL
100 mM PMSF (100 mL)
Store at -20 °C
Reagent Quantity
PMSF 1.74 g
100% ethanol to 100 mL
0.5 M ATP (20 mL)
Store filter-sterilized frozen aliquots at -20 °C
Reagent Quantity
ATP 6.05 g
KOH to pH ~ 7.2
H2O to 20 mL
1 M DTT (50 mL)
Store filter-sterilized frozen aliquots at -20 °C
Reagent Quantity
DTT 7.71 g
H2O to 50 mL
0.5 M EGTA (100 mL)
Reagent Quantity
EGTA
NaOH
19.0 g
to pH 8
H2O to 100 mL
Laboratory supplies
50 mL conical tubes (Thermo Fisher Scientific, Falcon, catalog number: 1495949A)
15 mL conical tubes (Thermo Fisher Scientific, Falcon, catalog number: 1495949B)
Pierce 5 mL disposable columns, including polyethylene discs, stoppers, and end caps (Thermo/Pierce, catalog number: 29922)
Funnels for Pierce columns (Thermo/Pierce, catalog number: 29923)
Petri dishes (Kord-Valmark, catalog number: 2900)
2 L Erlenmeyer flasks
Syringe filters, 0.45 μm (Genesee Scientific, catalog number: 25-242)
Slide-A-Lyzer dialysis cassettes, 7,000 or 10,000 MWCO (Thermo Fisher Scientific, catalog numbers: 66370 and 66380, respectively)
Dialysis tubing, 12,000–14,000 MWCO, 25 mm width (Fisher Scientific, catalog number: 21-152-18)
Protein concentrators: Vivaspin-6, 5,000 MWCO (Sartorius, catalog number: VS0611); Amicon Ultra-4, 10,000 MWCO (Millipore-Sigma, catalog number: UFC801024)
Equipment
Magnetic stir plate (optional; for making YPD plates)
Shaking incubator capable of holding multiple 2 L flasks at 30 °C
Spectrophotometer for cell density measurement (Bio-Rad, model: SmartSpec 3000)
Rocker or rotator, preferably at 4 °C (e.g., in cold room or chromatography cabinet)
Centrifuges:
Refrigerated tabletop centrifuge (Beckman Coulter, model: Allegra X-15R) for preparing resin
Superspeed centrifuge (Thermo Scientific/Sorvall, model: Lynx 4000) and rotor (Thermo Scientific, model: F10-4x1000 LEX) for harvesting 1–2 L of yeast
Ultracentrifuge (Beckman Coulter, model: Optima LE-80K) and rotor (Beckman Coulter, model: 70 Ti) for clarifying yeast lysates
Tabletop ultracentrifuge (Beckman Coulter, model: Optima Max-XP) and rotors (Beckman Coulter, models: TLA100, TLA100.2, TLA100.3) for clarifying actin solutions
French press and pressure cell with 1-inch diameter piston (American Instrument Company/SIM-Aminco)
Protein gel apparatus and power supply (Bio-Rad, models: Mini-Protean Tetra, PowerPac Basic Power Supply)
Optional: spectrophotometer for determining protein concentration (Thermo Scientific, model: NanoDrop 2000)
Procedure
Cell growth
Streak strains BHY845 and BHY848 (expressing β- and γ1-actin, respectively) from frozen glycerol stocks onto YPD plates (4% glucose) and incubate at 30 °C until colonies form (4–6 d; these strains are particularly slow growing, with doubling times ~4–6 times longer than wild-type laboratory strains).
Inoculate flasks containing 50 mL of YPD (4% glucose) with several colonies for each strain; grow on a shaker overnight at 30 °C.
Use 25–40 mL of the cultures from step A2 to inoculate 1 L of YPD cultures; incubate on a shaker at 30 °C.
Grow strains to OD595 ~1.5–2, which may take ~1.5–2 days.
Harvest yeast cells by centrifugation, e.g., in 250 mL bottles using a Beckman JA12 rotor at 7,000 rpm (7,900× g) for 7 min.
Decant culture medium and resuspend cells (e.g., by vortexing) in 10 mM Tris, pH 7.5, 0.2 mM CaCl2; transfer to 50 mL conical tubes. Pellets typically fill 6–7 mL of space in the tube.
Spin at 3,500× g for 10 min at 4 °C, decant, and then freeze pellets at -80 °C.
Coupling DNase I to Sepharose 4B (DNase I-resin preparation)
Swell 3 g of CNBr-activated Sepharose 4B in ice-cold 1 mM HCl in a 50 mL conical tube; place on a rocker or rotator to disperse persistent clumps.
Collect activated beads via centrifugation at ~2,000× g for 2 min; wash twice with 1 mM HCl and bring volume to 50 mL with each wash.
Resuspend 100 mg of DNase I in coupling buffer at 5 mg/mL final concentration; save a small aliquot (~100 μL) of DNase I suspension for later calculation of coupling efficiency and add the remainder to the resin. Incubate coupling reaction overnight at 4 °C on a rocker or rotator.
Allow the resin to settle on ice or collect via centrifugation at ~2,000× g for 2 min; remove and save the supernatant for calculation of coupling efficiency.
Add 25 mL of cold 0.1 M Tris pH 8.0 to block unreacted sites on beads. Rock at 4 °C for 2 h.
Collect resin via centrifugation at ~2,000× g for 2 min. Wash resin, alternating between 25 mL of each of the following solutions thrice:
Low pH wash buffer.
High pH wash buffer.
Wash resin twice with 25 mL of DNase I storage buffer; resuspend in DNase I storage buffer and store at 4 °C.
Determine coupling efficiency by comparing DNase I concentration of the pre- (step B3) and post-coupling (step B7) solutions; this can be done with standard protein determination reagents [e.g., Bio-Rad protein assay reagent, Bradford reagent, Lowry reagent, or direct A280 measurement (A280 of 1 = 0.5 mg/mL DNase I)]. Coupling efficiency should be ~ 95%–98%.
Column packing: 2 × 3 mL columns
The DNase I-Sepharose resin will be packed into two Pierce polypropylene columns, assembled from two 5 mL polypropylene columns, two filter discs, two column end caps, and two column top funnels.
Using an upside-down 1 mL pipette tip, push one filter disc to the bottom of each column and pass 5 mL of ddH2O through each.
Attach the funnel/reservoir to the top of the column: use parafilm to seal potential leak points between the funnel neck and the column (Figure 1A and 1B).
Figure 1. Diagram of disposable column assembly. (A) Schematic and (B) photo depicting column apparatus. See Protocol section D for details.
Cap the columns, add 3 mL of water, and note the top water level on the column using a marker. Uncap the column and remove the water via gravity flow.
Gradually add the resin to the columns, allowing it to slowly settle via gravity as the column flows. Add enough resin to reach the 3 mL mark.
Rinse columns and reservoirs with DNase I storage buffer to settle remaining resin via gravity flow.
Wash the column with 3–5 column volumes (CV) of DNase I storage buffer; then, cap the column (e.g., by covering with parafilm) and store at 4 °C until use (Section D).
Columns are generally reusable for several (3–5) purifications. Following each purification, the column should be washed with at least five CV of G-buffer, then washed (3–5 CV) and stored in DNase I storage buffer.
Actin purification [modified from Goode (2002) and Aggeli et al. (2014)]
Thaw cell pellet obtained from 1 L media in ~15 mL of lysis buffer.
Lyse the cells by passing through French press 2× at press setting of 1,000 psi (16,000 actual psi in a pressure cell with 1-inch diameter piston).
Spin lysate at ~17,000–20,000× g for 30 min at 4 °C (e.g., in a Beckman JA20 rotor at 12 krpm).
Spin the supernatant of the previous step in a Beckman Ti70 rotor at 50,000 rpm (256,600× g) for 50 min at 4 °C.
Wash each 3 mL of DNase I-Sepharose column with 15 mL (five CV) of G-buffer supplemented with 0.2 M NH4Cl followed by 15 mL of G-buffer supplemented with 0.1 mM PMSF.
Filter the clarified supernatant through 0.45 μm syringe filters (Figure 2A, Lanes 1 and 2).
Figure 2. Coomassie stained SDS-PAGE gels of β- and γ-actin purification steps. (A–B) Two gels showing the purification steps of β- and γ-actin. Relative amounts post 50% formamide elution, as follows: Lanes 7 and 8: 1/900 of total eluates; Lanes 9 and 10: 1/100 of post-dialysis concentrated samples; Lanes 11 and 12: 1/10 of G-actin pellets (after suspension in gel loading buffer); Lanes 13 and 14: 1/75 of KCl-treated F-actin supernatants; Lanes 17 and 18: 1/10 of G-actin pellets (after suspension in gel loading buffer); Lanes 19 and 20: 1/200 of final actin preparations. Abbreviations: MW, molecular weight markers; HSS, high speed supernatant; Sup, supernatant; F-actin, filamentous actin; G-actin, globular (monomeric) actin.
Load supernatants on DNase I columns and allow to drip through (Figure 2A, Lanes 3 and 4).
Note: The “depleted” flowthrough can be collected and analyzed by western blotting to determine the degree of binding and whether actin has saturated the DNase I resin.
Wash each column with:
15 mL (i.e., five CV) of actin wash buffer 1 (Figure 2A, Lanes 5 and 6).
15 mL of actin wash buffer 2.
15 mL of G-buffer.
Elute actin from each column with actin elution buffer (formamide disrupts the actin–DNase I interaction, likely denaturing one or both proteins); for 3 mL of resin, add 2 mL (<1 CV) of actin elution buffer to the column and discard the resulting flowthrough, then add another 4 mL and collect the flowthrough, dripping into a tube containing 2 mL of G-buffer (this helps to immediately reduce the formamide concentration and overall actin denaturation) (Figure 2A, Lanes 7 and 8).
Dialyze overnight against 2 L of G-buffer at 4 °C in Slide-A-Lyzer cassettes (10,000 MWCO) or dialysis tubing (12,000–14,000 MWCO); this step reduces the formamide concentration to ~0.1% and introduces fresh G-buffer into the actin solution.
Concentrate the dialyzed actin solution to ~ 0.5–1.5 mL using centrifugation-based protein concentrators (Figure 2A, Lanes 9 and 10); we have successfully used Vivaspin 5000 MWCO and Amicon Ultra 10,000 MWCO centrifugal concentrators for this step.
Distribute to 1 mL ultracentrifuge tubes and clarify actin at 213,000× g (70,000 rpm in a Beckman TLA100.2 rotor) for 30 min at 4 °C. This step removes aggregated actin (Figure 1B, Lanes 11 and 12).
Note: Larger volumes can be accommodated here and at subsequent steps using the higher capacity TLA100.3 rotor and associated tubes.
Distribute a maximum of 760 μL of the G-actin supernatant per tube to new 1 mL ultracentrifuge tubes.
Add 20× F-buffer to 1× final concentration per 1 mL ultracentrifuge tube (i.e., 40 μL of F-buffer to the 760 μL of G-actin already in the tube from step D13). Incubate tubes at room temperature for 20 min to polymerize F-actin.
Add KCl to 600 mM final (i.e., 200 μL of 3 M KCl stock to 800 μL mixture in step D14). Incubate at room temperature for 1 h. This step promotes the dissociation of potential host cofilin contaminants.
Collect F-actin via centrifugation at 213,000× g for 30 min at 20 °C. Remove and discard the supernatant (Figure 2B, lanes 13 and 14).
Rinse the F-actin pellets with ~200 μL of G-buffer (Figure 2B, Lanes 15 and16) and then resuspend in ~500 μL of G-buffer. Depolymerize actin filaments on ice overnight or for at least 3 h. Mechanically sheering actin filaments with a P200 pipette or by dounce homogenization will further expedite depolymerization.
Remove depolymerization-incompetent actin and actin aggregates via centrifugation at 213,000× g for 30 min at 4 °C (Figure 2B, lanes 17 and 18).
Transfer supernatants (G-actin) to new 1 mL ultracentrifuge tubes. Determine concentration and yield (e.g., using Bio-Rad protein assay reagent and comparing to known quantities of BSA).
Run 200–300 ng of actin on a 10%–15% polyacrylamide gel to assess purity (Figure 2B, Lanes 19 and 20).
To further enhance the purity of actin, perform 2–3 additional polymerization/depolymerization “cycles” (concentration permitting). To cycle actin, add 20× F-buffer to 1× final concentration, incubate for 20 min at room temperature, then repeat steps D16–D19 as above. Running small aliquots of the supernatants and resuspended pellets on SDS-PAGE gels can be used to track the final purity and/or loss of actin with each polymerization cycle. The amount of G-buffer to add when repeating step D17 will depend on yield and desired final concentration (e.g., suspending 500 μg of actin in 500 μL of G-buffer results in a 24 μM actin solution).
Store actin monomers in one of two ways:
For long-term storage, distribute G-actin into small (20–50 μL) aliquots, flash freeze in liquid nitrogen, and store at -80 °C.
For short-term storage (1–2 weeks), add 20× F-buffer (i.e., 5 μL of 20× F-buffer per 95 μL of G-actin) and store polymerized F-actin at 4 °C.
When ready to use:
Thaw aliquot and clarify monomers via centrifugation at 213,000× g for 30 min at 4 °C to remove aggregates.
Spin F-actin at 213,000× g for 30 min at 20 °C, resuspend actin pellet in fresh G-buffer, and incubate on ice to depolymerize (see step D17).
Validation of protocol
This protocol has been used and validated in the following research article:
Haarer et al. (2023). Purification of human β- and γ-actin from budding yeast. Journal of Cell Science 136: jcs260540. Doi: 10.1242/jcs.260540.
General notes and troubleshooting
General notes
BHY848 is respiratory incompetent (petite). We recommend growing both strains (BHY848 and BHY845) with 4% glucose.
It is convenient to make β- and γ-actin preparations in parallel. We recommend having dedicated DNase I columns for each isoform to avoid cross-contamination.
Run each step of the purification (including column washes and spin supernatants and pellets) on polyacrylamide gels to assess effectiveness, actin loss, and relative purity (see Figure 2).
DNase I affinity columns lose effectiveness over time. We recommend using columns for 3–5 purifications.
Alternate yeast lysate preparation methods can be used in the absence of a French press with pressure cell, including microfluidizer, ball mill, or bead beater.
There are many methods for purifying actin. We see no reason why alternate methods would not be compatible with purification of β- or γ-actin from clarified yeast lysates.
Growing yeast in bioreactors or generally scaling cultures > 1 L permits additional polymerization/depolymerization cycles and gel filtration methodologies.
Troubleshooting
Problem 1: Low protein yield due to cell lysis (step D2).
Possible cause: Cell lysis is incomplete.
Solutions: After passing through French press (or comparable breaking procedure), observe cells under a microscope; if many cells remain unbroken, pass through breaking procedure one or two more times.
Problem 2: DNase I binding capacity is low.
Possible causes: Low efficiency coupling of DNase I to new resin or saturation/breakdown of a reused column.
Solution: Replace column.
Problem 3: Formide-related actin denaturation.
Possible cause: The ratio of Formide to actin was too high.
Solution: Limit exposure to formamide by dilution (leaving space for solution in step D9). Keep samples on ice whenever possible.
Problem 4: Final concentration of actin is too low (step D19 onward).
Possible cause: Loss of actin during cycling.
Solutions: Ensure that F-actin pellets have had sufficient time and agitation to convert to G-actin. To maximize recovery of low-yield actin, resuspend F-actin pellets in G-buffer to at least 1 μg/μL prior to converting to F-actin.
Acknowledgments
Components of the graphical overview were created with and exported from BioRender.com under a paid subscription purchased by SUNY Upstate. This work was supported by R35GM133485 to JLH-R, and by support from SUNY Upstate Medical University and the Research Foundation of SUNY to DCA. We are grateful to Thomas J. Black for his acerbic wit, energy, and peaches. This protocol was adapted from our previously published study (Haarer et al., 2023).
Competing interests
The strains described in this protocol and their use for purifying human cytoplasmic actin isoforms are the subject of a provisional patent application by the authors and SUNY Research Foundation (App No: 63/399,088).
References
Aggeli, D., Kish-Trier, E., Lin, M. C., Haarer, B., Cingolani, G., Cooper, J. A., Wilkens, S. and Amberg, D. C. (2014). Coordination of the filament stabilizing versus destabilizing activities of cofilin through its secondary binding site on actin. Cytoskeleton 71(6): 361–379.
Antoku, S., Wu, W., Joseph, L. C., Morrow, J. P., Worman, H. J. and Gundersen, G. G. (2019). ERK1/2 Phosphorylation of FHOD Connects Signaling and Nuclear Positioning Alternations in Cardiac Laminopathy. Dev. Cell 51(5): 602–616.e12.
Bamburg, J. R. and Bernstein, B. W. (2016). Actin dynamics and cofilin-actin rods in alzheimer disease.Cytoskeleton 73(9): 477–497.
Belin, B. J., Lee, T. and Mullins, R. D. (2015). Correction: DNA damage induces nuclear actin filament assembly by Formin-2 and Spire-1/2 that promotes efficient DNA repair. eLife 4: e11935.
Ceron, R. H., Carman, P. J., Rebowski, G., Boczkowska, M., Heuckeroth, R. O. and Dominguez, R. (2022). A solution to the long-standing problem of actin expression and purification. Proc. Natl. Acad. Sci. U. S. A. 119(41): e2209150119.
De La Cruz, E. M. (2005). Cofilin Binding to Muscle and Non-muscle Actin Filaments: Isoform-dependent Cooperative Interactions. J. Mol. Biol. 346(2): 557–564.
Goode, B. L. (2002). Purification of yeast actin and actin-associated proteins. Methods Enzymol. 351: 433–441.
Haarer, B. K., Pimm, M. L., de Jong, E. P., Amberg, D. C. and Henty-Ridilla, J. L. (2023). Purification of human β- and γ-actin from budding yeast. J. Cell Sci. 136(9): e260540.
Hatano, T., Alioto, S., Roscioli, E., Palani, S., Clarke, S. T., Kamnev, A., Hernandez-Fernaud, J. R., Sivashanmugam, L., Chapa-Y-Lazo, B., Jones, A. M. E., et al. (2018). Rapid production of pure recombinant actin isoforms in Pichia pastoris. J. Cell Sci. 131(8): jcs213827.
Hatano, T., Sivashanmugam, L., Suchenko, A., Hussain, H. and Balasubramanian, M. K. (2020). Pick-ya actin: a method to purify actin isoforms with bespoke key post-translational modifications.J. Cell Sci. 133(2): jcs241406.
Hyrskyluoto, A. and Vartiainen, M. K. (2020). Regulation of nuclear actin dynamics in development and disease. Curr. Opin. Cell Biol. 64: 18–24.
Izdebska, M., Zielińska, W., Hałas-Wiśniewska, M. and Grzanka, A. (2020). Involvement of Actin and Actin-Binding Proteins in Carcinogenesis. Cells 9(10): 2245.
Kadzik, R. S., Homa, K. E. and Kovar, D. R. (2020). F-Actin Cytoskeleton Network Self-Organization Through Competition and Cooperation. Annu. Rev. Cell Dev. Biol. 36(1): 35–60.
Kilmartin, J. V. and Adams, A. E. (1984). Structural rearrangements of tubulin and actin during the cell cycle of the yeast Saccharomyces. J. Cell Biol. 98(3): 922–933.
Kron, S. J., Drubin, D. G., Botstein, D. and Spudich, J. A. (1992). Yeast actin filaments display ATP-dependent sliding movement over surfaces coated with rabbit muscle myosin. Proc. Natl. Acad. Sci. U. S. A. 89(10): 4466–4470.
Perrin, B. J. and Ervasti, J. M. (2010). The actin gene family: function follows isoform. Cytoskeleton 67(10): 630–634.
Rivière, J. B., van Bon, B. W. M., Hoischen, A., Kholmanskikh, S. S., O’Roak, B. J., Gilissen, C., Gijsen, S., Sullivan, C. T., Christian, S. L., Abdul-Rahman, O. A., et al. (2012). De novo mutations in the actin genes ACTB and ACTG1 cause Baraitser-Winter syndrome. Nat. Genet. 44(4): 440–444.
Rubenstein, P. A. and Wen, K. K. (2014). Insights into the effects of disease-causing mutations in human actins.Cytoskeleton 71(4): 211–229.
Schafer, D. A., Jennings, P. B. and Cooper, J. A. (1998). Rapid and efficient purification of actin from nonmuscle sources. Cell Motil. Cytoskeleton 39(2): 166–171.
Schaks, M., Giannone, G. and Rottner, K. (2019). Actin dynamics in cell migration. Essays Biochem. 63(5): 483–495.
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Peer-reviewed
H2 Production from Methyl Viologen–Dependent Hydrogenase Activity Monitored by Gas Chromatography
NK Nuttavut Kosem
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4895 Views: 562
Reviewed by: Chhuttan L MeenaKarem A Court Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Applied Catalysis A: General Feb 2023
Abstract
Bio-hydrogen production is an eco-friendly alternative to commercial H2 production, taking advantage of natural systems. Microbial hydrogenases play a main role in biological mechanisms, catalyzing proton reduction to molecular hydrogen (H2) formation under ambient conditions. Direct determination is an important approach to screen bacteria with active hydrogenase and accurately quantify the amount of H2 production. Here, we present a detailed protocol for determining hydrogenase activity based on H2 production using methyl viologen (MV2+) as an artificial reductant, directly monitored by gas chromatography. Recombinant Escherichia coli is used as a hydrogenase-enriched model in this study. Even so, this protocol can be applied to determine hydrogenase activity in all biological samples.
Key features
• This protocol is optimized for a wide variety of biological samples; both purified hydrogenase (in vitro) and intracellular hydrogenase (in vivo) systems.
• Direct, quantitative, and accurate method to detect the amount of H2 by gas chromatography with reproducibility.
• Requires only 2 h to complete and allows testing various conditions simultaneously.
• Kinetic plot of H2 production allows to analyze kinetic parameters and estimate the efficiency of hydrogenase from different organisms.
Graphical overview
Keywords: Hydrogenase Hydrogen Methyl viologen Gas chromatography Biocatalyst
Background
Although hydrogen (H2) has been considered as a clean energy carrier, most of today’s commercial H2 production is from fossil fuels, which emit greenhouse gases into the atmosphere. Finding a more sustainable alternative remains a challenge. It is well known that, for over a million years of evolution, microorganisms have acted as a natural H2 production plant through the action of enzymes, namely the hydrogenase family (Tard and Pickett, 2009). With efficient biochemical mechanisms, it is possible to develop an environmentally friendly process for H2 production at ambient conditions by taking advantage of hydrogenase-expressing microorganisms. Among the enzymes with different catalytic sites, [FeFe]-hydrogenase is more efficient than [NiFe]- and [Fe]-types in H2 production with NADH as a reductant in a natural mechanism, as shown in Figure S1 in the Supporting materials (Ogo et al., 2020; Xuan et al., 2023). To screen microorganisms carrying active hydrogenase in a laboratory scale, methyl viologen (MV2+), which is compatible with many enzymes (Orgill et al., 2015), can also be used as an artificial reductant in this reaction. According to the theory, a more negative redox potential of methyl viologen [E(MV2+/MV•+) = -0.446 V vs. normal hydrogen electrode (NHE)] provides a favorable potential scale for proton reduction [E(H+/H2) = -0.41 V vs. NHE] at physiological pH.
In this protocol, a reduced methyl viologen (MV•+) is formed in sodium dithionite (Na2S2O4) solution to serve as an artificial chemical reductant. With cell permeability, MV•+ can penetrate and transfer electrons to intracellular hydrogenase (Kosem et al., 2023), as shown in Figure 1.
Figure 1. Methyl viologen (MV2+) as an artificial reductant in hydrogenase activity assay. (A) MV2+ reduction to MV•+ formation in Na2S2O4 solution. (B) H2 production from the MV•+-dependent reaction of [FeFe]-hydrogenase.
The amount of H2 generated is directly monitored by gas chromatography. To avoid the detrimental effect of atmospheric oxygen and enhance biocatalytic activity, the reactions are carried out under anaerobic conditions at 37 °C. This method can be applied to determine H2 production capacity of various biological samples such as whole cells, crude cell extracts, or purified enzymes.
Materials and reagents
Biological materials
Hydrogenase-expressing Escherichia coli carrying hydA, hydE, hydF, and hydG genes (this recombinant bacterial strain was constructed in our laboratory according to our previous report in Kosem et al., 2023).
Note: The function of each protein expressed from specific genes in the processes of hydrogenase maturation was reported in many published articles, such as Broderick et al. (2014) and Lubitz et al. (2014).
Reagents
LB broth, Miller (Nacalai Tesque, catalog number: 20068-75)
Methyl viologen (Tokyo Chemical Industry, catalog number: D3685)
Sodium dithionite (Na2S2O4) (Sigma-Aldrich, catalog number: 71699-250G)
PierceTM BCA Protein Assay kit (Thermo Scientific, catalog number: 23227)
Sodium chloride (NaCl) (Wako Chemical, catalog number: 191-01665)
Tris (hydroxymethyl) aminomethane (Tris) (Nacalai Tesque, catalog number: 35434-21)
Hydrochloric acid (HCl) (Wako Chemical, catalog number: 080-0106)
Trichloroacetic acid (TCA) (Nacalai Tesque, catalog number: 34637-14)
3-(N-morpholino)propanesulfonic acid (MOPS) (Nacalai Tesque, catalog number: 23415-25)
Sodium hydroxide (NaOH) (Chameleon Reagent, catalog number: 000-75165)
Ampicillin sodium (Wako, catalog number: 014-23302)
Streptomycin sulfate (Wako, catalog number: 194-08512)
Glucose (Nacalai Tesque, catalog number: 16805-35)
Ferric ammonium citrate (Nacalai Tesque, catalog number: 19425-12)
L-cysteine (Nacalai Tesque, catalog number: 10309-12)
Sodium fumarate (TCI, catalog number: F0070)
Isopropyl-β-D-thiogalactopyranoside (IPTG) (FujiFilm, catalog number: 094-05144)
Solutions
LB (Luria-Bertani) broth (see Recipes)
1 M MOPS-NaOH (see Recipes)
100 mg/mL Ampicillin sodium (see Recipes)
40 mg/mL Streptomycin sulfate (see Recipes)
50% (w/v) Glucose (see Recipes)
250 mg/mL Ferric ammonium citrate (see Recipes)
1 M Sodium fumarate (see Recipes)
1 M IPTG (see Recipes)
0.9% (w/v) NaCl (see Recipes)
50 mM MV2+/Na2S2O4 solution (see Recipes)
1 M Tris-HCl pH 7 (see Recipes)
Recipes
LB broth
Note: Dissolve the medium and all reagents in a 500 mL Erlenmeyer flask and then autoclave at 121 °C for 15 min before use.
Reagent Final concentration Quantity
LB broth 2.5 % (w/v) 5 g
1 M MOPS-NaOH (pH 7.4) (Recipe 2) 100 mM 20 mL
Water n/a 180 mL
Total n/a 200 mL
The following (see Recipes 3–8) are added after autoclaving:
100 mg/mL Ampicillin sodium 100 μg/mL 0.2 mL
40 mg/mL Streptomycin sulfate 40 μg/mL 0.2 mL
50% (w/v) Glucose 0.5% (w/v) 2 mL
250 mg/mL Ferric ammonium citrate 250 μg/mL 0.2 mL
L-cysteine 2 mM 50 mg
1 M Sodium fumarate 20 mM 4 mL
1 M IPTG 1 mM 0.2 mL
1 M MOPS-NaOH pH 7.4
Note: Dissolve in a 100 mL beaker on a magnetic stirrer.
Reagent Final concentration Quantity
MOPS 1 M 20.9 g
Water n/a 80 mL
NaOH (8 M) n/a Slowly add until pH 7.4
Total n/a Make up final volume to 100 mL
100 mg/mL Ampicillin sodium
Note: Dissolve in a 5 mL microcentrifuge tube and mix thoroughly with gentle shaking by hand. Aliquot the stock solution in 1 mL microcentrifuge tubes (500 μL per tube) and store at -20 °C until use.
Reagent Final concentration Quantity
Ampicillin sodium 100 mg/mL 0.5 g
Sterilized water n/a 5 mL
Total n/a 5 mL
40 mg/mL Streptomycin sulfate
Note: Dissolve in a 5 mL microcentrifuge tube and mix thoroughly with gentle shaking by hand. Aliquot the stock solution in 1 mL microcentrifuge tubes (500 μL per tube) and store at -20 °C until use.
Reagent Final concentration Quantity
Streptomycin sulfate 40 mg/mL 0.2 g
Sterilized water n/a 5 mL
Total n/a 5 mL
50% (w/v) Glucose
Note: Dissolve glucose powder with warm sterilized water in a 100 mL beaker on a magnetic stirrer. Once it has completely dissolved, bring the volume up to 100 mL total. Aliquot the stock solution in 15 mL tubes (10 mL per tube) and store at -20 °C until use.
Reagent Final concentration Quantity
Glucose 50% (w/v) 50 g
Sterilized water n/a 60 mL
Total n/a Make up final volume to 100 mL
250 mg/mL Ferric ammonium citrate
Note: Dissolve in a 5 mL microcentrifuge tube and mix thoroughly with gentle shaking by hand. Aliquot the stock solution in 1 mL microcentrifuge tubes (500 μL per tube) and store at -20 °C until use.
Reagent Final concentration Quantity
Ferric ammonium citrate 250 mg/mL 1.25 g
Sterilized water n/a 5 mL
Total n/a 5 mL
1 M Sodium fumarate
Note: Dissolve in a 50 mL tube and mix thoroughly with gentle shaking by hand. Aliquot the stock solution in 15 mL tubes (10 mL per tube) and store at -20 °C until use.
Reagent Final concentration Quantity
Sodium fumarate 1 M 8.0 g
Sterilized water n/a 50 mL
Total n/a 50 mL
1 M IPTG
Note: Dissolve in a 5 mL microcentrifuge tube and mix thoroughly with gentle shaking by hand. Aliquot the stock solution in 1 mL microcentrifuge tubes (500 μL per tube) and store at -20 °C until use.
Reagent Final concentration Quantity
IPTG 1 M 1.2 g
Sterilized water n/a 5 mL
Total n/a 5 mL
0.9% (w/v) NaCl solution
Note: Dissolve in a 100 mL beaker on a magnetic stirrer.
Reagent Final concentration Quantity
NaCl 0.9% (w/v) 0.9 g
Water n/a 100 mL
Total n/a 100 mL
50 mM MV2+/Na2S2O4 solution
Note: Prepare fresh in an anaerobic sealed vial inside a glove box and mix thoroughly with gentle shaking by hand.
Reagent Final concentration Quantity
Methyl viologen 50 mM 0.0257 g
Na2S2O4 250 mM 0.0870 g
Water n/a 2 mL
Total n/a 2 mL
1 M Tris-HCl pH 7
Note: Dissolve in a 100 mL beaker on a magnetic stirrer. Adjust pH with 3 M HCl in a fume hood and strictly following the Kyushu University Guidelines for Safety in Education: Laboratory Activities. A safety data sheet from the chemical company as shown in Guideline 1 in the Supporting materials.
Reagent Final concentration Quantity
Tris 1 M 12.1 g
Water n/a 80 mL
HCl (3 M) n/a Slowly add until pH 7
Total n/a Make up final volume to 100 mL
Laboratory supplies
Glass vial No. 3, 10 mL size (ASONE, catalog number: 5-111-03)
Rubber stopper (ASONE, catalog number: 5-112-01)
Aluminum cap (ASONE, catalog number: 5-112-03)
1 mL Syringe (Terumo, catalog number: SS-01T)
Needle No. 27 G × 3/4" (Terumo, catalog number: NN-2719S)
10 μL pipette tip (Molecular BioProducts, catalog number: 3510-05)
200 μL pipette tip (Violamo, catalog number: 3-6504-12)
1,000 μL pipette tip (Violamo, catalog number: 3-6504-13)
1 mL microcentrifuge tube (Quality Scientific Plastics, catalog number: L-510-GRD-Q)
5 mL microcentrifuge tube (ASONE, model: AST0500)
96-well plate (ASONE, catalog number: 2-8085-02)
99.999% N2 gas (Air Liquide, catalog number: JAGA 13038)
Gloves (Showaglove, catalog number: 882.L.BLUE)
Paper towels
Disposable cuvettes (Violamo, model: UVC-Z8.5)
Equipment
2–20 μL autoclavable micropipette (Nichipet Ex II, Nichiryo, catalog number: J16Y01961)
20–200 μL autoclavable micropipette (Nichipet Ex II, Nichiryo, catalog number: J16Z00871)
100–1,000 μL autoclavable micropipette (Nichipet Ex II, Nichiryo, catalog number: J16X11751)
accu-jet® pro pipette controller (BRAND®, catalog number: Z671533)
High-speed micro centrifuge (Hitachi, model: CF16RN)
Personal centrifuge (Front Lab, model: FLD2012)
Gas chromatography (Shimadzu Corp., model: GC-8A)
Thermal conductivity detector (Shimadzu Corp., model: GC-8AIT)
Integrator C-R6A chromatopac (Shimadzu Corp., catalog number: 223-04500-38)
Molecular sieve 5A beads (GL Sciences Inc., catalog number: 1001-11503)
Stainless steel column, 2 m length × 3 mm diameter (Shimadzu Corp., catalog number: 201-48705-20)
Shaking water bath (Yamato Scientific, model: BW101)
Shaking incubator (EYELA, model: FYC-100)
Anaerobic glove box (MIWA, model: 1ADB-3)
Clean bench (Hitachi Appliances Inc., model: CCV-1300E)
Microplate reader (Corona Electric, model: SH-1000)
500 mL Erlenmeyer flask with baffle (IWAKI, model: CTE33)
250 mL centrifuge bottle (Nalgene, catalog number: B1033)
20 mm vial crimper (Chromatography Research Supplies, catalog number: 320990)
20 mm vial decapper pliers (Kebby, catalog number: D-20)
1 mL gas tight syringe (ITO Corporation, catalog number: MS-GAN100)
Weighing balance with 0.0001 g accuracy (Metter Toledo, model: XS204)
Vortex mixer (Scientific Industries, model: SI-0286)
pH meter (Horiba Scientific, model: 9625)
Water distillation apparatus (Advantec, model: RFD240NA)
Magnetic stirrer (Pasorina Stirrer, model: CT-MINI)
100 mL Beaker (AGC Iwaki, model: CTE33)
Fume hood (Oriental, model: TNV-STZ-1800HCS)
Software and datasets
Microsoft Excel
SF6 (version 5.6.0, Copyright© 2005 Corona Electric) for spectrophotometer
Procedure
Cultivation of hydrogenase-expressing bacteria
In this protocol, a recombinant E. coli encoding hydA, hydE, hydF, and hydG genes for hydrogenase expression was utilized as a H2-producing model constructed in our laboratory as reported in Honda et al. (2016) and Kosem et al. (2023).
Note: This protocol can be applied to determine the efficiency of other hydrogenase-producing bacteria, as reported in Benoit et al. (2020) and Kosem et al. (2024).
Aerobically pre-culture the recombinant E. coli in a 500 mL Erlenmeyer flask with 200 mL of LB broth supplemented with 100 μg/mL of ampicillin sodium, 40 μg/mL of streptomycin sulfate, 0.5% (w/v) of glucose, 250 μg/mL of ferric ammonium citrate, and 100 mM MOPS/NaOH pH 7.4. Incubate the pre-culture flask at 37 °C placed on a shaking incubator at 120 rpm until the cell density reaches an optical density of 0.4 at OD600, monitored by a spectrophotometer using a 1 mL cell suspension in a disposable cuvette. Then, transfer the 200 mL pre-cultured E. coli suspension into a 250 mL centrifuge bottle inside an anaerobic glove box supplemented with 2 mM L-cysteine (50 mg), 20 mM sodium fumarate (4 mL of 1 M stock solution), and 1 mM IPTG (0.2 mL of 1 M stock solution), and further incubate for 18 h in the glove box for [FeFe]-hydrogenase expression (Figure 2A).
Figure 2. Experimental process of MV2+-dependent hydrogenase activity
Cell harvesting and preparation
In the glove box, transfer 200 mL of cell suspension cultivated in LB medium from the Erlenmeyer flask to a centrifuge bottle with a silicone cap to preserve an anaerobic environment.
Centrifuge the bottle at 2,000× g for 10 min, discard the supernatant, and collect cell pellet.
Wash the cell pellet once with 5 mL of 0.9% NaCl solution and centrifuge at 2,000× g for 10 min.
To prepare cell suspension for experiments, resuspend the washed cell pellet in 5 mL of 0.9% NaCl solution (Figure 2A).
To standardize the protocol, the concentration of cells used in the reaction is based on total protein contents measured by PierceTM BCA Protein Assay kit in a 96-well plate.
Note: The protocol can be modified when different bacterial strains or biological samples are used.
Hydrogenase activity assay
Degas all chemical reagents with pure N2 for 5 min and place them in the anaerobic glove box before use.
Prepare the following reagents as shown in Table 1 and Figure 2B.
Table 1. Reaction mixture preparation of MV•+-hydrogenase activity assay
Reagents Applied volume Final concentration
1 M Tris-HCl pH 7 0.2 mL 100 mM
10 mg/mL protein of cell suspension or extract 0.2 mL 1 mg/mL
Sterile deionized water 1.4 mL
50 mM MV2+ in 250 mM Na2S2O4 (reduced MV•+ solution) 0.2 mL 5 mM
Total 2 mL
Notes:
The reaction mixture is prepared in a 10 mL glass vial and sealed with a rubber stopper and aluminum cap inside the glove box.
The protein concentration can be adjusted in different samples.
In case of a negative control, the same volume of sterile water is added instead of cell suspension or extracts.
The reduced MV•+ solution is prepared in a separate vial and sealed with a rubber stopper and aluminum cap inside the glove box.
Take the reaction vial and the MV•+ vial out of the glove box.
Purge the contaminated gases in each vial with pure N2 for 5 min.
Initiate the reaction by adding 0.2 mL of 50 mM MV2+/Na2S2O4 solution into the reaction vial using a 1 mL syringe with needle No. 27 G × 3/4" and further incubate in a shaking water bath at 100 rpm and 37 °C.
Note: The reaction mixture turns dark blue after adding MV2+/Na2S2O4 solution into the reaction vial.
Every 20 min after incubation, terminate the reaction vial by adding 0.1 mL of 100% TCA (concentration of the commercial stock solution) using a 1 mL syringe with needle No. 27 G × 3/4".
Sample 1 mL of H2 produced in a headspace of the reaction vial using a 1 mL gas tight syringe and vertically inject into a gas chromatograph for analysis. Here, a GC-8A gas chromatograph equipped with a thermal conductive detector and an integrator C-R6A chromatopac was used. The produced gas goes through molecular sieve 5A beads packed in a stainless steel column (2 m length × 3 mm diameter) with a carrier gas of argon. Operating parameters are shown in Table 2.
Table 2. Chromatographic operating parameters
Parameters Units
Pressure of carrier gas, Ar 100 kPa
Injection volume 1 mL
Detector temperature 50 °C
Injection temperature 50 °C
Column temperature 50 °C
Detector current 60 mA
Data analysis
Quantification of H2:
The accurate amount of H2 produced from the reaction is calculated from a calibration curve between H2 concentration vs. peak area obtained from the chromatogram of GC analysis (Figure 3).
Figure 3. Calibration plot of standard H2 vs. peak area by gas chromatography. According to the calibration curve, the amount of H2 can be calculated from the following equation: H2 (mol) = (Peak area – Intercept)/Slope.
Calculation of hydrogenase activity
(1)
In which the amount of H2 (μmol) in the headspace (Figure 4A) is measured at each time point and plotted to obtain the H2 production rate (Figure 4B).
Time (minutes or hours) is the incubation time of the reaction.
Note: Normally it takes 1–2 h for the incubation period. The amount of H2 production rate is presented in μmol per min.
Cprotein (mg/mL) is the final concentration of protein in the reaction mixture.
Vreaction (mL) is the total volume of the reaction mixture.
Figure 4. Methyl viologen (MV•+)-dependent hydrogenase activity assay. (A) Reaction vial containing the liquid phase of reaction mixture and the gas phase of H2 produced in a headspace. (B) H2 production rate is calculated from a kinetic plot between the amount of H2 in the y-axis vs. time in the x-axis. Hydrogenase activity is calculated from equation (1).
Validation of protocol
This protocol was validated in Kosem et al. (2023). Applied Catalysis A: General, DOI: 10.1016/j.apcata.2022.119019.
General notes and troubleshooting
To preserve the biological function of O2-sensitive enzymes, biological samples containing hydrogenase must be protected from air. Therefore, the whole process of cell preparation and experiments must be performed under anaerobic environment or in the glovebox.
Kinetic parameters of different hydrogenases from various biological sources can be analyzed by varying the concentrations of methyl viologen as a substrate according to the Michaelis-Menten equation: , where v is velocity, Vmax is the maximum velocity, Km is the Michaelis constant, and [S] is the concentration of the substrate (methyl viologen).
Acknowledgments
I would like to thank Prof. Tatsumi Ishihara for his supervision on my previous research, which originally validated this protocol in Kosem et al. (2023) that was funded by a Grant-in-Aid for Specially Promoted Research (No. 21K18213) from the Ministry of Education, Culture, Sports, Science and Technology (MEXT), Japan through the Japan Society for the Promotion of Science. I would also like to acknowledge the Department of Applied Chemistry, Faculty of Engineering, Kyushu University, and the International Institute for Carbon-Neutral Energy Research (I2CNER), Kyushu University for research facilities and instruments. I am thankful to Dr. Yuki Honda from whom I adapted this protocol.
Competing interests
The authors declare no competing interests.
References
Benoit, S. L., Maier, R. J., Sawers, R. G. and Greening, C. (2020). Molecular Hydrogen Metabolism: a Widespread Trait of Pathogenic Bacteria and Protists. Microbiol. Mol. Biol. Rev. 84(1): e00092-19.
Broderick, J. B., Byer, A. S., Duschene, K. S., Duffus, B. R., Betz, J. N., Shepard, E. M. and Peters, J. W. (2014). H-Cluster assembly during maturation of the [FeFe]-hydrogenase. J. Biol. Inorg. Chem. 19(6): 747–757.
Honda, Y., Hagiwara, H., Ida, S. and Ishihara, T. (2016). Application to Photocatalytic H2 Production of a Whole‐Cell Reaction by Recombinant Escherichia coli Cells Expressing [FeFe]‐Hydrogenase and Maturases Genes. Angew. Chem. Int. Ed. 55(28): 8045–8048.
Kosem, N., Watanabe, M., Song, J. T., Takagaki, A. and Ishihara, T. (2023). A comprehensive study on rational biocatalysts and individual components of photobiocatalytic H2 production systems. Appl. Catal. A: Gen. 651: 119019.
Kosem, N., Shen, X.-F., Ohsaki, Y., Watanabe, M., Song, J. T. and Ishihara, T. (2024). Photobiocatalytic conversion of solar energy to NH3 from N2 and H2O under ambient condition. Appl. Catal. B: Environ. 342: 123431.
Lubitz, W., Ogata, H., Rüdiger, O. and Reijerse, E. (2014). Hydrogenases. Chem. Rev. 114(8): 4081–4148.
Ogo, S., Kishima, T., Yatabe, T., Miyazawa, K., Yamasaki, R., Matsumoto, T., Ando, T., Kikkawa, M., Isegawa, M., Yoon, K. S., et al. (2020). [NiFe], [FeFe], and [Fe] hydrogenase models from isomers. Sci. Adv. 6(24): eaaz8181.
Orgill, J. J., Chen, C., Schirmer, C. R., Anderson, J. L. and Lewis, R. S. (2015). Prediction of methyl viologen redox states for biological applications. Biochem. Eng. J. 94: 15–21.
Tard, C. and Pickett, C. J. (2009). Structural and Functional Analogues of the Active Sites of the [Fe]-, [NiFe]-, and [FeFe]-Hydrogenases. Chem. Rev. 109(6): 2245–2274.
Xuan, J., He, L., Wen, W. and Feng, Y. (2023). Hydrogenase and Nitrogenase: Key Catalysts in Biohydrogen Production. Molecules 28(3): 1392.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Catalytic sites of different hydrogenases.
Guideline 1. Safety procedure in handling HCl.
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Correction Notice: Correlative Conventional and Super-resolution Photoactivated Localization Microscopy (PALM) Imaging to Characterize Chromatin Structure and Dynamics in Live Mammalian Cells
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Targeted Delivery of Chemogenetic Adeno-Associated Viral Vectors to Cortical Sulcus Regions in Macaque Monkeys by Handheld Injections
KO Kei Oyama *§
YN Yuji Nagai *§
TM Takafumi Minamimoto
(*contributed equally to this work, § Technical contact)
Published: Vol 13, Iss 23, Dec 5, 2023
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Jan 19, 2022
Original Research Article:
The authors used this protocol in Science Advances Jun 2021
Abstract
Recent advancements in chemogenetic tools, such as designer receptors exclusively activated by designer drugs (DREADDs), allow the simultaneous manipulation of activity over a specific, broad brain region in nonhuman primates. However, the introduction of DREADDs into large and complexly shaped cortical sulcus regions of macaque monkeys is technically demanding; previously reported methods are time consuming or do not allow the spatial range of expression to be controlled. In the present report, we describe the procedure for an adeno-associated viral vector (AAV2.1) delivery via handheld injections into the dorsolateral prefrontal cortex (Brodmann’s area 9/46) of macaque monkeys, with reference to pre-scanned anatomical magnetic resonance images. This procedure allows the precise delivery of DREADDs to a specific cortical region.
Key features
• This article describes the procedures for injecting viral vectors encoding functional proteins for chemogenetic manipulation into targeted cortical sulcus regions.
• The protocol requires magnetic resonance imaging for the accurate estimation of the injection sites prior to surgery.
• Viral vector solutions are injected using a handheld syringe under microscopic guidance.
• This protocol allows for the precise introduction of designer receptors exclusively activated by designer drugs (DREADDs) to large and complex cortical regions.
Keywords: Monkeys Chemogenetics Surgery Handheld injection Viral vector MRI
Background
Genetic approaches that enable the expression of desired functional proteins in order to manipulate the activity of specific brain regions or cell types have become a central method in systems neuroscience [1, 2]. In particular, chemogenetic tools such as designer receptors exclusively activated by designer drugs (DREADDs) have become a promising means to study the relationship between animal behavior and the neuronal activity of specific cell populations. DREADDs allow the manipulation of activity in specific brain regions in behaving animals following systemic administration of an agonist, without any requirements for special devices or implants [3]. This feature is particularly advantageous for manipulating the brain activity of nonhuman primates such as macaque monkeys, which have relatively large and complexly shaped brains.
The introduction of functional proteins to an intended brain region is typically achieved by the injection of viral vector solutions. To manipulate the neuronal activity of a specific region, these vectors need to be precisely delivered over the entire target region. However, if the target region is large and deep within the brain, such as for the cortical sulcus regions, it is technically demanding to inject viral vectors to cover the entire target region. For example, the introduction of DREADDs into a cortical region surrounding a sulcus [e.g., the dorsolateral prefrontal cortex (dlPFC) or Brodmann’s area 46] requires injection of vectors into a broad area along the banks of the principal sulcus (i.e., from the bottom to the surface; Figure 1). A conventional and typical stereotaxic approach using a manipulator ensures accurate localization of viral delivery. However, completing one injection with this procedure takes several tens of minutes, which makes the total surgical duration very long, with considerable surgical stress to the animal. An alternative approach termed convection-enhanced delivery enables widespread cortical viral delivery via a single injection, which considerably shortens the procedural time and minimizes damage [4]. However, its extent of spatial delivery is uncontrollable, which leads to concerns about off-target expression.
Figure 1. Illustration of the dorsolateral prefrontal cortex (dlPFC). Left, lateral view of the monkey brain. Right, coronal section around the dlPFC (colored area).
In the present report, we describe a procedure for viral vector injection into deep sulcus regions (such as the dlPFC) of macaque monkeys using a handheld injection technique. This procedure enables flexible injection of vectors within 1 min per injection track, each of which has several injection points. This procedure is based on a method described in previous reports [5, 6]. The injections cover 5–10 mm horizontally around the principal sulcus, as we have previously reported [7–9]. Our protocol uses a magnetic resonance (MR) image obtained before surgery to identify the position of the intended injection area, injection tracks and points, and angles for each track. The successful expression of the DREADDs was validated using conventional histological methods.
Materials and reagents
Biological materials
Macaque monkey (5.7 kg; age = 6.0 years old at the beginning of experiments; provided by the National Bio-Resource Project “Japanese Monkeys” of the Ministry of Education, Culture, Sports, Science and Technology, Japan)
Adeno-associated viral vector (e.g., AAV2.1-hSyn-hM4Di-IRES-AcGFP, provided by the Takada Lab, Kyoto University, Japan)
Reagents
Ketamine hydrochloride (Daiichi Sankyo Chemical Pharma, product name: Ketalar for intramuscular injection 500 mg)
Xylazine hydrochloride (Bayer, product name: Seraktar or Rompun)
Sodium cefmetazole (Alfresa Pharma, product name: Cefmetazon for intramuscular injection 0.5 g)
Ketoprofen (Kissei Pharmaceutical, product name: Capisten for intramuscular injection 50 mg)
Lidocaine 2% solution including epinephrine (Sandoz K.K., product name: Xylocaine injection 2% with epinephrine)
Lidocaine spray (Sandoz K.K., product name: Xylocaine pump spray 8%)
Sterilized saline (Otsuka Pharmaceutical, catalog number: 1326)
Povidone-iodine solution (Meiji Seika Pharma, product name: Povidone-iodine 10%)
Propofol (Nichi-Iko, product name: Propofol 1% intravenous injection)
Mannitol (Yoshindo, product name: Mannitol-s injection)
Atropine sulfate (Nipro, product name: Atropine sulfate hydrate 0.5 mg/mL)
Isoflurane [Mylan, product name: Isoflurane inhalation solution (Pfizer)]
Infusion solution (Terumo, catalog number: TP-AB05NR)
Bone wax (TOKYO M.I., catalog number: J901)
Sodium lactate Ringer’s solution (Terumo, product name: Solulact infusion)
Sterilized distilled water (Otsuka Pharmaceutical, catalog number: 1323)
Laboratory supplies
Endotracheal tube (Fuji Systems, catalog number: FR-26)
IV catheter (Terumo, catalog number: SR-FF2419)
25G needle (Terumo, catalog number: NN-2525R)
Surgical tape (Nichiban, catalog number: 21N)
Surgical cotton (Hakujuji, catalog number: 11371)
Disposable electrode for electrocardiogram (Nihon Kohden, catalog number: M-150)
1 mL syringe (Terumo, catalog number: SS-01T)
50 mL syringe (Terumo, catalog number: 5SS-50ESZ)
Valve syringe (Eastsidemed, catalog number: ES-17005)
Micro syringe (Hamilton, model: 1701RN directly connected to a 2 inch PT3 type 30G needle)
Depilatory cream (Center Shoji, product name: Fi-i-mo Epi DX Plus)
Scalpel
No. 11 (Akiyama Medical MFG, catalog number: FS11)
No. 21 (Akiyama Medical MFG, catalog number: FS21)
Suture with needle
Vicryl Plus 3-0 (Ethicon, catalog number: VCP460H)
Vicryl Plus 5-0 (Ethicon, catalog number: VCP303H)
Monosof 2-0 (COVIDIEN, catalog number: SN-628)
Infusion tube (Terumo, catalog number: TK-U200L)
Sterile gown (Nissho Sangyo, catalog number: 24503)
Sterile glove
Overglove (Mölnlycke Health Care, catalog number: 42175)
Underglove (Mölnlycke Health Care, catalog number: 41670)
Sterile drape
1,200 mm × 1,200 mm, no hole (Nissho Sangyo, catalog number: 23203)
900 mm × 900 mm, with hole and tape (Nissho Sangyo, catalog number: 23265)
Ioban antimicrobial incise drape (3M, catalog number: 6661EZ)
Sterile surgical marker (Muranaka Medical Instruments, catalog number: 2730PBX)
Sterile gauze (Hakujuji, catalog number: 14824)
Freer elevator (Roboz, catalog number: RS-8820)
Ruskin rongeur (Biomedical Research Instruments, catalog number: 46-1655)
Knapp scissors (Roboz, catalog number: RS-5965)
Drill bit
Diamond disc, Ø 6–9 mm disc (Biomachinery, custom made)
Twist drill, Ø 2 mm tip (Nakanishi Inc., catalog number: ES-TD-S20)
Equipment
MR imaging scanner (Bruker, model: 7 Tesla 400 mm/SS system)
X-ray computed tomography (CT) scanner (J. Morita, model: 3D Accuitomo170)
Operating microscope (Leica Microsystems GmbH, model: M220)
Patient monitor (Fukuda, model: Bio-Scope AM140)
Shadowless operating lamp (Yamada, model: CRV0404V)
Ventilator (Shin-Ei Industries, Inc., model: A.D.S. 2000)
Vaporizer (Shin-Ei Industries, Inc., model: I-200)
Stereotaxic device (David Kopf Instruments, model: 1530)
Electrolyzed water generator (Omco, model: NDX-70KMW)
Micromanipulator (David Kopf Instruments, model: 2166A)
Micro syringe pump (World Precision Instruments, model: UMP3T-1)
Circulating thermal water system (KIMURAMED, model: T-CARE)
Surgical drill control unit (Nakanishi Inc. model: Primado2)
Software and datasets
PMOD (PMOD Technologies Ltd. ver. 3.6 or above)
Procedure
Planning the surgery using MR and CT images
MR and CT scans (General note 1).
Anesthetize the monkey by intramuscular (i.m.) injection of ketamine (5–10 mg/kg) and xylazine (0.2–0.5 mg/kg) at 5 min after the i.m. injection of atropine sulfate (0.02–0.05 mg/kg) to reduce salivation, which is followed by continuous intravenous (i.v.) infusion of propofol (0.2–0.6 mg/kg/min) to achieve stable anesthesia during the scan.
Monitor the animal’s heart rate and peripheral oxygen saturation (SpO2) until the animal awakens.
Spray lidocaine into the trachea before securing the airway with an endotracheal tube.
Perform an MR imaging on the monkey under anesthesia.
Note: Any MR scan protocol is acceptable as long as the target location is clearly visible. As an example, the scanner, sequences, and parameters used in our institute are as follows: Bruker Biospec 40 cm bore 7T-MRI scanner; three-dimensional fast low angle shot gradient echo sequence; repetition time/echo time = 30/3.7 ms; flip angle = 10°; spatial resolution = 0.5 mm isotropic; field of view = 100 × 100 × 70 mm3; acquisition time = 6 min; fat suppression ON; band width = 347 Hz/Px; acceleration factor = 2.
Perform a CT scan of the head.
Note: We typically take a CT scan as a reference for aligning the MR images to the stereotaxic coordinates. The typical conditions are as follows: 90 kV, 5 mA, and 18.5 s rotation time for Ø 140 mm × 100 mm. The CT image should include the ear and orbit so that it can be virtually aligned to the stereotaxic coordinates. Any other methods that allow such alignment (e.g., MR imaging with a stereotaxic landmark) are also acceptable.
Estimation of injection sites.
Align an MR image to the stereotaxic coordinates using image analysis software (e.g., PMOD). In our typical procedure, we first align the CT image to the stereotaxic coordinates by matching the vertical level of the ear holes and the infraorbital margin (the lower margin of the eye socket), and then fuse the MR image to the stereotaxically aligned CT image (Figure 2).
Note: Any other software that allows the alignment of an MR image to stereotaxic coordinates is acceptable.
Figure 2. Alignment of the magnetic resonance (MR) image to the stereotaxic coordinates. A. Vertically aligned ear holes (white arrows) and the infraorbital margin (the lower margin of the eye socket) (yellow allows). B. Fused MR and computed tomography (CT) images on the horizontal (i), coronal (ii), and sagittal (iii) planes. Scale bar (white bar below the brain image): 10 mm.
Verify the location of the target area based on the stereotaxically aligned MR images. Critical: We typically check the depth of the sulcus from the surface, the distance from the interaural line, and the horizontal angle of the sulcus (which often varies along the anterior–posterior axis and between the left and right hemispheres). Based on the images, we then design the injection tracks and points. For example, in the case of Figure 3, the depth of the cortex (from the bottom to the surface) along the dorsal bank of the principal sulcus is approximately 8 mm, and the angle of the sulcus is 35°. For this case, we designed five injection points along the sulcus (at 1.5 mm intervals) to allow the vector solution to spread throughout the dlPFC (Figure 3, also see Figure 1).
Figure 3. Coordination of injection points on the magnetic resonance image. The red dots and yellow bar represent the intended injection points and injection track, respectively. Scale bar (white bar below the brain image): 10 mm.
Surgical procedures
Preparation for surgery.
Anesthetize the monkey with i.m. injection of ketamine (5–10 mg/kg) and xylazine (0.2–0.5 mg/kg) after the i.m. injection of atropine sulfate (0.02–0.05 mg/kg) to reduce salivation.
Note: The animal injected with an appropriate dose of ketamine and xylazine may be immobilized within 10 min after the injection, and the effects of ketamine and xylazine can continue for 30–40 min. Therefore, the next two steps should be done within that timeframe to maintain immobilization.
Move the animal to the preparation room.
Maintain immobilization with anesthesia using isoflurane (2%–3%) through a mask until the airway is secure.
Remove the hair around the head (first coarsely with hair clippers and then with depilatory cream).
Secure vascular access from the right foot vein using an IV catheter.
Spray lidocaine into the trachea before securing the airway with an endotracheal tube.
Start the anesthesia with isoflurane (1%–3%) through the secured airway and maintain it throughout the surgery.
Note: The surgery can be performed with the continuous use of a mask. We recommend the use of an endotracheal tube to secure the airway, which allows fine-tuning of anesthetic control by reducing the dead space.
Monitor the animal’s electrocardiogram, SpO2, end-tidal CO2, body temperature, and respiratory rate throughout the surgery.
Note: The surgery should be suspended immediately and appropriate measures taken if the following are noted: body temperature > 40 °C or < 35 °C, heart rate >150 beats/min or < 60 beats/min, SpO2 < 95%, and end-tidal CO2 > 50 mmH2O or < 20 mmH2O.
Maintain body temperature at 36–38 °C using a circulating thermal water system throughout and after surgery until the animal awakens.
Fix the monkey’s head to the stereotaxic device and check whether the head is leveled.
Note: The vertical level of the ear holes and the infraorbital margin should be matched with the alignment of the MR image aligned to stereotaxic coordinates.
After fixation, close the monkey’s eyes and cover them with surgical cotton for protection.
Connect the infusion tube to the vascular access and provide an i.v. injection of infusion solution (sodium lactate Ringer’s solution, approximately 10 mL/kg/h) during the surgery.
Administer prophylactic antibiotics (sodium cefmetazole, 25–50 mg/kg, i.m.).
Clean the monkey’s head by brushing with hypochlorite water (produced using the electrolyzed water generator). Dry the head and wipe with disinfectant (povidone-iodine) solution and sterile gauze.
Note: When cleaning with the sterile gauze, the operator should wear sterile gloves.
The following surgical procedures should be performed under sterile operating conditions.
From the beginning of surgery until before the craniotomy:
Put on a sterile gown and sterile gloves (undergloves and overgloves).
Note: Two people wearing sterile equipment are required for the handheld injections in the flow protocol—one to hold the syringe and one to push the plunger. Additionally, a third person is required to handle the viral vector.
Prepare sterile surgical instruments on sterile drapes.
Mark the incision line with the sterile surgical marker, which is typically approximately 10 mm from the tip of the forehead to the lambdoid suture at the midline of the head.
Cover the monkey with the Ioban antimicrobial incise drape and the sterile drape (900 mm × 900 mm, with a 90 mm hole), with the head exposed.
Begin the i.v. injection of hypotensive agent (i.e., 15%–20% mannitol, 4 mL/kg for 30 min).
Note: Glycerol is also acceptable (recommended dose: 8 mL/kg for 30 min, i.v. injection).
Subcutaneously inject the lidocaine 2% solution including epinephrine along the intended incision line to induce an analgesic effect and prevent bleeding.
Incise the skin and subcutaneous tissue with the scalpel (No. 21) and exfoliate the muscles with the elevator to expose the skull.
Spread and secure the incised skin and exfoliated muscles on both sides using suture thread (Vicryl Plus 3-0) to maintain the operative field.
Note: Cover the spread tissues (such as skin, subcutaneous tissue, and muscle) with damp gauze to keep them moist throughout the surgery. Keep the gauze damp by adding saline using the valve syringe.
Craniotomy, dura incision, and viral vector injections
Mark the contour of the intended cranial extraction site with the sterile surgical marker (Figure 4).
Figure 4. Contour of the intended cranial extraction site. Illustration of the intended cranial extraction site (red dashed line) on the monkey’s head from the lateral (left) and top (right) views. Red dots indicate the location of the suture holes to be made.
Craniotomy. Cut the edge of the intended area with the disc drill and extract the bone flap from the skull using the elevator.
Note: When targeting both sides, a wide bilateral craniotomy is acceptable. However, when cutting the midline, be very careful not to damage the superior sagittal sinus. Tilt the disc slightly from vertical so that the brain is not compressed when the bone is put back in place.
Smooth the edge of the opened skull (especially on the cerebral side) using the Ruskin rongeur.
Stop any bleeding on the incision surface of the skull using bone wax.
Make suture holes in the skull and bone flap with the drill to allow for the return of the bone flap into place (Figure 4, General note 2).
Note: The holes are drilled first to prevent bone fragments from entering under the dura mater during the drilling process.
Cut the dura with the scalpel (No. 11) and Knapp scissors so that the operator can see the landmarks of the brain surface, such as the arcuate and principal sulci for Brodmann’s area 46 (Figure 5).
Note: The incised dura mater should be moved aside (the dura flap folded back) for later suturing. After cutting the dura, particular care should be taken to keep the brain surface and dura mater moist with sterile saline.
Figure 5. Incision line of the dura mater. Illustration of the intended incision area of the dura mater (red dashed line). The dura is incised to visualize the rostral tip of the ascending limb of the arcuate sulcus (as) and the caudal tip of the principal sulcus (ps).
Prepare the viral vector solution.
Note: We typically use adeno-associated virus vectors expressing DREADDs under the control of general or specific promoters, with marker proteins or a tag sequence (e.g., AAV2.1-hSyn-hM4Di-IRES-AcGFP, as used in our previous studies [10–12]). For more details on the high-efficacy viral vector used in our recent work, see also [13].
Fill the injection syringe (10 μL Hamilton syringe with PT3 type 30 G needle) with the vector solution to be used.
Note: A PT2 needle or 25 μL micro syringe is also acceptable. Before filling the syringe with the vector solution, wash it first with ethanol and then with distilled water.
Perform the injection. One operator should manually hold the syringe and insert the needle to the intended depth under the guidance of an operating microscope. The other operator then pushes the plunger (Figure 6). Critical: The needle that we use is marked at 3 and 7 mm from the tip so that we can easily recognize the depth (Figure 7). For one hemisphere, perform nine penetrations surrounding the principal sulcus (Figure 8), with three to five injections at different depths (1 μL per injection) per penetration (see also Figure 3). It takes approximately 2 s to perform one injection. In total, 35–44 μL of vector solution is injected into one hemisphere.
Note: To avoid movement of the syringe during injection, the hands holding the syringe should be placed on the stereotaxic frame and the skull and supported by the other hand also placed on the stereotaxic frame.
Figure 6. Handheld injection. The needle is inserted into the intended area by one person (front). A second person (back) presses the plunger to expel approximately 1 µL of solution. The angle of the syringe is visually calibrated to be parallel to the manipulator, which is tilted to the angle of the sulcus.
Figure 7. Customized needle with marker for injection. Arrows indicate markers at 3 and 7 mm from the needle tip.
Figure 8. Injection area. Arrows indicate injection tracks along the principal sulcus.
Closing the head.
Suture the dura with the fusible sutures (Vicryl Plus 5-0) before checking for cerebrospinal fluid (CSF) leakage.
Note: Sutures should be evenly spaced at approximately 2 mm to prevent CSF leakage. If CSF leakage is noted, continue suturing until no leakage occurs.
Suture the bone flap back in place with the fusible sutures (Vicryl Plus 3-0) (General note 2).
Suture the fascia and subcutaneous tissue with the fusible sutures (Vicryl Plus 3-0) and then suture the skin with the nylon sutures (Monosof 2-0).
Administer analgesia (ketoprofen, 1–2 mg/kg, i.m.).
Upon awakening, return the monkey to its home cage.
Note: To prevent being harmed by other monkeys, it is preferable to keep the monkey alone until the sutures are removed.
Post-surgical procedures
Perioperative management
Administer prophylactic antibiotics by i.m. injections (sodium cefmetazole, 25–50 mg/kg/day) and analgesics (ketoprofen, 1–2 mg/kg/day) for seven days.
Remove the sutures when the skin has healed (after ≥ two weeks), after which experiments can be initiated.
Validation of protocol
The successful expression of DREADDs was confirmed by post-mortem histological inspection (Figure 9) in each study using our previously reported protocol [7–9]. We also confirmed DREADDs expression in vivo by conducting positron emission tomography (PET) scans with a radiolabeled DREADDs ligand at 45 days after the surgery [8, 9].
Figure 9. Histological section. 3,3’-diaminobenzidine-stained section showing immunoreactivity against a reporter protein (AcGFP).
We have also observed robust chemogenetic effects on behavior (i.e., working memory impairment) following the administration of a DREADD agonist (deschloroclozapine) [7–9].
General notes and troubleshooting
General notes
MRI and CT scans are performed on the same day. Following estimation of the injection sites based on MRI, the surgery is performed on a later day (typically ≥ one week later).
Insert an elevator or flat instrument between the skull and the dura mater to prevent injury of the dura mater and brain when making suture holes and when suturing to restore the bone flap.
Acknowledgments
This study was supported by the Ministry of Education, Culture, Sports, Science and Technology (MEXT)/ Japan Society for the Promotion of Science (JSPS) KAKENHI grant numbers JP21K07268, JP22H05521 (to KO), JP19K08138, JP23H02405 (to YN), and JP20H05955 (to TM); JST PRESTO grant number JPMJPR22S3 (to KO); and AMED grant number JP18dm0307007 (to TM). The two Japanese monkeys used in the original work (Oyama et al., 2021) were provided by the National Bio-Resource Project “Japanese Monkeys” of MEXT, Japan. We thank Jun Kamei, Ryuji Yamaguchi, Yuichi Matsuda, Yoshio Sugii, Takashi Okauchi, Rie Yoshida, Risa Suma, and Tomomi Kokufuta for their technical assistance. We also thank Mark A. G. Eldridge and Richard C. Saunders for providing us with tremendous guidance and advice on surgical techniques. This protocol was adapted from our previous works [7–9].
Competing interests
The authors declare no competing interests.
Ethical considerations
All experimental procedures involving animals were carried out in accordance with the Guide for the Care and Use of Nonhuman Primates in Neuroscience Research (The Japan Neuroscience Society; https://www.jnss.org/en/animal_primates) and were approved by the Animal Ethics Committee of the National Institutes for Quantum Science and Technology.
References
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Armbruster, B. N., Li, X., Pausch, M. H., Herlitze, S. and Roth, B. L. (2007). Evolving the lock to fit the key to create a family of G protein-coupled receptors potently activated by an inert ligand. Proc. Natl. Acad. Sci. U. S. A. 104(12): 5163–5168. doi: 10.1073/pnas.0700293104
Roth, B. L. (2016). DREADDs for Neuroscientists. Neuron 89(4): 683–694. doi: 10.1016/j.neuron.2016.01.040
Khateeb, K., Griggs, D. J., Sabes, P. N. and Yazdan-Shahmorad, A. (2019). Convection Enhanced Delivery of Optogenetic Adeno-associated Viral Vector to the Cortex of Rhesus Macaque Under Guidance of Online MRI Images. J. Vis. Exp. (147): e3791/59232-v. doi: 10.3791/59232-v
Lerchner, W., Corgiat, B., Der Minassian, V., Saunders, R. C. and Richmond, B. J. (2014). Injection parameters and virus dependent choice of promoters to improve neuron targeting in the nonhuman primate brain. Gene Ther. 21(3): 233–241. doi: 10.1038/gt.2013.75
Eldridge, M. A. G., Lerchner, W., Saunders, R. C., Kaneko, H., Krausz, K. W., Gonzalez, F. J., Ji, B., Higuchi, M., Minamimoto, T., Richmond, B. J., et al. (2016). Chemogenetic disconnection of monkey orbitofrontal and rhinal cortex reversibly disrupts reward value. Nat. Neurosci. 19(1): 37–39. doi: 10.1038/nn.4192
Oyama, K., Hori, Y., Nagai, Y., Miyakawa, N., Mimura, K., Hirabayashi, T., Inoue, K. i., Takada, M., Higuchi, M., Minamimoto, T., et al. (2022). Chronic Behavioral Manipulation via Orally Delivered Chemogenetic Actuator in Macaques. J. Neurosci. 42(12): 2552–2561. doi: 10.1523/jneurosci.1657-21.2021
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Nagai, Y., Miyakawa, N., Takuwa, H., Hori, Y., Oyama, K., Ji, B., Takahashi, M., Huang, X. P., Slocum, S. T., DiBerto, J. F., et al. (2020). Deschloroclozapine, a potent and selective chemogenetic actuator enables rapid neuronal and behavioral modulations in mice and monkeys. Nat. Neurosci. 23(9): 1157–1167. doi: 10.1038/s41593-020-0661-3
Miyakawa, N., Nagai, Y., Hori, Y., Mimura, K., Orihara, A., Oyama, K., Matsuo, T., Inoue, K. i., Suzuki, T., Hirabayashi, T., et al. (2023). Chemogenetic attenuation of cortical seizures in nonhuman primates. Nat. Commun. 14(1): e1038/s41467-023-36642-6. doi: 10.1038/s41467-023-36642-6
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A Methodology for the Enzymatic Isolation of Embryonic Hypothalamus Tissue and Its Acute or Post-Culture Analysis by Multiplex Hybridisation Chain Reaction
KC Kavitha Chinnaiya
MP Marysia Placzek
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4898 Views: 421
Reviewed by: Chiara AmbrogioMayank GautamRupkatha Banerjee
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Original Research Article:
The authors used this protocol in eLIFE Jan 2023
Abstract
The hypothalamus is an evolutionarily ancient part of the vertebrate ventral forebrain that integrates the dialogue between environment, peripheral body, and brain to centrally govern an array of physiologies and behaviours. Characterizing the mechanisms that control hypothalamic development illuminates both hypothalamic organization and function. Critical to the ability to unravel such mechanisms is the skill to isolate hypothalamic tissue, enabling both its acute analysis and its analysis after explant and culture. Tissue explants, in which cells develop in a manner analogous to their in vivo counterparts, are a highly effective tool to investigate the extrinsic signals and tissue-intrinsic self-organising features that drive hypothalamic development. The hypothalamus, however, is induced and patterned at neural tube stages of development, when the tissue is difficult to isolate, and its resident cells complex to define. No single molecular marker distinguishes early hypothalamic progenitor subsets from other cell types in the neural tube, and so their accurate dissection requires the simultaneous analysis of multiple proteins or mRNAs, techniques that were previously limited by antibody availability or were arduous to perform. Here, we overcome these challenges. We describe methodologies to precisely isolate early hypothalamic tissue from the embryonic chick at three distinct patterning stages and to culture hypothalamic explants in three-dimensional gels. We then describe optimised protocols for the analysis of embryos, isolated embryonic tissue, or cultured hypothalamic explants by multiplex hybridisation chain reaction. These methods can be applied to other vertebrates, including mouse, and to other tissue types.
Key features
• Detailed protocols for enzymatic isolation of embryonic chick hypothalamus at three patterning stages; methods can be extended to other vertebrates and tissues.
• Brief methodologies for three-dimensional culture of hypothalamic tissue explants.
• Optimised protocols for multiplex hybridisation chain reaction for analysis of embryos, isolated embryonic tissues, or explants.
Graphical overview
Keywords: Embryonic hypothalamus dissection Enzymatic tissue dissection Explant culture Multiplex hybridisation chain reaction (HCR) Stripping and re-probing protocol for HCR
Background
In the developing embryo, dynamic cellular and molecular processes direct early tissue patterning and subsequent organogenesis. These events have been deciphered through multiple experimental approaches, but one of the most indispensable tools in such investigations is the use of tissue or organ explants. While organs—including the central nervous system—can be isolated manually (Moore and Kennedy, 2008; Morrison et al., 2021), tissue isolation at patterning stages requires an enzymatic treatment. A variety of different enzymes can be used, including Collagenase, Trypsin, Dispase I, or Dispase II (Davies, 2010; Lowery et al., 2012; Morales et al., 2016; Shirahama et al., 2019; Ye et al., 2022), amongst which Dispase I is particularly effective (Placzek et al., 1990). This neutral protease can separate intact tissue layers, preserving the viability of each layer, and has frequently been used to isolate the posterior neural tube (Placzek et al., 1990; Yamada et al., 1993; Ericson et al., 1997; Tozer et al., 2013). Explants from isolated posterior neural tube regions, or adjacent tissues, have enabled an exquisite dissection of the cellular interactions and signalling events that orchestrate spinal cord patterning. In combination with post-hoc analysis through immunohistochemistry or in situ hybridisation, their use has helped to reveal how distinct progenitor subtypes are established along the dorso–ventral axis of the prospective spinal cord in response to dynamic morphogen gradients (Grega, 1984; Placzek et al., 1990; Yamada et al., 1993; Ericson et al., 1997; Tozer et al., 2013; Morales et al., 2016; Placzek and Briscoe, 2018).
The anterior neural tube that will give rise to the brain is more difficult to isolate due to its small size and dynamic morphogenesis. Nonetheless, where anterior neural tube explants have been used, they, too, have provided new insights into the tissue, cell, and molecular interactions that mediate brain patterning (Ericson et al., 1995; Guinazu et al., 2007; Placzek and Briscoe, 2018; Shirahama et al., 2019). In particular, their use has significantly improved our understanding of the development of the hypothalamus (Dale et al., 1997; Manning et al., 2006; Kim et al., 2022; Chinnaiya et al., 2023). This intricate region of the ventral forebrain regulates physiological processes that maintain homeostasis and orchestrate complex behaviours. The enzymatic isolation of entire embryonic chick hypothalamic tissues at different patterning stages, and their acute analysis through scRNA-seq, has generated a comprehensive roadmap of hypothalamic development, from induction to patterning and to neurogenesis, and revealed many previously uncharacterized candidate regulators of hypothalamic patterning (Kim et al., 2022). A powerful new development has been the use of multiplex hybridisation chain reaction (HCR) (Choi et al., 2018) in the analysis of wholemount and isolated hypothalamic tissues or hypothalamic tissue explants (Kim et al., 2022; Chinnaiya et al., 2023), for detecting and visualising specific DNA or RNA sequences in a sample. In the protocol we describe here, RNA probes specific to a target gene trigger chain reaction events between two sets of hairpin molecules, allowing for the rapid detection of mRNA (Choi et al., 2014). Multiple different mRNAs can be analysed on a single sample, and the sample can be repeatedly stripped and re-probed. Multiplex HCR has, therefore, significant advantages over more traditional in situ methods. In the complex environment of the developing brain, multiplex HCR has provided an incisive tool to define distinct progenitor subtypes and examine dynamic gene expression patterns. Multiplex HCR can similarly be applied to cultured hypothalamic tissue explants to reveal how naïve or pre-patterned tissues respond to extrinsic signals and/or self-organise (Kim et al., 2022; Chinnaiya et al., 2023). To date, these techniques have provided particularly powerful insights into the development of the tuberal hypothalamus (Chinnaiya et al., 2023).
Here, we describe how to isolate chick hypothalamus at three different stages, culture it as explanted tissue, and process acutely dissected tissue, or explants, by multiplex HCR. The protocols build on previous studies and cover Hamburger-Hamilton (HH) stage 6 to HH stage 25, the time span over which the hypothalamic tissue is induced, regionalised, and begins to undergo neurogenesis. These timepoints equate approximately to E7 and E11.5 of mouse development. We discuss key considerations in deciding how to approach dissection at different stages and steps that aim to minimise technical error and ensure that different samples can be compared. Importantly, the same methodologies can be widely applied to different vertebrates (including rodents), different regions of the brain, and tissues from different germ layers (e.g., Placzek et al., 1993).
Materials and reagents
Biological material
Brown Bovan fertilised chicken eggs (Medeggs Ltd)
Reagents for isolation and culture
Leibovitz’s L-15 medium with phenol red (Fisher Scientific, catalog number: 11415056)
Dispase Grade 1 (Roche, catalog number: 4942086001)
Corning collagen (Fisher Scientific, catalog number: 11563550)
Corning Matrigel GFR membrane matrix (Fisher Scientific, catalog number: 354236)
Explant medium:
Opti-MEM (Thermo Fisher Scientific, catalog number: 31985-062)
1% L-Glutamine (Fisher Scientific, catalog number: 11574466)
1% Penicillin/Streptomycin (Fisher Scientific, catalog number: 11528876)
< 3% Fetal calf serum (Sigma, catalog number: F2442)
0.8 M Sodium bicarbonate (NaHCO3), sterile (Sigma, catalog number: S5761)
10× DMEM (without bicarbonate, with phenol red, filtered, sterile) (Thermo Fisher Scientific, catalog number: 31600091)
Heat-inactivated goat’s serum (HINGS) (Scientific Laboratory Supplies, catalog number: G9023)
Bovine serum albumin (BSA) (Thermo Fisher Scientific, catalog number: B14)
Phosphate buffered saline tablets (Fisher Scientific, catalog number: 18912014)
Tween 100 (VWR, catalog number: A4974.0250)
70% Ethanol
Tissue paper
Reagents for multiplex hybridisation chain reaction
0.2 M Phosphate buffer pH 7.4 (PB): 6 g/L Sodium dihydrogen phosphate (NaH2PO4) (VWR Chemicals, catalog number: 28013.264), 21.8 g/L Di-sodium hydrogen phosphate (Na2HPO4) (VWR Chemicals, catalog number: 102494C)
4% Paraformaldehyde (PFA) (Sigma, catalog number: P6148) in 0.12 M PB: heat 10 mL of H2O to 65 °C, add 1 g of paraformaldehyde, mix by inverting, and add two drops of 1 M NaOH (VWR chemicals, catalog number: 28244.262). Invert until solution clears, make up to 25 mL with 0.2 M PB, and filter
Methanol (MeOH) (Fisher Scientific, catalog number: M/4450/17) series in PBS + 0.1% Tween 100 (VWR chemicals, catalog number: A4974.0250) for dehydration and rehydration of explants (25% MeOH/75% PBST; 50% MeOH/50% PBST; 75% MeOH/25% PBST; 100% MeOH)
Custom probes (Molecular Instruments)
Custom amplifiers (hairpin h1 and hairpin h2) (Molecular Instruments)
Hybridisation buffer (Molecular Instruments, catalog number: BPH01825)
Wash buffer (Molecular Instruments, catalog number: BPW02025)
Amplification buffer (Molecular Instruments, catalog number: BAM012125)
Proteinase K (Thermo Fisher Scientific, catalog number: 11444822)
Saline-sodium citrate (SSC) (Sigma, catalog number: S/3320/53)
SSC + 0.1% Tween 100 (SSCT)
DNase I Recombinant RNase-free Sol (ROCHE, catalog number: 4716728001)
Formamide (SLS, catalog number: 47670-1L)
OCT cryo-embedding matrix (Fisher Scientific UK, catalog number: 12678646)
Acetic anhydride (Sigma Aldrich, catalog number: 320102)
Triethanolamine (Sigma Aldrich, catalog number: 90279)
Imaging
Spinning disk confocal microscope (Nikon, model: W1)
Microscope (Zeiss, model: Apotome 2)
Microscope (Leica, model: MZ16F)
2% Low melting point agarose (Promega, catalog number: V2831)
4’,6-Diamidino-2-Phenylindole, dihydrochloride (DAPI) (Thermo Fisher Scientific, catalog number: D1306)
Anti-fade mounting medium (SLS, catalog number: F6182-20ML)
Glass-bottomed dishes (VWR, catalog number: 734-2906)
Equipment
Incubators at 18 ℃ for storing fertilised eggs and at 37 ℃ for incubating eggs to the right stages (Panasonic, PHCbi Programmable Cooled Incubator 238l)
37 ℃, 5% CO2 incubators for primary explant cultures
Dissection microscope with overhead and under lights (Leica MZ6 Stereomicroscope; Zeiss KL1500 LCD)
Dissection tools
Standard curved forceps (Fine Science Tools, catalog number: 11001-20)
Hardened fine scissors (Fine Science Tools, catalog number: 14090-11)
Vannas scissors (Fine Science Tools, catalog number: 15000-01)
Dumont No. 5 fine forceps (Fine Science Tools, catalog number: 11254-20)
Micro knives plastic handle (Fine Science Tools, catalog number: 10316-14)
Tungsten needles (holding and cutting) (Fine Science Tools, catalog numbers: 10130-05 and 20616-12)
Hamburger Hamilton staging guide (Hamburger and Hamilton, 1951)
Nunc 4-well dishes (Sigma, catalog number: 734-1175)
10 cm, 30 mm Petri dishes (SLS, catalog number: PET2000)
Eppendorfs (VWR, catalog number: 525-0794)
Tissue chopper (McILWAIN Tissue Chopper)
Razor blade (Wilkinson’s) (optional)
Water bath at 65 ℃
3 mL wide-bore Pasteur pipettes (VWR, catalog number: SLIN200C)
Gilson pipettes
Software and datasets
Axiovision software (Zeiss)
LAS X1.1.0.12420 imaging software
Fiji ImageJ2 Version 2.3.0/1.53q
Adobe Photoshop 2023 Version 24.7.0
Procedure
Removal of embryo from egg
The following section describes the removal of a chick embryo from the egg. The method is applicable to all patterning stages of development. Isolation can be performed outside a laminar flow hood, but eggs and instruments should be sterilised with 70% ethanol and, between use, instruments should be placed on sterilised tissue paper (autoclaving dissection tools is not necessary).
Incubate embryos in a 37 °C incubator to appropriate HH stage. Use standard fine scissors to window the eggshell and No. 5 fine-tipped forceps to carefully peel away outer and inner shell membranes and vitelline membrane, without damaging the embryo.
Use No. 5 fine-tipped forceps and standard fine scissors to cut around the embryo. Lift out and place in ice cold L-15 medium in a 10 cm Petri dish (Figure 1B).
Figure 1. Experimental setup for harvesting chick embryos. (A) Required dissection tools. (B) L-15 medium on ice and Brown Bovan fertilised eggs. (C) Examples of HH10 chick embryos after harvesting from the eggs and removal of membranes, yolk, and excess extra-embryonic tissue. Scale bar: 1 mm.
Remove the chorioallantoic membrane and any contaminating yolk. Transfer embryo (with a minimal amount of medium) to clean L-15 medium in a 30 mm Petri dish using a wide-bore 3 mL Pasteur pipette and assess developmental stage.
Cut away excess extra-embryonic tissue using Vannas scissors or an electrolytically sharpened tungsten needle, to leave the embryo surrounded by a small amount of extra-embryonic tissue (Figure 1C).
If required, analyse a small piece of embryonic tissue by PCR to determine sex. The protocol we describe below is equally applicable to male and female animals.
Top tip: 5 min rule. After removal from the egg, isolate the embryo from extraembryonic tissues as rapidly as possible (within 5 min). Transfer embryo to ice and keep on ice at all times to avoid tissue and RNA degradation.
Hypothalamic isolation
B1. Method 1
Isolation of neuroectoderm
The following section describes the isolation of anterior neuroectoderm at three different stages: neural plate [Hamburger-Hamilton (HH6–HH8)], neural tube (HH9–HH12), and phylogenetic (HH13–HH25) stages. The same protocol can be adapted to isolate tissues from other germ layers and other species. Dispase is particularly effective in rapidly and gently separating the neuroectoderm from other tissue layers, while preserving its integrity and viability.
Using tungsten needles, trim embryo in cold L-15 medium to isolate anterior embryonic regions, discarding posterior portions of the embryo and any remaining extraembryonic tissue (Figure 2A, 2F, 2K). Leave sufficient excess tissue to avoid damaging the hypothalamus at subsequent steps in the procedure, so make a posterior cut at the level of Hensen’s node (at HH6), somite 1 (at HH7–HH12), and midbrain (HH13–HH25).
Figure 2. Steps involved in Dispase treatment and sub dissection of hypothalamic tissue at neural plate, neural tube, and phylogenetic stages of development. (A, F, K) Ventral views of whole embryos (A, F) or side view of head (K) harvested from eggs before Dispase treatment. At late phylogenetic stages, the eye is removed prior to Dispase treatment. (B–D) Isolation at neural plate stages. (B) Boxed region in (A) following isolation and Dispase treatment. (C) HH6 neuroectoderm or mesendoderm, separated after Dispase treatment. Dotted outline shows translucent medial midline neuroectoderm cells. (D) Same isolated neuroectoderm as in (C), trimmed to remove head fold and ectoderm. Boxed region shows prospective hypothalamus. (G–I) Isolation at neural tube stages. Note: routinely, the boxed region in (F) would be isolated and treated with Dispase. However, to better show tissue separation, the view in (G) shows an entire HH10 embryo after Dispase treatment. Dotted line in (G) outlines neuroectoderm separating from adjacent mesendoderm. (H, I) Ventral views of isolated anterior neuroectoderm, visualised under the dissecting microscope (H) or brightfield illumination after DAPI labelling (I). Boxed regions show developing hypothalamus, which develops around morphologically distinct neuroepithelial folds. (K–O) Isolation at phylogenetic stages. (L) Side view of HH24 head following Dispase treatment. Optic stalks and hypothalamus start to become morphologically distinct. (M) Isolated neuroectoderm: circled region marks hypothalamus and optic stalks. (N) Side view shows sub-dissected neuroectoderm (circled region in (M). Ellipse outlines prospective hypothalamus which protrudes ventrally between the morphologically distinct optic stalks and the cephalic flexure. (O) Open-book preparation of tissue shown in (N). The violin-shaped hypothalamus is morphologically distinct. (E, J, P) Schematics of HH6, HH10, and HH24 neuroectoderm and highlighted region shows the dissected region of interest. Ellipse outlines hypothalamus. Abbreviations: HYP: hypothalamus; ME: mesendoderm; NE: neuroectoderm; OS: Optic stalk, CF: cephalic flexure. Scale bar: 250 μm.
Transfer the anterior embryonic region into 0.5 mL of freshly made Dispase (1 mg/mL) in L-15 medium in a Nunc 4-well dish at room temperature. Depending on the size of tissue, transfer using siliconised glass pipette, appropriately sized Gilson pipette (P200 or P1000), or forceps (from small to large size, respectively), taking care to transfer minimal amounts of L-15 medium. The length of time of incubation is highly stage dependent (see Table 1). A rule of thumb is to observe the embryos until the tissue layers begin to visibly separate: the mesendoderm shrinks relative to ectoderm (Figure 2B, 2G, 2L). Once the reaction is complete, transfer the embryo regions from Dispase solution into a large volume of cold L-15 medium in a 10 cm dish. Allow embryos/embryonic regions to rest for 5–10 min.
Table 1. Duration of Dispase treatment for the different chick developmental stages
HH stage Duration of 1 mg/mL Dispase treatment
HH6 5 min
HH8 6–8 min
HH10 10–12 min
HH13/14 15–17 min
HH17–20 18–20 min
HH21–25 20–25 min
Top tip 1: Transferring embryo regions between dishes must be done with care to ensure that they do not become stuck in the pipette or rise and burst at the meniscus [the latter occurs readily if embryos are transferred from a warm to a cold solution, e.g., Dispase solution (room temperature) to cold L-15 medium]. To avoid, transfer under the microscope, avoid air bubbles, and transfer only a few embryo regions at a time (3–4).
Top tip 2: It is essential to optimise the Dispase concentration and incubation period. Tissues exposed to low concentrations of Dispase (even for longer incubation periods) will fail to separate; those exposed to high concentrations of Dispase, or incubated for too long, will become sticky, leading to a failure to obtain cleanly isolated tissue layers.
Top tip 3: For mouse tissues, which are more delicate than chick, reduce time in Dispase by 20%.
Top tip 4: Dispase-treated tissue fragments are transferred with a minimal volume of liquid into a large volume of L-15 medium, negating the need to inactivate the enzyme. However, if necessary, Dispase can be inactivated using a drop of 10% HINGS or BSA.
Hypothalamus dissection
The following section describes the isolation of hypothalamic tissue at neural plate, neural tube, and phylogenetic stages.
Use tungsten needles (some prefer a combination of tungsten needles and micro knives) to dissect the neural tissue (i.e., neural plate, neural tube, or brain, depending on embryonic stage) away from other germ layers. To do so, use a holding needle in one hand to pin down the embryo region (avoid pinning down the hypothalamus), and use the long edge of the second needle to gently stroke away the mesendoderm, paring it from adjoining neuroectoderm. The ease and details through which this is achieved is stage dependent. At neural plate stages, the mesendoderm can be easily stroked away from adjacent neuroectoderm. The tissues can be readily distinguished: neuroectoderm is smoother and larger than mesendoderm (Figure 2C). The anterior–posterior axis can be distinguished through the headfold; the neural plate medial midline (which harbours prospective floor plate and hypothalamus) is obvious through its translucent appearance (Figure 2C, dotted outline). At neural plate stages, the neuroectoderm is contiguous with ectoderm, which will provide a useful holding tissue at subsequent stages of dissection. At neural tube and phylogenetic stages, mesendoderm and neuroectoderm do not separate as easily, and mesendoderm must be carefully peeled away around the optic vesicles, optic stalk and eyes, and hypothalamus. This can be achieved using tungsten needles and (where tissues are larger) sharp No. 5 forceps. Once isolated, the neural tissue should appear smooth and epithelial-like, with no contamination of round mesenchymal/mesendoderm cells (Figure 2H, 2M). Transfer isolated neuroectoderm into cold L15 medium using a siliconised glass Pasteur pipette or an appropriately sized Gilson pipette (P10, P20), again ensuring that the tissue does not rise to meniscus and bursts.
Once the neural tissue is dissected, sub-dissect the hypothalamus (Figure 2D, 2H, 2I, 2M, 2N) using morphological criteria to define its limits. At HH6, the prospective hypothalamus is situated in and around anterior-most medial midline cells. Like more posterior midline cells, these are translucent but can be distinguished from these because they are slightly wider (Figure 2D). At HH10, the nascent hypothalamus occupies an area centred around a series of neuroepithelial folds in the ventral neuroectoderm. While most obvious after contrast microscopy (e.g., Figure 2I), the folds are morphologically visible under the dissecting microscope (Figure 2H) and provide a precise reference point for hypothalamic position. At later phylogenetic stages, the hypothalamus protrudes ventrally between the morphologically distinct optic chiasm and cephalic flexure (Figure 2L, 2M). Isolation of hypothalamic tissue is achieved in two steps. First, a domain encompassing the optic stalk, hypothalamus, and more dorsal diencephalic tissue is dissected using tungsten needles (limits shown by ellipse in Figure 2M, isolated region shown in Figure 2N). Second, this tissue is prepared as an open book. The hypothalamus is obvious as a violin-shaped domain, composed of a translucent medial domain and a thicker surrounding area, situated posterior to the optic stalks/developing optic chiasm (ellipse in Figure 2O).
Top tip 1: 5 min rule. Except for the Dispase step, avoid removing the embryo from ice-cold L-15 medium for more than 5 min. With practice, the neuroectoderm can be isolated from the mesendoderm within 30–60 s (early stages) and 2–3 min (late stages). A good rule of thumb is to dissect one embryo at a time.
Top tip 2: At neural plate stages, proceed to dissect out the hypothalamus (Figure 2C) as rapidly as possible, or midline cells lose their translucent appearance.
Top tip 3: Routinely sharpen the cutting tungsten needle.
Top tip 4: Dissect in a relaxed and calm environment. Remember to breathe and keep your shoulders loose and back straight.
B2. Alternate slicing method
In the previous method, the anterior neural plate/neural tube is first isolated using Dispase, and the hypothalamus (or prospective hypothalamus) is then sub-dissected. Some find this approach difficult and prefer the following alternate method, in which embryos are first sliced into fragments, including a hypothalamic-containing fragment that is then treated with Dispase, and hypothalamic tissue isolated. This method is particularly useful for neural tube–stage embryos, where some find it difficult to pare away mesendoderm from the neuroepithelial folds that characterise the nascent hypothalamus at this stage.
Transfer the dissected embryo to a clean 5 cm square plastic plate with as little liquid as possible (it must not dry out but must not float) and arrange it so it is flat (Figure 3A).
Figure 3. Isolation of hypothalamus tissue using the slicing method. (A) HH9 embryo laid flat on a plastic plate. (B) Top view of tissue chopper, with plastic plate and embryo (dotted outline and arrow) ready for slicing. (C) Front view of tissue chopper. (D) HH9 embryo sliced into 150 μm thick slices. (E) Anterior-most slices, removed from the plastic holder, arranged from anterior (top) to posterior (bottom). Arrows point to hypothalamus-containing slice. (F) Examples of HH10 slices, each containing hypothalamus, after Dispase treatment. (G) Slice containing hypothalamus from a HH9 embryo after Dispase treatment: dotted outline demarcates neuroectoderm and mesendoderm. (G’–G”) Same slice after isolation of neuroectoderm (G’) and sub-dissection to obtain hypothalamic explant (G”). Scale bar: 250 μm.
Set the plate with the embryo onto the base of the tissue chopper, with the anterior–posterior axis of the embryo at a right angle to the tissue chopper blade (Figure 3B).
Set the thickness of each slice by turning the thickness dial at the front (Figure 3C) and use a new razor blade for each experiment to keep the sample sterile.
Move the holder base to the start position and press start.
Once the tissue is sliced (Figure 3D), carefully transfer anterior slices to L-15 medium in a 30 mm Petri dish using a siliconised glass pipette or Gilson (P2 or P10). Each slice has a distinctive morphology. Slices containing the hypothalamus are thicker than more anterior slices, due to the folded neuroepithelium and the underlying mesendoderm, and have a wider neural tube than more posterior slices (Figure 3E).
Transfer hypothalamus slices to Dispase for 5 min (Figure 3F); then, isolate the neural tissue by gently stroking away the adjacent mesendoderm, as above. Once freed from adjacent tissue, the neuroepithelium will constrict slightly (Figure 3G, 3G’), but the ventral midline containing the nascent hypothalamus remains distinct and can be sub-dissected with tungsten needles (Figure 3G”).
Isolated hypothalamic tissue can be subjected to a wide range of further investigations, ranging from tests of function in vivo (e.g., grafting it to ectopic locations or cell dissociation for scRNA-Seq analyses) to tests of function ex vivo (e.g., neurospherogenic competence or ex vivo cultures). Here, we focus on the investigation of hypothalamic tissue explants in 3D culture systems.
Tissue embedding for three-dimensional explant culture
Tissue explants are a highly effective tool to investigate the extrinsic signals and tissue-intrinsic self-organising features that orchestrate hypothalamus development. They are versatile and can be used to assay patterning, proliferation, and migration. Explant cultures may consist of the entire hypothalamus or smaller, sub-dissected domains (limited by size). Good viability and the development of explants in a manner analogous to their in vivo counterparts occurs when explants are relatively small. Explants of this size can be cultured for up to seven days, after which tissue crowding and cell death begin to be observed in the explant centre. When used to assay the effects of signalling ligands or inhibitors, these factors can be added along with explant medium. Explants can be embedded in different matrices, most frequently in collagen or Matrigel. Explants are placed on a bottom layer of gelled collagen or Matrigel, then covered with a second layer, and positioned at the interface of the layers. Previous reports have provided detailed methodologies for how to embed explants in 3D matrices (Placzek and Dale, 1999; Placzek, 2008), so here we provide only brief details.
For collagen:
Prepare the desired volume (25 μL/bed/well) of 90% collagen and 10% 10× DMEM in an Eppendorf and vortex for 15–20 s. Add 0.8M NaHCO3 to make the solution turn pale yellow after vortexing. Typically, 2–6 μL of NaHCO3 is added to 100 μL of collagen/10× DMEM. Collagen, if unused, should be left on ice as it will start to set at room temperature. Always make fresh batches of collagen for bottom and top layers.
In a Nunc 4-well dish, prepare collagen beds by pipetting 20–25 μL of collagen, spreading it into a flatbed (Figure 4A). Allow to set (20–30 min at room temperature). Flat collagen beds are easier to embed the tissue as the tissue tends to slide off a convex bed.
Figure 4. Hypothalamic tissue embedding and culture. (A) Nunc 4-well dishes with Matrigel beds. (B) Single hypothalamic explant after dissection (inset) and embedding (arrow), prior to culturing. (C) Hypothalamic explants cultured after 72 h. (D) High power view of explant cultured for 72 h. Scale bar: (B, C) 250 μm, (D) 100 μm.
Transfer explants onto the collagen bed using a siliconised glass pipette/appropriately sized Gilson (P2, P10, P20), position them, and remove excess medium using a fine pipette (Figure 4B). Overlay with 25 μL of collagen, spreading this to ensure it covers the bottom bed. Reposition/manipulate the explants as required while the collagen is setting. Collagen sets quickly at room temperature so work rapidly and add top collagen layer to one well at a time, providing the opportunity to reposition explants before the collagen begins to set.
Allow the top collagen bed to set completely at room temperature (check by prodding the collagen with a tungsten needle; set collagen feels firm) before adding 400–500 μL of explant medium with or without factors and transfer to incubator for the desired time of incubation (Figure 4C, 4D). Routinely, we culture explants for 3 h to 7 days.
For Matrigel:
Prepare desired volume (25 μL/bed/well) of 50% Matrigel and 50% explant medium in an Eppendorf and vortex for 15–20 s. If unused, Matrigel should be left on ice as it will start to set at room temperature. Always make a fresh batch of Matrigel for bottom and top beds.
Proceed with steps 2–4 as above for collagen, but beds must be set by placing dish at 37 °C for a minimum of 30 min (check by prodding Matrigel with a tungsten needle; set Matrigel feels firm).
Data analysis
Analysis by multiplex HCR
Embryos and cultured explants can be analysed by immunohistochemistry (Manning et al., 2006), flow cytometry (Perez et al. 2023), chromogenic in situ hybridisation (Manning et al., 2006), or multiplex hybridisation chain reaction (HCR). In our hands, we have found that immunohistochemistry and chromogenic in situ work equally well for explants embedded in either collagen or Matrigel, but HCR works well only in explants embedded in Matrigel. Previous reports have described methodologies for multiplexed quantitative HCR (Choi et al., 2020) and for its extension to immunohistochemistry (Schwarzkopf et al., 2021). Here, we extend these reports to provide details on how to take embryos and explants through repeated rounds of multiplex HCR. Such stripping and re-probing enables samples to be routinely analysed with 10–12 genes per sample (Chinnaiya et al., 2023) (Figure 5).
Figure 5. Wholemount multiplex hybridisation chain reaction (HCR) analysis on embryos and explants (A–D) HCR analysis on hemi-dissected HH20. Views show hypothalamic region in four sequential rounds of HCR following stripping and re-probing. Tuberal regions of the hypothalamus express ISL1, SIX6, SHH, FGF10, TBX2, RAX, and NKX2-1; hypothalamic mammillary and supramammillary regions express FOXA2, PITX2, and EMX2. NKX2-1 is additionally expressed in the ventral telencephalon and PAX6 in the dorsal diencephalon (from (Chinnaiya et al., 2023). (B) Multiplex HCR analyses on wholemount HH6 explants embedded in Matrigel and cultured for 72 h. The explants have been taken through four rounds of HCR following stripping and re-probing. In this example, the explant included prospective tuberal hypothalamus and some eye tissue. Scale bar: 100 μm.
Embryos and explants can each be processed as wholemounts or as cryosections, as detailed below. After completion of the HCR, neuroectoderm can be particularly well visualised either by hemi-dissecting embryos along a sagittal midline (we refer to these as hemiviews), or by isolating the neuroectoderm. Depending on the stage, this can be achieved manually or may require the use of Dispase, as above. Explants are not removed from the Matrigel during the HCR.
Hybridisation chain reaction
Analysis of wholemount embryos or explants
Pre-incubation stage
Fix, dehydrate, then rehydrate the samples with a series of Methanol/PBST washes for 5 min each on ice.
Incubate samples in Proteinase K (10 μg/mL) for 2–5 min (depending on sample developmental stage) at room temperature.
Postfix samples in 4% PFA at room temperature for 20 min.
Wash samples with PBST, PBST + 5× SSCT (50:50), and then with 5× SSCT for 5 min each on ice.
Detection stage
Pre-incubate samples in hybridisation buffer for 30 min at 37 °C.
Make up the desired probe solution by adding 2–10 μL of 1 μm stock probe in 500 μL of hybridisation buffer and incubate samples in the probe solution overnight at 37 °C.
Next day, remove excess probes by washing samples 4 × 5 min in wash buffer at 37 °C.
Wash samples 2 × 5 min with 5× SSCT at RT.
Amplification stage
Incubate samples in amplification buffer for 5 min at room temperature.
Prepare 30 pmol of hairpin h1 and 30 pmol of hairpin h2 in separate tubes by snap-cooling 10 μL of 30 μM stock (heat at 95 °C for 90 s and cool to room temperature in a dark drawer for 30 min).
Prepare hairpin mixture by adding snap-cooled h1 and h2 hairpins to 500 μL of amplification buffer at room temperature.
Incubate samples in hairpin mixture in the dark overnight at room temperature.
Next day, remove hairpins by washing with 5× SSCT at room temperature.
2 × 5 min
2 × 30 min
1 × 5 min
Incubate samples in 1:1,000 DAPI (1 mg/mL) for 5–10 min in 5× SSCT.
Remove excess DAPI by washing with 5× SSCT.
Samples can be stored at 4 °C protected from light before microscopy.
Note: Leave the samples on a rocker for all the steps except when in Proteinase K.
Stripping and re-probing
Following imaging the first round of probes, wash samples 2 × 5 min with 5× SSCT.
Pre-incubate samples in 1× DNase buffer (100 μL/mL in H2O) solution for 5 min.
Incubate samples in DNase1 solution (1:50, 10,000 U/mL) overnight at 37 °C.
Note: For 500 μL of DNase1 solution: 50 μL of DNase1 + 50 μL of 10× DNase buffer + 400 μL of ddH2O.
Check samples under the microscope to confirm that the signal from the previous round is gone.
Next day, wash samples 3× 10 min with 30% Formamide in 2× SSCT at 37 °C.
Wash samples (3× 10 min) with 2× SSCT at 37 °C.
Follow the detection and amplification steps.
Note: Stripping and re-probing can be repeated successfully for 3–6 rounds with different probe sets.
Image acquisition and analysis
Samples taken through HCR analyses can be imaged as flat mounts on a depression slide with anti-fade mounting medium or after embedding in 2% low melting point agarose on a glass-bottomed dish.
Samples can be stored at 4 °C protected from light before microscopy.
Samples are imaged as z-stacks using Nikon NIS elements or Zeiss Axiovision software and displayed as maximum intensity projections.
Image analysis is performed using Fiji ImageJ software.
Analysis of cryo-sectioned embryos or explants
For detailed protocol on OCT mounting and cryosection of samples, refer to Manning et al. (2006).
Pre-incubation stage
Pre warm humidification chamber at 37 °C.
Place slides flat on a slide holder and wash with 1× PBS for 5 min to remove any OCT.
Post-fix with 4% PFA for 10 min at room temperature. Wash slides 3 × 5 min with PBS.
Incubate samples in Proteinase K (10 μg/mL) for 2–5 min (depending on sample developmental stage) at room temperature and fix sections again for 10 min. Wash slides 3 × 5 min in PBS.
Treat slides with acetylation mix for 10 min: 11.2 μL of Triethanolamine + 2.5 μL of acetic anhydride in 1 mL of ddH2O. Wash slides 3 × 5 min in PBS.
Detection stage
Pre-incubate slides in hybridisation buffer for 1 h at 37 °C in a humidified chamber.
Prepare probe solution as described above in Detection stage.
Remove hybridisation solution, apply 200 μL of probe solution to each slide, coverslip the slides, seal the sides of the humidified chamber with Sellotape, and incubate at 37 °C overnight.
Next day, immerse slides in probe wash buffer at 37 °C to float off the coverslip.
Remove excess probe by incubating slides at 37 °C in:
75% of probe wash buffer/25% 5× SSCT for 15 min.
50% of probe wash buffer/50% 5× SSCT for 15 min.
25% of probe wash buffer/75% 5× SSCT for 15 min.
100% 5× SSCT for 15 min.
Wash slides in 5× SSCT for 5 min at room temperature.
Dry slides by blotting edges on tissue paper.
Amplification stage
Add 200 μL of amplification buffer and pre-amplify for 30 min at room temperature.
Prepare hairpin mixture as described above and add 200 μL of hairpin mixture, coverslip, and incubate overnight at room temperature.
Next day, immerse slides in 5× SSCT to float off the coverslip.
Remove excess hairpins by washing with 5× SSCT at RT for:
1× 5 min.
2× 30 min.
1× 5 min with DAPI.
1× 5 min.
Dry slides by blotting edges on tissue paper.
Add 50–100 μL of anti-fade mounting medium and mount the coverslip onto the slide for microscopy.
Stripping and re-probing
Following imaging the first round of probes, wash samples 2× 5 min with 5× SSCT.
Pre-incubate in 1× DNase buffer (100 μL/mL in H2O) solution for 5 min.
Incubate samples in DNase1 solution (1:20, 1,000 U/mL) for 4 h at 37 °C.
Note: For 500 μL of DNase1 solution: 25 μL of DNase1 +50 μL of 10× DNase buffer + 425 μL of ddH2O.
Check slides under the microscope to confirm that the signal from the previous round is gone.
Next day, wash samples 3× 10 min with 30% Formamide in 2× SSCT at 37 °C.
Wash 3 × 10 min wash with 2× SSCT at 37 °C.
Follow the detection and amplification steps.
Note: Stripping and re-probing can be repeated successfully to 3–6 rounds with different probe sets.
Image acquisition and analysis
Samples taken through HCR analyses can be mounted with anti-fade mounting medium and coverslipped.
Slides can be stored at 4 °C protected from light before microscopy.
Samples are imaged as single stack or z-stacks with Zeiss Axiovision software and displayed as maximum intensity projections.
Image analysis is performed using Fiji ImageJ software.
Top tip 1: In some cases, morphological criteria can be used to align images from different rounds. When this is not possible, a reference probe can be re-deployed in each round.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Chinnaiya et al. (2023). A neuroepithelial wave of BMP signalling drives anteroposterior specification of the tuberal hypothalamus. eLife (Figure 1, panel T-T’’; Figure 3; Figure 4; Figure 5 panel E–H; Figure 6.
The robustness and reproducibility of our protocol can be evidenced from the above research article.
Acknowledgments
This research was funded in whole, or in part, by the Wellcome Trust [212247/Z/18/Z] to M.P. For the purpose of Open Access, the author has applied a CC BY public copyright licence to any Author Accepted Manuscript version arising from this submission.
This protocol was derived from the original work of Chinnaiya et al. (2023).
Competing interests
The authors declare that no competing interests exist.
References
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An Improved Protocol for the Matrigel Duplex Assay: A Method to Measure Retinal Angiogenesis
KB Kathleen C. Brown
RL Reagan S. Light *
KM Kushal J. Modi *
KC Kaitlyn B. Conely *
AS Amanda M. Sugrue
AC Ashley J. Cox
SM Sarah L. Miles
MV Monica A. Valentovic
Piyali Dasgupta
(*contributed equally to this work)
Published: Vol 13, Iss 23, Dec 5, 2023
DOI: 10.21769/BioProtoc.4899 Views: 543
Reviewed by: Vivien J. Coulson-ThomasTarsis Gesteira FerreiraSudhir Verma
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Original Research Article:
The authors used this protocol in Investigative Ophthalmology & Visual Science Jun 2011
Abstract
Neovascular diseases of the retina, such as diabetic retinopathy (DR) and age-related macular degeneration (AMD), are proliferative retinopathies involving the growth of new blood vessels on the retina, which in turn causes impairment and potential loss of vision. A drawback of conventional angiogenesis assays is that they are not representative of the angiogenic processes in the retina. In the retina, the new blood vessels grow (from pre-existing blood vessels) and migrate into a non-perfused region of the eye including the inner limiting membrane of the retina and the vitreous, both of which contribute to vision loss. The Matrigel Duplex Assay (MDA) measures the migration of angiogenic capillaries from a primary Matrigel layer to a secondary Matrigel layer, which resembles the pathological angiogenesis in AMD and DR. The methodology of MDA is comprised of two steps. In the first step, the human retinal microvascular endothelial cells (HRMECs) are mixed with phenol red–containing Matrigel (in a 1:1 ratio) and seeded in the center of an 8-well chamber slide. After 24 h, a second layer of phenol red–free Matrigel is overlaid over the first layer. Over the course of the next 24 h, the HRMECs invade from the primary Matrigel layer to the secondary layer. Subsequently, the angiogenic sprouts are visualized by brightfield phase contrast microscopy and quantified by ImageJ software. The present manuscript measures the angiogenesis-inhibitory activity of the Src kinase inhibitor PP2 in primary HRMECs using the MDA. The MDA may be used for multiple applications like screening anti-angiogenic drugs, measuring the pro-angiogenic activity of growth factors, and elucidating signaling pathways underlying retinal angiogenesis in normal and disease states.
Graphical overview
Keywords: Angiogenesis MDA HRMEC PP2 Quantification
Background
The retina consists of organized layers of photoreceptors, interneurons, glia, epithelial cells, and endothelial cells. The proper maintenance of vascular networks is critical to normal visual function [1]. Aberrant angiogenesis is the hallmark of several ocular diseases including age-related macular degeneration (AMD) and diabetic retinopathy (DR) [2]. The angiogenic process in the retina is a complex, multistep process involving endothelial cell invasion, adhesion, chemotactic migration, proliferation, and differentiation into capillary tube–like structures and the production of a basement membrane around the vessel [3]. A survey of literature shows that the Matrigel capillary tube assay is one of the most prevalent angiogenesis assays in cell culture models [4, 5]. In this assay, human microvascular endothelial cells are seeded on a three-dimensional layer of solidified Matrigel (or any other reconstituted basement membrane extracellular matrix). Over a period of 24 h, the cells differentiate into capillary tube–like networks, which can be quantified by digital image analysis. A caveat of this assay is that it does not represent pathological angiogenesis in the eye. The Matrigel assay is more representative of vasculogenesis, which is defined as the differentiation of endothelial cells to yield de novo primitive vascular networks rather than angiogenesis where new capillaries are generated from existing vasculature [6]. Another disadvantage is the lack of a lumen in the capillaries obtained by this assay [7, 8]. Other methods used to measure retinal angiogenesis include the measurement of endothelial cell proliferation or endothelial cell chemotaxis or migration. Although these individual processes are useful indicators of angiogenic activity, they do not provide a holistic representation of the angiogenic cascade [7, 8]. These drawbacks are circumvented by using the Matrigel Duplex Assay (MDA). In the MDA, the angiogenic sprouting (in the secondary Matrigel layer) arises from pre-formed vascular networks in the primary layer [9, 10].
The MDA represents a combination of all the steps of retinal angiogenesis and provides a highly relevant model for the study of pro- and anti-angiogenic agents in vitro. In-depth morphological studies have shown that the angiogenic sprouting (occurring in the MDA) closely mimics retinal angiogenesis in vivo. Electron microscopy studies have demonstrated the presence of a lumen and elaborate cell–cell junctions within the endothelial cell aggregates observed in the MDA [9, 10]. As the endothelial capillary tube–like structures invade into the secondary Matrigel layer, the leading edge of the capillary sprouts is associated with long filopodia. The structure of these filopodia resembles those seen at the angiogenic front during developmental angiogenesis in the neonatal retina [9]. The retinal endothelial cells grown within the two layers of Matrigel reflect the biological characteristics of the neovascular retina, such as diffusion gradient of oxygen, nutrients, and pH. The growth of the cells inside the Matrigel duplex system allows for complex cell–cell and cell–matrix interaction. Furthermore, these retinal endothelial sprouts may be characterized and quantified by confocal microscopy [9].
Stitt et al. (2005) were the first to report the protocol for the MDA to study retinal angiogenesis in diabetic retinopathy [10]. During the application of the original protocol of Stitt et al. (2005) in our laboratory, some technical challenges occurred. The original protocol uses phenol red–containing Matrigel for both the primary and the secondary Matrigel layers [9, 10]. We observed that the interface between the two layers was difficult to visualize by phase contrast microscopy. This is especially true when there is a dense network of retinal capillaries migrating from the primary to the secondary Matrigel layer in the MDA. In order to circumvent this issue, we used a black Sharpie marker pen (in our early studies) to draw a boundary encircling the first layer before adding the second Matrigel layer [11]. This approach had its limitations as it was difficult to accurately demarcate the boundary of the first layer. Secondly, the black boundary around the primary layer appeared as a thick black band under the brightfield microscope, increasing the difficulty of accurately visualizing and quantifying the angiogenic capillary sprouts in the second layer. We drew the dotted line in the center of this black band to quantify the invasion of endothelial capillary networks into the secondary layer [11] and the whole process became arduous and cumbersome.
We devised a novel strategy to clearly differentiate between the two Matrigel layers in the MDA by utilizing phenol red–free Matrigel (PR-free-Matrigel) and phenol red–containing Matrigel (PR-Matrigel) for the individual layers. Initially, we tried to use PR-free-Matrigel for the primary layer and PR-Matrigel for the secondary layer but found that this arrangement resulted in poor quality photographs under a microscope. Subsequently, we reversed the layers (using the PR-Matrigel in the primary layer and PR-free-Matrigel in the secondary layer) and obtained clearer pictures under phase contrast microscopy, providing readily reproducible images and assay results. The present manuscript describes the ability of the Src Kinase inhibitor PP2 [12] (at a concentration of 10 µM) to suppress retinal angiogenesis of human retinal microvascular endothelial cells (HRMECs) using our modified MDA protocol. We hope that the MDA will be a useful tool for researchers working in the field of physiological and pathological angiogenesis in the retina.
Materials and reagents
Primary human retinal microvascular endothelial cells (HRMECs) (Cell Systems, catalog number: ACBRI 181). Grow HRMECs in endothelial growth media-2 (EGM-2) [EGMTM-2 Endothelial Cell Growth Medium-2 BulletKitTM (Lonza, catalog number: CC-3162)]. The BulletKitTM contains EBM-2 basal medium (catalog number: CC-3156) and SingleQuots growth factor supplements (catalog number: CC-4176) needed for the growth of HRMECs. The BulletKitTM also contains a vial of antibiotic (Gentamycin) and fetal bovine serum (FBS). Store the EBM-2 at 4 °C and the SingleQuots and FBS at -20 °C. Prepare the EGM-2 media following the manufacturer’s instructions provided in the BulletKitTM. Add FBS to the media at a final concentration of 2% (see Recipes). The EGM-2 medium with 2% FBS will be hereafter referred to as EGM-2 complete medium
RPMI-1640 medium (ATCC, catalog number 30-2001)
Trypsin-EDTA (0.25% Trypsin, 0.53 mM EDTA) (ATCC, catalog number: 30-2101). Long-term storage at -20 °C; however, aliquot Trypsin-EDTA and store at 4 °C for immediate use
Dulbecco’s Phosphate buffered saline (DPBS) without calcium and magnesium (Corning, Fisher, catalog number: 21-031-CM). Store at 4 °C
Absolute anhydrous ethanol (200 proof) (Pharmaco, Greenfield Global, catalog number: 111000200). Dilute ethanol with distilled water to obtain a 70% ethanol solution for wiping down the laminar flow hood
PR-Matrigel membrane matrix (Fisher, catalog number: CB-40234). Aliquot the Matrigel in Corning sterile freezing vials and store at -70 °C for long-term storage. Before the experiment, keep the Matrigel aliquot overnight at 4 °C (in a refrigerator) for thawing. During the assay, Matrigel should be kept on ice throughout the experiment
PR-free-Matrigel membrane matrix (Fisher, catalog number: CB-40234C)
PP2 (Enzo Biosciences, catalog number: 50-201-0681). PP2 is a potent and selective inhibitor of the Src family tyrosine kinases). Store PP2 at -20 °C and solubilize in DMSO (Fisher, catalog number: MT-25950CQC) to yield the stock solution at concentration of 10 mM (see Recipes). Aliquot the PP2 stock solution into microcentrifuge tubes and store at -20 °C. Immediately before use, thaw and dilute the PP2 in EBM-2 basal media (without FBS or growth factors) to a working concentration of 10 µM. After use, discard the remaining solution of 10 µM PP2
Tissue culture plasticware and related supplies
Corning cell counting chamber (Fisher, catalog number: 07-200-988)
Nunc Lab-Tek 8-well Permanox plastic chamber slide system (Fisher, catalog number: 12-565-22)
Corning BioCoat Collagen-coated T-25 (Fisher, catalog number: 08-774-327) and T-75 (Fisher, catalog number: 08-774-327) tissue culture flask. The BioCoat flasks are stored at 4 °C
Corning sterile cryogenic vials (Fisher, catalog number: 13-700-504)
Sterile polypropylene centrifuge tubes, 50 mL (Fisher, catalog number: 06-443-20)
Sterile polypropylene centrifuge tubes, 15 mL (Fisher, catalog number: 05-539-5)
Safety vented wash bottles (1,000 mL) for storing 70% ethanol (Fisher, catalog number: 11-865-170)
Microcentrifuge tubes, 1.5 mL (Fisher, catalog number: 05-408-129). Autoclave the microcentrifuge tubes before using them for the MDA
Sterile 5 mL microcentrifuge tube (Eppendorf, catalog number: 0030119487)
Sterile plastic serological pipettes (individually Wrapped, paper peel packaging, plugged):
25 mL pipette (Fisher, catalog number: 12-567-604)
10 mL pipette (Fisher, catalog number: 12-567-603)
5 mL pipette (Fisher, catalog number: 12-567-602)
2 mL pipette (Fisher, catalog number: 12-567-601)
1 mL pipette (Fisher, catalog number: 12-567-600)
Fisherbrand 5 3/4 inches’ disposable borosilicate glass Pasteur pipettes (Fisher, catalog number: 13-678-20B). Autoclave the Pasteur pipettes before using them for the MDA
Fisherbrand sterile high precision #22 style scalpel blade (Fisher, catalog number: 12-000-161)
DrummondTM Portable Pipet-AidTM XP pipette Controller (Fisher, catalog number: 13-681-15E)
PIPETMAN® 4-Pipette kit P2, P20, P200, P1000 (Gilson, catalog number: F167360)
Fisherbrand SureOne sterile aerosol barrier tips (100–1,000 µL) (Fisher, catalog number: 02-707-404)
Fisherbrand SureOne sterile aerosol barrier tips (20–200 µL) (Fisher, catalog number: 02-707-430)
Fisherbrand SureOne sterile aerosol barrier tips (0.1–20 µL) (Fisher, catalog number: 02-707-470)
Other consumable laboratory supplies
Labcoat (Fisher, catalog number: 19-181-596)
Ansell MICROFLEXTM NeoProTM NEC-288 Neoprene small size (Fisher, catalog number: 19-350-340F)
and medium size gloves (Fisher, catalog number: 19-350-340A)
Freezer boxes for long-term storage of Matrigel and PP2 (Cole-Parmer Essentials 81-Place Freezer Boxes, catalog number: UX-04396-63)
Kimtech Kimwipe delicate task wipers (Fisher, catalog number: 06-666A)
Sterile nonwoven gauze sponges 4 × 4 inch (Fisher, catalog number: 22-028-558)
Hayman style microspatula (Fisher, catalog number: 21-401-25A)
Solutions
EGM-2 complete medium (see Recipes)
EGM-2 media (with growth factors) (see Recipes)
10 mM PP2 (see Recipes)
200 µM PP2 (see Recipes)
VEH-20X (see Recipes)
Recipes
EGM-2 complete medium
The EGMTM-2 Endothelial Cell Growth Medium-2 BulletKitTM contains the following ingredients:
1× EBM-2 basal medium (CC-3156), 500 mL
1× EGM-2 SingleQuots supplement pack (CC-4176) containing:
1× white cap vial with VEGF, 0.50 mL
1× gray cap vial with hFGF-B, 2 mL
1× yellow cap vial with R3-IGF-1, 0.50 mL
1× green cap vial with hEGF, 0.50 mL
1× orange cap vial with heparin, 0.50 mL
1× blue cap vial with ascorbic acid, 0.50 mL
1× red cap vial with GA-1000, 0.50 mL
1× natural cap vial with hydrocortisone, 0.20 mL
1× bottle FBS, 10 mL
Please use the following table to prepare the required volume of EGM-2 complete medium:
EGM-2 complete media (mL) VEGF (µL) FGF (µL) IGF-1 (µL) EGF (µL) Heparin (µL) Ascorbic acid (µL) Gentamycin (GA-1000) (µL) Hydrocortisone (µL) FBS (mL)
50 50 200 50 50 50 50 50 20 1
100 100 400 100 100 100 100 100 40 2
250 250 1,000 250 250 250 250 250 100 5
400 400 1,600 400 400 400 400 400 160 8
500 500 2,000 500 500 500 500 500 200 10
EGM-2 media (with growth factors)
We usually make EGM-2 media (with growth factors) in a bottle and add the FBS to the T-75 flask directly. This is because we often do experiments in serum-free conditions, so we add the FBS separately during cell culture as required.
10 mM PP2
Molecular weight of PP2 = 301.78
Weigh 6 mg of PP2 in a sterile microcentrifuge tube. In the laminar flow hood, dissolve the PP2 in 2 mL of DMSO. Vortex briefly to obtain 10 mM PP2 stock solution. Aliquot this PP2 stock solution (as 50 µL aliquots) into microcentrifuge tubes and store at -20 °C.
200 µM PP2
Thaw out one aliquot of PP2 (concentration = 10 mM) in a water bath (held at 37 °C). Add 2 µL of PP2 in 100 µL of basal EBM-2 medium. Vortex vigorously. Now the concentration of PP2 is 200 µM (called PP2-20X in the text). Use this PP2 solution as described in Section D, Step 8. Discard the remaining solution of PP2-20X.
VEH-20X
Add 2 µL of DMSO in 100 µL of basal EBM-2 medium. Vortex vigorously to obtain VEH-20X solution (the concentration of DMSO in this solution is 2% v/v), which is referred to as VEH-20X. Use the VEH-20X solution as described in Section D, Step 10. Discard the remaining solution of VEH-20X.
Equipment
NU-540 (LabGard® ES NU-540 class II, type A2) laminar-flow biosafety cabinet (NuAire, Plymouth, MN, catalog number: NU-540)
Cell culture incubator (Heracell VIOS 150i cell culture incubator) (Thermo Scientific, Waltham, MA, catalog number: 51-032-872)
Leica DM IL LED inverted phase contrast microscope with camera (VWR International, catalog number: 76382-982)
Fisher Isotemp 220A water bath (Fisher, catalog number: FSGPD05)
Thermo Electron Corporation IEC Centra CL2 benchtop centrifuge (Thermo Scientific, catalog number: 004260F)
Microbalance (Sartorius, Model 1712 MP8, silver edition, catalog number: CUBIS_II_ANALYTICAL)
Vortex mixer (Fisher, catalog number: 02-215-414)
Software
LAS Image Capture Software (Leica Microsystems)
Adobe Photoshop 2023 (Adobe Creative Cloud for Windows)
Adobe Illustrator 2023 (Adobe Creative Cloud for Windows)
NIH ImageJ Version 1.47 (National Institutes of Health, Bethesda)
GraphPad Prism Version 9
Procedure
Culturing HRMECs: thawing HRMECs from freezing vial
Order cryopreserved HRMECs from Cell Systems. These HRMECs are isolated from normal, healthy donor tissue and supplied at passage 3 (<12 cumulative population doublings). A certificate of analysis is provided with each vial, which contains approximately 1 × 106 cells. After obtaining the cryopreserved cells from the vendor, they should be immediately stored in liquid nitrogen. We usually thaw the HRMECs by adding one frozen cell vial into one BioCoat T-25 and one T-75 flask. All steps are done in a laminar flow hood under aseptic conditions.
Rehydration of BioCoat Collagen-coated T-25 and T-75 tissue culture flasks:
The BioCoat flasks are stored at 4 °C. Take the packet containing the flasks out and allow them to come to room temperature prior to use. Warm some bicarbonate-based serum-free culture medium (such as RPMI-1640) in a 37 °C water bath. Add 5 mL of warm bicarbonate-based serum-free culture medium (such as RPMI) to the T-25 flask. Similarly, add 15 mL of warm serum-free media to the T-75 flask. We usually use warm RPMI-1640 for rehydrating the flasks. Allow the flasks to rehydrate for 2 h in a humidified tissue culture incubator maintained at 37 °C with 5% CO2. After 2 h, aspirate the media from the flask, taking care not to disrupt the collagen coating.
Add 10 mL of EGM-2 complete media to the rehydrated T-25 flask. Similarly, add 20 mL of EGM-2 complete media to the rehydrated T-75 flask. Important: Cryopreserved cells are very delicate. The HRMECs are typically cryopreserved in complete EGM-2 media supplemented with 5% DMSO. Thaw the vial in a 37 °C water bath by swirling the vial gently until the contents are completely thawed. Each vial contains approximately 1 × 106 cells at passage 3 (<12 cumulative population doublings).
As soon as the vial is thawed, immediately remove the vial from the water bath, wipe it dry, and rinse it with 70% ethanol. Subsequently, transfer the vial inside a laminar flow hood. Remove the cap, being careful not to touch the interior threads with gloved fingers.
Using a 1 mL sterile serological pipette, gently dispense the contents of the vial into the rehydrated BioCoat T-25 and T-75 flasks containing EGM-2 complete media (Figure 1) at the recommended seeding density of 6,000 cells/cm2. Rinse the freezing vial with 1 mL of EGM-2 complete medium and add it to the tissue culture flasks. Important: Do not centrifuge the cells from the freezing vial. The DMSO present in the cryopreserved cells will get diluted in the EGM-2 complete media present in the flask. Therefore, the cells will not be harmed by the DMSO.
Figure 1. Schematic diagram representing the establishment of human retinal microvascular endothelial cell (HRMEC) cultures from cryopreserved cells. The flasks should be placed flat (horizontally) in the cell culture incubator.
Return the tissue culture flasks to the humidified tissue culture incubator maintained at 37 °C with 5% CO2. Do not disturb the culture for at least 24 h after the culture has been initiated. After 24 h, aspirate and discard the media in the flask and add fresh EGM-2 complete media to remove any residual DMSO and unattached cells. A healthy culture will display cobblestone morphology and non‐granular cytoplasm [9, 10].
Culturing HRMECs: maintenance and subculturing of HRMEC cultures
Culture HRMECs to approximately 70%–80% confluence. The media should be replenished once every three days.
Rehydrate the required number of BioCoat T-75 flasks (see Section A, Step 2).
Rinse the cells with DPBS. Use 5 mL of DPBS for T-25 and 10 mL of DPBS for T-75 flasks.
Add 3 mL of 0.25% Trypsin/EDTA solution into the T-75 flask (in the case of T‐25 flask, use 2 mL).
Gently rock the flask to ensure that the cells are covered by Trypsin/EDTA solution. Incubate the flask at 37 °C incubator for 1–2 min or until cells are completely rounded up (monitored with inverted microscope). Gently tap the side of the flask to fully detach the cells if necessary.
Add 2 mL of FBS to the T-75 flask (use 1 mL for a T-25 flask) to neutralize the Trypsin-EDTA. Using a 5 mL serological pipette, gently rinse the flask completely and transfer the cells in a 15 mL centrifuge tube.
Rinse the growth surface of the flask with 10 mL of DPBS to collect the remainder of the detached cells and combine in the same 15 mL centrifuge tube from Step B6. Close and examine the flask under an inverted microscope to verify that the cell harvesting is complete. There should be less than 5% cells (of the initial density of the flask) remaining in the flask.
Centrifuge the harvested cell suspension at 800× g for 5 min at room temperature using a benchtop centrifuge. Remove (aspirate) the FBS/DPBS liquid from the pellet. Gently flick the tube to dislodge the cells and resuspend them in 1 mL of EGM-2 complete medium.
Count the cells using a hemocytometer and then dispense the required volume of cell suspension into the hydrated T-75 flask as recommended in see Section A, Steps 5–6. Return the flasks back to the cell culture incubator maintained at 37 °C and 5% CO2.
Working with Matrigel
The Matrigel is aliquoted in ice-cold sterile Corning freezing vials and stored at -70 °C.
Before starting the MDA, the required amount of Matrigel should be thawed overnight at 4 °C. The Matrigel is always kept on ice during the experiment.
Matrigel is a liquid at 4 °C, and it polymerizes into a gel-like solid at 37 °C. We recommend that plasticware and reagents should be kept ice cold while handling the Matrigel.
The thawed Matrigel is a thick, viscous liquid. Pipette tips should be trimmed at the tip (to increase the diameter of the tip) to transport the Matrigel in/out of the freezing vials. All microcentrifuge tubes and pipette tips should be kept ice cold while handling the Matrigel to avoid polymerization. In our laboratory, we use ice-cold serum-free RPMI-1640 as a cooling media. All pipette tips are dipped into the ice-cool media to chill them down before using them to handle Matrigel.
Matrigel Duplex Assay
Day 1
The assay is set up in an 8-well chamber slide. Before starting the experiment, draw the schema of the assay (Figure 2). The region (with the checkered pattern) of the right side of the chamber slide is used to lift and transport the slide.
Figure 2. Schema of the Matrigel Duplex Assay
All experiments are performed with HRMECs between passage 3–7. The entire experiment is performed in a laminar flow hood under aseptic conditions.
Grow the HRMECs cells to approximately 70%–80% confluence (in hydrated collagen-coated BioCoat T-75 flasks (please see Section A, Step 2 on instructions for rehydration of flasks) in complete EGM-2 media in a humidified environment at 37 °C (with 5% CO2) in a cell culture incubator.
Wash the HRMECs once with DPBS and trypsinize the flask using 0.25% Trypsin-EDTA.
Add 2 mL of FBS to neutralize the trypsin and transfer the cell suspension to a 15 mL tube. Rinse the flask with DPBS and transfer the solution to a 15 mL tube.
Gently spin down the cells at 800× g for 5 min at room temperature in a benchtop centrifuge. Aspirate the media and resuspend the HRMECs in EGM-2 containing 4% FBS.
Count the cells using the Corning cell counting chamber. Adjust the concentration of the cells to 1.6 × 107 cells/mL (using EGM-2 media containing 4% FBS) in a sterile 5 mL microfuge tube. This tube is labeled as TUBE A.
The stock solution of PP2 (10 mM) should be diluted to a concentration of 200 µM (see Recipes) in basal EBM-2 media in a sterile microfuge tube. The PP2 is prepared at 20× working concentration and labeled as PP2-20X. The PP2-20X is the concentration of PP2 used to set up the MDA. The final concentration of PP2 to be used for the MDA is 10 µM.
In a separate 1.5 mL microfuge tube, mix the cells (from TUBE A) with PP2-20X at the ratio of 10:1 (v/v). This dilutes the PP2-20X to a concentration of 20 µM. As an example, you may add 45 µL of cell suspension (from TUBE A) to 5 µL of the PP2-20X. Gently flick the tube so that the drug mixes well with the cell suspension. Do not vortex because this will shear (and lyse) the cells. Therefore, the new concentration of PP2 is 20 µM. This tube is labeled as TUBE B.
The vehicle for the PP2-20X is 2% DMSO (hereafter referred to as VEH-20X, see Recipes). Just like in the previous step, mix the cells (from TUBE A) with VEH-20X at the ratio of 10:1 (v/v). Do not vortex because this will shear (and lyse) the cells. Therefore, the new concentration of the vehicle is now 0.2% DMSO. This tube is labeled as TUBE B-VEH.
Keep a few empty 1.5 mL microfuge tubes on ice to chill them down. Combine the cells with Matrigel at a ratio of 1:1 in an ice-cold microfuge tube. For example, mix 20 µL of Matrigel with 20 µL of cell suspension (taken from TUBE B). Mix the cells with the Matrigel by flicking the tube. Do not allow the tube to warm up and do not generate air bubbles in the Matrigel–cell suspension while flicking. The final PP2 concentration in the tube is now 10 µM. This tube is labeled as TUBE C.
Similarly, 20 µL of HRMECs (treated with 0.2% DMSO) are mixed with 20 µL of Matrigel in a separate microfuge tube. The final concentration of vehicle in the tube is now 0.1% DMSO. This tube is labeled as TUBE C-VEH.
Table 1 describes the tubes A, B, and C in detail.
Table 1. Description of tubes A, B, and C in the Matrigel Duplex Assay
Tube name Composition of vehicle Contents of tube
A HRMECs resuspended in EGM-2 media supplemented with 4% FBS at a concentration of 1.6 × 107 cells/mL.
B 45 µL of HRMEC suspension (a concentration of 1.6 × 107 cells/ml, taken from TUBE A) mixed with 5 µL of 200 µM PP2 (referred in the text as PP2-20X). Final concentration of PP2 is now 20 µM. Gently flick the tube so that the drug mixes well with the cell suspension.
B-VEH The vehicle for the PP2-20X is 2% DMSO (referred in the text as VEH-20X) 45 µL of HRMEC suspension (a concentration of 1.6 × 107 cells/mL, taken from TUBE A) and mixed with 5 µL of 2% DMSO (which is the vehicle for PP2-20× and also called VEH-20% in the text).
C Mixture of 20 µL of Matrigel with 20 µL of cell suspension (treated with PP2 and taken from TUBE B). Mix the cells with the Matrigel by flicking the tube.
C-VEH TUBE C-VEH contains a mixture of 20 µL of Matrigel with 20 µL of cell suspension (treated with vehicle and taken from TUBE B-VEH). Mix the cells with the Matrigel by flicking the tube.
Take a chamber slide out of the packet. Place the chamber slide in a dish over a piece of tissue paper inside the laminar flow hood. Use a Kimwipe or a sterile gauze pad to prevent the chamber slide from sticking to the surface of the hood, as this may cause a disruption of the primary Matrigel layer during the transfer to the incubator.
Take 6 µL of the cell suspension from TUBE C and plate it as a drop in the center of the chamber slide. Each sample is assayed in duplicate. A similar process is followed for TUBE C-VEH. The chamber slide is then incubated at 37 °C for 1 h in a humidified cell culture incubator with 5% CO2. Below is a schematic representation of the chamber slide with the drop in the middle of the well (Figure 3). We also provide a photograph of the chamber slide with the drop in the middle of the well (Figure 4).
Figure 3. Schematic diagram of the chamber slide with the drop in the middle of the well
Note: The drop does not need to be exactly at the geometric center of the slide. The important fact to remember is that it should be centered enough so that the cells can invade radially into the secondary layer. Even if the drop (the primary layer) is slightly off center, that would be okay for the assay.
Figure 4. Photograph of the chamber slide with the primary layer in the middle of the slide
After 1 h, add 250 µL of EGM-2 complete media to each well. Leave the chamber slide at 37 °C (with 5% CO2) for 24 h in a cell culture incubator.
Thaw a vial of PR-free Matrigel overnight at 4 °C. If the assay is set up as shown in the schematic diagram above (Figure 5), 1 mL of PR-free Matrigel will be required for the experiment.
Figure 5. Schematic diagram of the side view of the chamber slide. (A) The components of the chamber slide have been labeled. (B) The procedure of adding the primary layer (PR-containing Matrigel and HRMECs) in the center of the chamber slide. (C) The drop is allowed to polymerize for 1 h at 37 °C (with 5% CO2) in a cell culture incubator. (D) After 1 h, 250 µL of EGM-2 complete media is added to each well.
Day 2
After 24 h, dilute the PR-free Matrigel with an equal volume of EGM-2 supplemented with 4% FBS. Keep this solution on ice.
Look at the chamber slide under a phase contrast microscope to confirm the presence of a network of retinal endothelial cell tube–like structures within the primary Matrigel layer (the drop).
Aspirate the medium from the chamber slide. Use a pipette to gently remove the medium from the cell without disturbing the drop at the center of the chamber slide.
Gently overlay 200 µL of the 1:1 solution of phenol red-free Matrigel (diluted in EGM-2 supplemented with 4% FBS) over the drop. Figure 6 represents a schematic diagram of the chamber slide. The pink color drop represents the primary layer of Matrigel, and the light grey region (around the pink drop) represents the secondary layer comprised of phenol red–free Matrigel. A side view of the chamber slide is provided in Figure 7. The chamber slide is incubated at 37 °C for 1 h (with 5% CO2) in a cell culture incubator. After 1 h, add 250 µL of EGM-2 complete media to each well. Leave the chamber slide at 37 °C for 24 h in a cell culture incubator.
Figure 6. Schematic diagram of the chamber slide with both the primary and secondary layers
Figure 7. Schematic diagram of the side view of the chamber slide with the primary and secondary layers in the well. (A) Side view of chamber slide with the primary layer (drop in the center) covered with EGM-2 media (light red color). (B) The media is aspirated carefully. (C) The second layer comprising a 1:1 solution of EGM-2 media (containing 4% FBS) and PR-free Matrigel is overlaid on the primary layer. (D) The secondary layer (indicated by grey color) is allowed to polymerize for 1 h at 37 °C in a cell culture incubator. (E) Subsequently, EGM-2 media (indicated in light red color) is added to the well, and the slide is incubated at 37 °C for 24 h (with 5% CO2) in a cell culture incubator.
Day 3
After 24 h, observe the chamber slide under a phase contrast microscope. The angiogenic tube–like structures (from the HRMECs in the primary Matrigel layer) should be clearly seen invading radially into the secondary Matrigel layer. Photograph three independent fields (at 10× magnification) for each sample for quantitative analysis.
Processing the images using Adobe Illustrator and ImageJ
The figure shown below shows the representative images obtained from the MDA. The black arrows indicate the sprouting angiogenic HRMEC tube–like structures, which have invaded the secondary Matrigel layer from the primary layer (Figure 8).
Figure 8. Representative photographs obtained from the Matrigel Duplex Assay (MDA). The black arrows indicate the angiogenic tube–like structures invading into the secondary layer.
Open the images in Adobe Illustrator 2023. Using the pen curvature tool, draw a dotted line across the interface between the two layers. A representative image with the dotted lines is shown below (Figure 9).
Figure 9. Drawing the dotted line at the interface of the two layers using Adobe Illustrator 2023
Export the images to Adobe Photoshop 2023 and save them in jpeg format. Open the jpeg files in ImageJ. Using the line tool, draw a set of six lines marking the distance that the angiogenic tube–like structures have invaded into the secondary Matrigel layer (Figure 10). After drawing each line, press CTRL-M to obtain the length of the line drawn. Cease drawing the line upon encountering a gap larger than two cells. Our microscope images were analyzed using the ImageJ program by three independent observers in a double-blind manner. The raw data is provided in Supplementary information.
Figure 10. Representative image of the Matrigel Duplex Assay (MDA) with lines drawn using Adobe Illustrator
Open GraphPad Prism (Version 9). Using the numbers obtained by the ImageJ program, create a column graph of the data and perform statistical analysis (Figure 11). The statistical analysis of the data is also described in the Data analysis section.
Figure 11. Microscope images were analyzed by the ImageJ program (by three independent observers). Data were graphically represented using GraphPad Prism (Version 9). Data were analyzed through an unpaired non-parametric t-test followed by the Mann-Whitney test. Values represented by the same letter are not statistically significantly different from each other (P ≤ 0.05).
Data analysis
Each sample was tested in duplicate in the MDA. The entire assay was repeated four independent times. The assay was quantified using phase contrast microscopy to obtain images for analysis. Three representative images (at 10× magnification) were captured for each sample. These images were quantified by Adobe Illustrator 2023 and Adobe Photoshop 2023, followed by ImageJ analysis by three independent observers in a randomized double-blind manner. The data is graphed using GraphPad Prism (Version 9). All data are plotted as mean ± standard deviation. Data were analyzed through an unpaired non-parametric t-test followed by the Mann-Whitney test. All analyses should be completed using a 95% confidence interval. Data is considered significant when P < 0.05.
Note: Our laboratory uses Adobe Illustrator and Adobe Photoshop combined with ImageJ to perform the analysis of data obtained in the MDA. We realize that the Adobe Illustrator and Adobe Photoshop software are not readily available to all laboratories. Alternative programs for data analysis include open-source image processing programs like ImageJ or Fiji. These platforms offer a vast array of tools and macros, such as the Drawing Tools set available at this GitHub location: https://github.com/fiji/fiji/blob/master/macros/toolsets/Drawing%20Tools.txt. Such open-source alternatives would broaden the protocol's accessibility, benefiting a larger number of users. However, we have no experience in using these open-source software programs, so we cannot comment on their efficacy for data analysis.
Validation of protocol
This MDA protocol was used in the following research papers:
Dom, A.M., Buckley, A.W., Brown, K.C., Egleton, R.D., Marcelo, A.J., Proper, N.A., Weller, D.E., Shah, Y.H., Lau, J.K., Dasgupta, P. (2011) The α7-nicotinic acetylcholine receptor and MMP-2/-9 pathway mediate the proangiogenic effect of nicotine in human retinal endothelial cells. Invest. Ophthalmol. Vis. Sci., 52, 4428-4438.
General notes and troubleshooting
Troubleshooting tips
HRMECs should be cultured to approximately 70%–80% confluence. Replace culture medium at least once in three days. Avoid growing the cells to more than 80% confluence to prevent contact inhibition and senescence of cultures.
It is very important that the HRMECs are healthy for the assay. Cells should display a cobblestone morphology and non‐granular cytoplasm. Therefore, we usually use HRMECs between passage 3 and 7 for all our experiments.
The HRMECs should be carefully trypsinized. Excessive trypsinization (or insufficient neutralization of trypsin) promotes cell aggregation and cell death and releases cellular debris, which will compromise the quality of the assay.
The Matrigel aliquot should be thawed overnight at 4 °C. Alternately, the Matrigel aliquot may be thawed on ice (for 5–6 h) before performing the experiment. Avoid rapid thawing of the Matrigel.
When working with Matrigel, use pre-chilled pipette tips. Alternatively, fill a 15 mL centrifuge tube with ice-cold sterile PBS or base media to cool the tip by filling and ejecting the cold liquid several times immediately before use with the Matrigel.
It is very important to aliquot the Matrigel. Do not re-freeze Matrigel more than twice. Repeated freeze and thaw of the Matrigel stock or aliquots will degrade the basement membrane proteins within the Matrigel. Similarly, storage of Matrigel at room temperature will cause it to polymerize (solidify), rendering it unusable.
Keep the mix of HRMECs and Matrigel (for the primary layer) on ice to prevent the solidification of Matrigel. Make sure to pipette up and down to keep the cells uniformly suspended before plating. Do not vortex the mixture of HRMECs and Matrigel. Perform gentle pipetting to avoid forming any bubbles in the Matrigel during the process.
The secondary layer comprises a 1:1 solution of phenol red–free Matrigel diluted in EGM-2 supplemented with 4% FBS. Mix the phenol red–free Matrigel and EGM-2 (supplemented with 4% FBS) by gently pipetting up and down. Avoid agitating the tubes or vortexing them as this will create air bubbles in the secondary layer.
Acknowledgments
We acknowledge Dr. S. Chellappan and his laboratory for their continuous support. PD and MAV are supported by a National Institutes of Health R15 Academic Research Enhancement Award (2R15CA161491-03). MAV is supported by NIH R15AI15197-01 and R15HL145573-01. This work was supported in part by the West Virginia IDeA Network of Biomedical Research Excellence (WV-INBRE) grant (NIH grant P20GM103434; PI: Dr. G. Rankin).
Competing interests
The authors declare no competing interests.
References
Usui, Y., Westenskow, P. D., Murinello, S., Dorrell, M. I., Scheppke, L., Bucher, F., Sakimoto, S., Paris, L. P., Aguilar, E., Friedlander, M., et al. (2015). Angiogenesis and Eye Disease. nnu. Rev. Vision Sci. (1): 55–184.
Dreyfuss, J. L., Giordano, R. J., Regatieri, C. V. (2015). Ocular Angiogenesis. J Ophthalmol. doi: 10.1155/2015/892043.
Eelen, G., Treps, L., Li, X. and Carmeliet, P. 2020). Basic and Therapeutic Aspects of Angiogenesis Updated. irc. Res. 27(2): 10–329.
Arnaoutova, I., George, J., Kleinman, H. K. and Benton, G. 2009). The endothelial cell tube formation assay on basement membrane turns 20: state of the science and the art. Angiogenesis 12(3): 267–274.
Stryker, Z. I., Rajabi, M., Davis, P. J. and Mousa, S. A. (2019). Evaluation of Angiogenesis Assays. Biomedicines 7(2): 37.
Chan-Ling, T. (2008). Vasculogenesis and Angiogenesis in Formation of the Human Retinal Vasculature. In: Penn, J. (eds) Retinal and Choroidal Angiogenesis. Springer, Dordrecht. https://doi.org/10.1007/978-1-4020-6780-8_6.
Moleiro, A. F., Conceição, G., Leite-Moreira, A. F. and Rocha-Sousa, A. (2017). A Critical Analysis of the AvailableIn VitroandEx VivoMethods to Study Retinal Angiogenesis. J Ophthalmol 2017: 1–19.
Rezzola, S., Belleri, M., Gariano, G., Ribatti, D., Costagliola, C., Semeraro, F. and Presta, M. (2014). In vitro and ex vivo retina angiogenesis assays. Angiogenesis 17(3): 429–442.
Browning, A. C., Dua, H. S. and Amoaku, W. M. (2008). The effects of growth factors on the proliferation and in vitro angiogenesis of human macular inner choroidal endothelial cells. Br J Ophthalmol 92(7): 1003–1008.
Stitt, A. W., McGoldrick, C., Rice-McCaldin, A., McCance, D. R., Glenn, J. V., Hsu, D. K., Liu, F. T., Thorpe, S. R. and Gardiner, T. A. (2005). Impaired retinal angiogenesis in diabetes: role of advanced glycation end products and galectin-3. Diabetes 54(3):785-94.
Dom, A. M., Buckley, A. W., Brown, K. C., Egleton, R. D., Marcelo, A. J., Proper, N. A., Weller, D. E., Shah, Y. H., Lau, J. K., Dasgupta, P., et al. (2011). The α7-nicotinic Acetylcholine Receptor and MMP-2/-9 Pathway Mediate the Proangiogenic Effect of Nicotine in Human Retinal Endothelial Cells. Investigative Opthalmology & Visual Science 52(7): 4428.
Bain, J., McLauchlan, H., Elliott, M. and Cohen, P. (2003). The specificities of protein kinase inhibitors: an update. Biochem. J. 371(1): 199–204.
Supplementary information
Raw data for the MDA obtained from ImageJ(CTRL-M) function
Each sample was tested in duplicate in the MDA. The entire assay was repeated four independent times. The assay was quantified using phase contrast microscopy to obtain images for analysis. Three representative images(at 10× magnification) were captured for each sample. These images were quantified by Adobe Illustrator 2023 and Adobe Photoshop 2023 followed by ImageJ analysis by three independent observers in a randomized double-blind manner.
The terms Control-1, Control-2, and Control-3(in EXCELOBSERVER-1-MDA-COUNTS-RAW-DATA, indicated by the green colored rows R1, R2, R3) represent three images(of three independent fields) of the vehicle-treated HRMECs. Similarly, PP2-10 µM-1, PP2-10 µM-2, and PP2-10 µM-3, indicated by orange-colored rows R4, R5, R6(EXCELOBSERVER-1-MDA-COUNTS-RAW-DATA), represent three images(of three independent fields) of PP2-treated HRMECs. A similar annotation has been used for EXCELOBSERVER-2-MDA-COUNTS-RAW-DATA and EXCELOBSERVER-3-MDA-COUNTS-RAW-DATA.
In the file EXCELOBSERVER-1-MDA-COUNTS-RAW-DATA, columns C1–C6 represent the dimensions of the six lines drawn for each image(analyzed by ImageJ followed by CRTL-M), as described by OBSERVER-1. The column C7 marked as“AVERAGE”(and indicated in yellow) represents the average of each row. The numbers in column C9(in pink color) show the average from the first three numbers(rows R1, R2, R3) in column C7. This number is the average of all vehicle-treated control HRMECs(indicated by pink color and referred as VEH in column C8) for Experiment-1. This process was repeated for Experiments 2-4. A similar annotation has been used for EXCELOBSERVER-2-MDA-COUNTS-RAW-DATA and EXCELOBSERVER-3-MDA-COUNTS-RAW-DATA.
The numbers in column C8 and row R5(in blue color) show the average of numbers in the last three rows(R4, R5, R6) of the column C7. This number is the average of the PP2-treated HRMECs(indicated by blue color and referred as PP2-Av in column C8) for Experiment-1. This process was repeated for Experiments 2-4. A similar annotation has been used for EXCELOBSERVER-2-MDA-COUNTS-RAW-DATA and EXCELOBSERVER-3-MDA-COUNTS-RAW-DATA.
The above-mentioned process yielded 12 values(from three observers over four experiments) for vehicle-treated control HRMECs, which are represented in Column A(yellow column) in the file EXCELDATA-PROCESSING-FROM-THREE-OBSERVERS. Similarly, the 12 values for PP2-treated HRMECs are shown in Column B(red column) in the file EXCELDATA-PROCESSING-FROM-THREE-OBSERVERS. The average of the numbers for vehicle-treated control(Column A) was calculated(= 595.5). Using this value, data was converted into percentage of control(Table 4, Columns C and D). The numbers in columns C and D were used to generate the graph(Figure 11) in GraphPad Prism.
The following supporting information can be downloaded here:
EXCELSUPPL-DATA-OBSERVER-1-MDA-COUNTS-RAW-DATA: Raw data obtained by first observer.
EXCELSUPPL-DATA-OBSERVER-2-MDA-COUNTS-RAW-DATA: Raw data obtained by second observer.
EXCELSUPPL-DATA-OBSERVER-3-MDA-COUNTS-RAW-DATA: Raw data obtained by third observer.
EXCEL SUPPL-DATA-PROCESSING-RESULTS-FROM-THREE-OBSERVERS: Processing the raw data from the observers to create the graph(Figure 11) using GraphPad Prism.
Article Information
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© 2023 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Developmental Biology > Cell growth and fate > Angiogenesis
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Streamlined Methods for Processing and Cryopreservation of Cell Therapy Products Using Automated Systems
YL Ye Li *
Hazel Y. Stevens *
SS Srikanth Sivaraman
LP Logan N. Porter
AH Allison R. Hoffman
SG Stuart L. Gibb
SS Shivaram Selvam
AB Annie C. Bowles-Welch
(*contributed equally to this work)
Published: Vol 13, Iss 24, Dec 20, 2023
DOI: 10.21769/BioProtoc.4900 Views: 1827
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Abstract
Streamlined procedures for processing and cryopreservation of cell therapies using good laboratory practices are integral to biomanufacturing process development and clinical applications. The protocol herein begins with the preparation of human cell types cultured as adherent (i.e., mesenchymal stromal cells, MSCs) or suspension cells (i.e., peripheral blood mononuclear cells, PBMCs) to comprehensively demonstrate procedures that are applicable to commonly used primary cell cultures. Cell processing steps consist of preparing high yields of cells for cryopreservation using instruments routinely used in cell manufacturing, including the Finia® Fill and Finish System and a controlled-rate freezer. The final steps comprise the storage of cells at subzero temperatures in liquid nitrogen vapor phase followed by the analysis of cell phenotypes before and after processing and cryopreservation, along with cell quality metrics for validation. Additionally, the protocol includes important considerations for the implementation of quality control measures for equipment operation and cell handling, as well as Good Laboratory Practices for cell manufacturing, which are essential for the translational use of cell therapies.
Key features
• The protocol applies to small- or large-scale manufacturing of cell therapy products.
• It includes streamlined procedures for processing and cryopreservation of cells cultured as adherent cells (MSCs) and suspension cells (PBMCs).
• Provides temperature control and rapid partitioning of sample in cryopreservation solution to maintain high viability of a range of cell types throughout the procedures.
• This protocol employs the Finia® Fill and Finish System and a controlled-rate freezer.
Graphical overview
Keywords: Cell therapies Cryopreservation Biomanufacturing Process development Mesenchymal stromal cells Peripheral blood mononuclear cells Automated systems Finia® Fill and Finish System
Background
Cell therapy investigations rely on ex vivo operations to adequately preserve the viability and functionality of the cell products prior to in vivo administration (Li et al., 2019). Process development for cell products requires standard operations with controlled systems, which should be monitored throughout the process of end-to-end manufacturing. Clinical laboratories routinely cryopreserve cells using cryopreservation solutions and operating conditions that need to be tightly controlled and amenable to the cell type for optimal cell viability and yield. The use of automated equipment that combines multiple manufacturing processes, such as fill-finish systems and controlled-rate freezers, can offer several advantages over manual operations including automation, better control of processing, and efficiency, thereby avoiding potential compromises to the viability and yield of cell products (van der Walle et al., 2021). Together, these systems can be employed and programmed to optimize the operations for cell cryopreservation at end-stage manufacturing as well as starting materials upstream of manufacturing.
Multiple cell types have been evaluated as cell therapies for clinical use to treat a broad spectrum of clinical indications. Depending on the cell type, specific reagents and operating conditions need to be considered for ex vivo manufacturing expansions to generate high yields for therapeutic dosages (Galipeau and Sensébé, 2018; Pigeau et al., 2018). For example, anchorage-dependent cells require adherence to substrates (e.g., plastic cultureware, microcarriers) to proliferate, whereas other cells proliferate in suspension without substrates (Merten, 2015; Cuesta-Gomez et al., 2023). Typically, cell therapy products are initially isolated from tissues or collected by apheresis from individuals and subsequently undergo various process development procedures to isolate, expand (i.e., proliferate), and harvest the cells (Bunnell et al., 2008; Palen et al., 2021). Herein, procedures that are applicable for both adherent (i.e., mesenchymal stromal cells, MSCs) and apheresis-derived suspension cells (i.e., peripheral blood mononuclear cells, PBMCs) were described to demonstrate how to prepare the cells (e.g., culture and harvest MSCs, isolated PBMCs) and subsequently execute automated and controlled process development procedures that are amenable to cell therapy product manufacturing.
The Finia® Fill and Finish System (Terumo Blood and Cell Technologies) is an automated, closed system that offers temperature-controlled processing for formulation and aliquoting of liquids, including cell suspensions, in preparation for cryopreservation (Sethi and Cunningham, 2021; Algorri et al., 2022). The system mixes and cools up to three materials (e.g., cells, buffers, cryopreservation solutions) to a specified temperature. The system offers flexible procedure programming to allow the user to define workstream needs, from simple aliquoting of a cell source to the stepwise cooling of cell solution, a relevant buffer, and a cryopreservation solution of choice. The final cell product is then aliquoted into individual product bags and automatically sealed. The single-use disposable set is composed of product bags appropriate for freezing, thawing, and administration in the clinic, and a quality control bag for testing (Sethi and Cunningham, 2020; Cunningham et al., 2022). The disposable set is offered in two volume configurations, allowing for the freezing of volumes from 10 to 70 mL per bag. The Finia system is governed by the cell processing application (CPA), which is a secure server-based software (i.e., protected from access by unauthorized individuals) for procedure management and record keeping, capable of tracking performance and protocol development for current Good Manufacturing Practices (cGMP) compliance.
The Finia system and controlled-rate freezer are two separate automated systems that together provide a closed system workflow, which has several advantages over common cell culture practices, including a reproducible cryopreservation procedure with >90% post-thaw cell viability and the prevention of risks related to contamination and operator error. In addition, product bags can store considerably more cells than cryovials (Sethi and Cunningham, 2021). Sethi and Cunningham performed studies to compare a robust manual process to the Finia system (automated process) using T cells and found that the targeted product volumes were more accurate using the automated processing, and cell viability (comparing pre-formulation, post-formulation, and post-thaw) was comparable between the two processes (Sethi and Cunningham, 2021). Cell therapy manufacturing can be further automated by the use of programmable, multi-step-controlled rate freezers, which can standardize and record freezing procedures (Meneghel et al., 2020). Based on these previous studies, our group has implemented streamlined methods for using the Finia system and a controlled-rate freezer for processing and cryopreservation of both adherent and suspension cell types commonly used in cell manufacturing as cell therapy products or as starting materials, respectively. Terumo Blood and Cell Technologies has performed internal testing on the Finia system to demonstrate it can be used for both adherent and suspension cells (data on file). Furthermore, the use of the Finia system in a workstream for a homogenous population of T cells was recently reported (Cunningham et al., 2022). This is the first published report on the use of the Finia system with adherent cells (i.e., MSCs) as well as heterogeneous suspension cells (i.e., PBMCs). Furthermore, we provide quality control results (based on cell count, viability, and phenotyping), troubleshooting guidance, and considerations for GMP and Good Laboratory Practice (GLP) compliance with application to automated and controlled process development procedures amenable to cell therapy product manufacturing.
Materials and reagents
Biological materials
Human MSCs derived from umbilical cord tissue cryovial, stored LN2 vapor phase (supplied by Duke University MC3)
Human peripheral blood one-tenth leukopak fresh (STEMCELL Technologies, 200-0092)
Reagents
Prime-XV MSC Expansion (XSFM) (Irvine Scientific, catalog number: 91149)
Penicillin/streptomycin (P/S) (Millipore Sigma, catalog number: P4333)
Phosphate buffered saline (PBS) Ca2+, Mg2+ free (Millipore Sigma, catalog number: TMS-012-A)
PLTGold® human platelet lysate (hPL) (Millipore Sigma, catalog number: SCM151;)
TrypLE Express 1× (Millipore Sigma, catalog number: 12605028)
Cryostor® CS-10 (Fisher Scientific, catalog number: NC9930384)
LymphoprepTM (STEMCELL Technologies, catalog number: 07801)
Fetal bovine serum (FBS) (any vendor)
Zombie UV Fixable Viability kit (BioLegend®, catalog number: 423107)
Human TruStain FcXTM (BioLegend®, catalog number: 422302)
BD CytofixTM fixation buffer (BD Biosciences, catalog number: 554655)
Solutions
Dilution buffer (see Recipes)
FC buffer (see Recipes)
Viability staining solution (see Recipes)
Fc block solution (see Recipes)
Recipes
Dilution buffer
Reagent Final concentration Quantity
PBS Ca2+/Mg2+ free (1×) 98% 98 mL
hPL 2% 2 mL
Total 100% 100 mL
FC buffer
Reagent Final concentration Quantity
PBS Ca2+/Mg2+ free (1×) 98% 98 mL
FBS 2% 2 mL
Total 100% 100 mL
Viability staining solution
Reagent Final concentration Quantity
PBS Ca2+/Mg2+ free (1×) 99.999% 9.99 mL
Zombie UV dye (reconstituted) 0.001% 10 μL
Total 100% 10 mL
Fc block solution
Reagent Final concentration Quantity
FC buffer 90% 9 mL
Human TruStain FcX 10% 1 mL
Total 100% 10 mL
Laboratory supplies
CellBIND® surface HYPERFlask® cell culture vessel (Corning®, catalog number: 10020)
50 mL conical tubes (any vendor)
Storage bottle, sterile (any vendor)
Serological pipettes and tips, assorted volumes (any vendor)
Micropipettes and tips, P10–P1000 μL (any vendor)
T-150 transfer bag (Fisher Scientific, catalog number: NC0470433)
FINIA 50 tubing set (Terumo Blood and Cell Technologies, catalog number: 22050)
Note: The tubing set comprises a mixing bag, a QC bag (10–40 mL), and three storage bags (10–84 mL total).
FINIA 250 tubing set (Terumo Blood and Cell Technologies; catalog number: 22250)
Note: The tubing set comprises a mixing bag, a QC bag (10–40 mL), and three storage bags (29–210 mL total).
Vapor phase storage cryovials for cell samples including QC (any vendor)
Microcentrifuge tubes low binding (any vendor)
Via-1-CassetteTM cartridges (Chemometec, catalog number: 941-0012)
Syringe 10 mL with luer lok (any vendor)
Blunt fill needle 18-gauge length 1.5 inch (any vendor)
Controlled rate freezer canister rack adjustable (Thermo Fisher Scientific, catalog number: 11679084)
Stainless steel freezing canister with bag capacity 250 mL (Thermo Fisher Scientific, catalog number: 4000335)
Frame to hold canisters in LN2 storage (Thermo Fisher Scientific, catalog number: 4R5461)
Tank of liquid nitrogen (any vendor)
96-well plates (any vendor)
Equipment
Biosafety cabinet (BSC) level A2 (e.g., Thermo Fisher Scientific, catalog number: 1377)
Incubator set to 5% CO2 and 37 °C (e.g., HeraCell VIOS; Thermo Fisher Scientific)
Cell counter (e.g., Chemometec, model: NucleoCounter NC-200TM)
Note: Cell counting methods can vary. It is important to use a reliable and reproducible method for acquiring cell counts and diameter.
Refrigerated centrifuge capable of reaching 800 rcf/g and using 50 mL conical tubes (any vendor)
Micropipette P10–1000 μL (any vendor)
Pipette controller (any vendor)
Inverted microscope (e.g., Thermo Fisher Scientific, model: EVOS M5000)
Note: Microscope needs to be used for phase contrast imaging of cells only.
Finia Fill and Finish System (Terumo Blood and Cell Technologies)
Sterile tubing welder (Terumo Blood and Cell Technologies, model: TSCD-II)
Tubing sealer (Terumo Blood and Cell Technologies, model: T-SEAL Mobile)
Flow cytometer, e.g., CytoFlex (Beckman Coulter), LSRFortessa (BD)
Liquid nitrogen (LN2) dewar or cryo unit (any vendor)
Controlled rate freezer (e.g., Thermo Fisher Scientific, CryoMed, catalog number: 7454)
Note: Freezer needs to be programmable to -90 °C.
Software and datasets
Cell Processing Application (CPA) software version 2.1 with Finia Fill and Finish System
Flow cytometry software for data analysis, e.g., FlowJoTM software (BD Biosciences), CytoBank (Beckman Coulter)
Procedure
All cell handling procedures should be performed in a BSC. The Finia system does not require the use of a BSC. The following procedures outline cell preparation practices for MSCs and PBMCs as the starting material for subsequent processing and cryopreservation using automated systems. More specifically, the procedures include thawing, cell culture, and harvest of cryopreserved MSCs as well as isolation of PBMCs from a fresh (i.e., not cryopreserved) one-tenth leukopak.
Note: It is not advisable to follow the procedures for processing and cryopreservation of PBMCs directly thawed and isolated from a cryopreserved leukopak. If a cryopreserved leukopak is used, allow for a 24–48 h culture rescue period prior to processing and cryopreservation.
Cell thawing (cryopreserved cells, MSCs)
Thaw XSFM media at 4 °C overnight prior to the day of use.
Prepare fresh media by adding 1% P/S to XSFM.
Add 8 mL of fresh media to a 50 mL conical tube.
Retrieve a vial of frozen cells from the LN2 storage.
Thaw the vial in a water bath set at 37 °C for 2 min or until 90% of the ice has melted.
Note: This may depend on the size of the vial and content volume.
Record the thawing time.
Gently transfer the cells from the cryovial to a 50 mL conical tube using a 1,000 μL micropipette with tip.
Wash the vial with 1 mL of fresh media and add this to the conical tube. The final volume of the thawing media and cells should be approximately 10 mL.
Gently resuspend the cells with a 10 mL serological pipette.
Perform a cell count #1 (“Thawed Pre-Spin Count”) by adding 100 μL of cell suspension to a microcentrifuge tube.
Insert the Via-1-Cassette tip into the microcentrifuge tube to uptake approximately 60 μL of cell suspension.
Load the cassette into the NucleoCounter and record the cell count and diameter.
Note: Please complete reading within 1 min after loading cells into cassette.
Centrifuge the remaining cell suspension at 500× g for 5 min at 4 °C.
Aspirate the supernatant using a 10 mL serological pipette and add enough media to give an approximate cell concentration between 1 and 2.5 × 106 cells/mL (optimal concentration for accurate determination of cell count).
Thoroughly resuspend the cells.
Perform and record cell count #2 (“Thawed Post-Spin Count”) by adding 100 μL of cell suspension to a microcentrifuge tube.
Note: Measurement of cells post spin can give an adjusted measure of viability due to dead cells being eliminated in the spin and can also be more accurate as the cells are usually less dilute.
Insert the Via-1-Cassette tip to uptake approximately 60 μL of cell suspension.
Load into the NucleoCounter and record the cell count and diameter.
Calculate the volume needed of cell suspension to give 1.72 × 106 cells (number of cells needed for each HYPERFlask).
Add 560 mL of media (XSFM + 1% P/S) into a storage bottle. Add 1.72 × 106 cells (1,000 cells/cm2) to the storage bottle and rock to ensure that the cells are evenly distributed in media.
Gently pour the cell suspension into the HYPERFlask tilted at a 30° angle to the horizontal.
Place the cap and distribute the cell suspension among all layers of the flask by placing on its long side and then its short side; then, place it lying flat and repeat these movements for even distribution of cells on all layers.
Ensure that there are no air bubbles in the body of the flask by adding more media if needed. Small bubbles are acceptable near the neck of the flask.
Incubate for five to seven days in an incubator set at 5% CO2 and 37 °C.
Cell expansion and harvest
Take daily microscopy images of the lowest layer of the flask to gauge confluence. Do not allow the MSCs to become overcrowded. Harvest MSCs when 80% confluence is achieved.
Carefully pour 50 mL of media from the flask into a 50 mL conical tube. Label and set aside. This is used later to wash the flask.
Remove all remaining media from the flask by pouring gently into a waste container.
Add 50 mL of PBS 1× Ca2+, Mg2+ free to the HYPERFlask and tilt the flask on its long end, short end, and flat side multiple times for an even distribution of the reagent.
Remove the PBS by pouring it gently into a waste container.
Add 50 mL of TrypLE 1× to the flask and distribute evenly by rotating the flask along each end to detach cells.
Incubate the flask for 5 min at 37 °C.
Check the cells under a microscope. Tap the flask lightly against the microscope base to dislodge cells. If incomplete detachment of the cells from the flask is observed, re-incubate for at least another 5 min.
Once the cells are completely detached, add 50 mL of XSFM to the flask and distribute evenly by rotating the flask along each end to subdue the activity of TrypLE.
Note: Unlike TrypLE, other dissociation reagents may require neutralization with media containing serum.
Gently pour the contents of the flask (i.e., cells in solution) into two 50 mL tubes.
Centrifuge the tubes at 500× g for 10 min at 4 °C.
Carefully remove the supernatant from the tubes and combine the pellets into a single tube by adding 5 mL of fresh media to one of the tubes, resuspending the cells, and transferring the cells in suspension to the second tube with the cell pellet.
Thoroughly resuspend the cells.
Perform and record cell count #3 (“Harvested Cell Count”) by adding 10 μL of cell suspension and 90 μL of XSFM to a microcentrifuge tube.
Note: Given an expected high number of cells, it is recommended to perform cell counting with a dilution of the cell suspension to obtain an accurate cell count that is within the range of the instrument’s capabilities. Make sure to factor the dilution into the final cell count.
Uptake the diluted sample into a Via-1-Cassette and load it into the NucleoCounter.
Record the cell count and diameter.
Keep the cells in media at room temperature (RT) until the Finia system is ready.
Note: It is advisable to prepare the cells once the Finia system has been prepared for processing to minimize the time cells are kept at RT.
Cell preparation (fresh cells, PBMCs)
Bring PBS Ca2+, Mg2+ free, hPL, and Lymphoprep to RT.
Bring the centrifuge to RT by switching off the cooling function and remove the brake by setting the deceleration to 0.
Note: Removing the brake is necessary for density separation.
Prepare 100 mL of dilution buffer (see Recipes).
Transfer the contents of the one-tenth leukopak into a 50 mL conical tube.
Note: The volume is between 10 and 30 mL for one-tenth leukopak.
Record the total volume and add 2× volume of dilution buffer (i.e., 7 mL leukopak + 14 mL dilution buffer = 21 mL total volume).
Perform and record cell count #1 (“Pre-Separation Cell Count”) by adding 10 μL of cell suspension and 90 μL of thawing buffer to a microcentrifuge tube.
Note: Given an expected high number of cells, it is recommended to perform cell counting with a dilution of the cell suspension to obtain an accurate cell count that is within the range of the instrument’s capabilities. Make sure to factor the dilution into the final cell count.
Prepare a 50 mL tube with an equal volume of Lymphoprep to the total volume of diluted blood preparation.
Carefully layer the diluted blood preparation over the Lymphoprep by holding the tube at a 45° angle and slowly adding the blood approximately an inch above the Lymphoprep using a 1,000 μL micropipette with tip (see Figure 1A).
Figure 1. Images of leukopak processing before (A) and after separation (B) using Lymphoprep by density centrifugation to isolate peripheral blood mononuclear cells (PBMCs)
Centrifuge the tube at 800× g for 20 min with the brake off and temperature set to RT.
Carefully remove and discard the upper plasma layer.
Collect the buffy coat containing the PBMCs in a 50 mL tube (see Figure 1B).
Add 10 mL of dilution buffer to the tube to wash the PBMCs.
Centrifuge at 300× g for 5 min at 4 °C with brake on by setting deceleration to 9.
Transfer the supernatant to a new tube and add 10 mL of dilution buffer to wash.
Retain and set aside the cell pellet.
Centrifuge the tube with supernatant at 300× g for 5 min at 4 °C to retrieve the remaining cells.
Remove the supernatant and combine the cell pellets from both centrifugations into a single one adding 5 mL to 10 mL of dilution buffer to one of the tubes, resuspending the cells, and transferring the cells in suspension to the second tube with cell pellet.
Perform count #2 (“Post-Separation Cell Count”). Take a 10 μL sample of the cell suspension and dilute 1:10 in a microcentrifuge tube.
Uptake the diluted sample into Via-1-Cassette and load it into the NucleoCounter.
Record the cell count and diameter.
Keep the cells in dilution buffer at RT until the Finia system is ready.
Preparation of the Finia system
Log into the Cell Processing Application (CPA) software, and under Configure Devices (see Figure 2), create a custom protocol to set up the material, product bag, and QC bag volume targets and temperature set point. Save the protocol.
Notes:
The temperature selected for MSCs and PBMCs was 8 °C. Temperature settings will affect the total process time. For example, setting the temperature of the protocol to 8 °C will reduce the time compared to setting to 4 °C.
For the FINIA 50 tubing set, the product bags hold 10–28 mL and the QC bag holds 10–40 mL, so the maximum product volume is 124 mL + 6 mL (retained in tubing). For the FINIA 250 tubing set, the product bags hold 29–70 mL, the QC bag holds 10–40 mL, and the maximum product volume is 250 mL + 6 mL (retained in tubing), including the QC volume.
Figure 2. Image of the Cell Processing Application (CPA) in the Finia system
Click Configure Materials and define material IDs and specific gravity of each material. Touch Save.
Create barcodes for each material.
Note: It is important to ensure the material ID on the barcode matches the material ID in the CPA. Barcodes are created using the following schema: for Material 1 (cells): [Material ID]|[Material Batch ID]|Batch ID, e.g., MSCs|20230207|BatchID. For Material 2 (solution): [Material Type]|[Lot_ID], e.g., CS10|Lot 123. Remember to create a unique batch ID for each experiment to be able to recover the data stored in the CPA. “BatchID” is used by the system to identify that Material batch ID as the batch ID for the procedure report in the CPA.
Optionally, mark the sterile PVC section of the transfer bag every 3 inches from its opening with a marker to aid in welding of the transfer bag to the tubing set.
Note: To ensure you have sufficient length of tubing, tube welding at intervals of 3 inches is recommended.
Turn on the welder.
Note: It will take 5 min for the welder to warm up.
Turn on the Finia device and log in to the system. Touch Start Procedure to advance to the protocol selection screen.
Note: If the device was previously on, turn the device off for 60 s and then power it back on. The device receives configuration and material updates from the CPA each time it is powered on and the user logs in.
The list of custom protocols is displayed. Touch the protocol that was created to use for the procedure and then touch Confirm.
Check the function of the load sensor by first checking the 0-gram setting. With no weight on the load sensor, touch Check 0 g.
Place the 500 g test weight on the load sensor. Once the test weight is at rest, touch Check 500 g.
Remove the test weight from the load sensor and close the doors.
The custom protocol values for material, volume, and QC bag targets and temperature set point appear on the screen. Visually confirm the values and touch Confirm.
Notes:
This step refers to a protocol generated by following step D1.
To change a target or the temperature set point, touch any field on the screen and enter the new values. Touch Next to advance to the next screen. Continue until the procedure targets are correct and touch Confirm.
Scan the barcode of the tubing set and place the mixing bag on the load sensor.
Note: Wipe the plate behind the mixing bag to remove any condensation that may cause the mixing bag to stick to the plate.
Connect all tubing via the pumps, tubing holders, and fluid sensor, and close the pump covers.
Slide the product bags and QC bag into the bag holders and load the bag lines into the valves. Pull the bag lines up through the valve until the port touches the bottom of the slide holder (see Figure 3).
Figure 3. The Finia Fill and Finish System. A. The Finia system with general user interface and barcode scanner underneath the equipment manifold. B. Finia system setup with FINIA 50 tubing set.
Separation of the cells into bags and starting the Finia procedure
Use the Harvested Cell Count (MSCs) or Post-Separation Cell Count (PBMCs) to calculate the amount of CryoStor CS-10 required.
Note: For MSCs, the total volume placed into the transfer bag was 30 mL (10 mL product bag + 10 mL QC bag + 6 mL tubing + 15% extra). For PBMCs, the total volume was 41 mL (20 mL product bag + 10 mL QC bag + 6 mL tubing + 15% extra.) Confirm that the transfer bag (materials) contains enough volume for the Finia system to prepare in the mixing bag.
Print cryolabels with donor information, cell concentration, operator initials, date, and any additional information desired (e.g., PDL/passage #).
Retrieve the tube containing the cells from step B17 or C21.
Centrifuge the cells to obtain the cell pellet.
Note: Centrifugation for MSCs was 500× g for 5 min and for PBMCs was 300× g for 5 min.
Remove the supernatant without disturbing the cell pellet.
Add an appropriate amount of Cryostor CS-10 to the cell pellet at a desired cell concentration.
Notes:
Record the duration for which the cells were exposed to the cryopreservation solution. Perform the following procedures quickly to minimize the duration for which cells are in cryopreservation solution, because cryopreservation solutions contain chemicals that may compromise cell viability.
The Finia system is designed to add cryoprotectant solutions to a cell suspension.
Prepare the transfer bag by puncturing the sample port with the spike (on the transfer bag) to open it and release pressure.
Thoroughly resuspend the cells and transfer them into a syringe with an 18-gauge needle.
Situate the needle in the sample port and fill the bag with the cell suspension.
Plug the sample port with the spike and twist it until tight to secure closure of sample port.
Use the T-SEAL Mobile to seal the tubing shut by the sample ports. This leaves the tubing available for welding to the tubing set.
Place the transfer bag on the hook outside the Finia system.
Connect the transfer bag to the Finia disposable set by placing the ends of the tubing into the welding device.
Note: The transfer bag line can be welded only onto the supply line/PVC portion of the Finia disposable set.
Close the welder and start the welding process.
Open the welder and open the weld.
Scan the barcode for the material and touch Confirm.
Confirm that the weld from the transfer bag to the disposable set is open and touch Continue to start the Finia procedure.
Note: The Finia system mixes and cools the cell suspension to the temperature set point. The system pumps three boluses of cell suspension into the mixing bag, after each bolus weighing the bag and reporting the measurement. Then, the system primes each product bag with cell suspension before extracting all air from the product bags. Once the product bags are filled, the bags are automatically sealed and the Finia process is complete.
Carefully remove the product bags and QC bag separately by pulling down the tubing at the top of each bag. Remove and process all product bags before removing any other material from the system. After the bags have been removed, touch Confirm on the screen to store all details of the process on the CPA.
In a BSC, open the QC bag and aliquot the contents into cryovials for downstream analysis of the cells.
Note: Air is not removed from the QC bag. The contents of the QC bag can be analyzed before or after cryopreservation.
On the Finia device, select View Procedure Summary to view the procedure details on the screen.
Remove the rest of the disposable set from the system.
Use the batch ID to retrieve the batch report from the CPA.
Cryopreservation of cells
Label and place the product bags into the stainless-steel canisters.
Place the canisters into the adjustable freezing rack.
Label and place QC cryovials in tube racks.
Perform freezing operations using the controlled-rate freezer set to a desired freezing program.
Note: The CryoMed program cools at a freezing rate of 1 °C/min from 4 to -40 °C, then at 10 °C per minute cooling rate, until reaching -90 °C end temperature.
Transfer the canisters to the accompanying frame.
Transfer the QC cryovials into storage boxes and place them in the storage rack.
Store the frame and rack into the LN2 vapor phase maintained at -140 °C to -180 °C.
Cell phenotype analysis by flow cytometry
Follow steps A1–A18 to thaw and record pre-and post-spin cell counts on MSCs and PBMCs.
Note: XSFM + 1% P/S was used to thaw MSCs or PBMCs.
Add 1 × 105 cells per well into a 96-well V-bottom plate [four technical replicates per sample and fluorescence minus one (FMO) control samples].
Centrifuge the plate at 500× g for 5 min and decant the supernatant.
Resuspend the cell pellets in 100 μL of viability stain solution (see Recipes) or PBS for controls (Zombie FMO and unstained control samples).
Incubate for 15 min at RT in the dark.
Perform a wash by resuspending the cells in 150 μL of FC buffer (see Recipes).
Centrifuge the plate at 500× g for 5 min and decant the supernatant.
For PBMC staining only, add 50 μL of Fc block solution (see Recipes) per well to all wells and incubate for 5 min at RT in the dark.
Resuspend cell pellets in surface marker stain mixture (see Table 1) or FMO mixture for controls and incubate for 20 min at 4 °C in the dark.
Table 1. Antibody information for cell phenotype panels
Surface marker Dilution (in FC buffer) Conjugate Vendor Catalog # Clone Excitation (nm) Emission (nm)
Human PBMCs
panel antibodies CD3 1:100 AF700 BioLegend 317340 OKT3 638 719
CD14 1:100 PE BioLegend 325606 HCD14 561 576
CD16 1:100 PerCP-Cy5.5 BioLegend 302028 3G8 488 679
CD19 1:100 PE-Cy7 BioLegend 302216 HIB19 561 780
CD56 1:100 APC BioLegend 362504 5.1H11 638 660
HLA-DR 1:100 FITC BioLegend 327006 LN3 488 517
Human MSCs panel antibodies CD14 1:100 PE-Cy7 BioLegend 367112 63D3 561 780
CD19 1:100 APC-Cy7 BioLegend 302218 HIB19 650 785
CD34 1:100 FITC BioLegend 343504 581 488 517
CD45 1:100 APC BioLegend 304012 H130 638 660
CD73 1:100 PE BioLegend 344004 AD2 561 576
CD90 1:100 PerCPCy5.5 BioLegend 328118 5E10 488 679
CD105 1:100 BV605 BD Biosciences 562664 266 405 421
HLA-DR 1:100 BV421 BioLegend 307636 L243 405 605
Centrifuge the plate at 500× g for 5 min and decant the supernatant.
Resuspend all cell pellets in BD CytofixTM fixation buffer.
Incubate for 15 min at 4 °C in the dark.
Centrifuge the plate at 500× g for 5 min and decant the supernatant.
Resuspend all cell pellets in the FC buffer and then perform data acquisition on the flow cytometer with appropriate settings and compensation for the designed panel.
Data analysis
Phenotypic analysis of cells “pre-cryopreservation” (post-Finia procedure, i.e., MSCs and isolated PBMCs prior to cryopreservation) and “post-cryopreservation” (post-Finia procedure, i.e., following thaw of cryopreserved MSCs and PBMCs) was performed by flow cytometry. The percentage of cells expressing surface markers outlined in Table 1 were analyzed following an initial gating strategy of forward and side scatter > singlets > live cells (Zombie UV live/dead discrimination) on >10K acquired events. Phenotypic analysis of MSCs showed comparable surface marker expressions for pre- and post-cryopreservation cells. Results showed positive expression (>95%) of markers CD73, CD90, and CD105 and negative expression (<5%) of markers CD14, CD19, CD34, CD45, and HLA-DR. For PBMCs, surface marker expressions of CD3 (34.0% for pre- and 33.0% for post-cryopreservation) and CD56 (16.3% for pre- and 16.9% for post-cryopreservation) were comparable (Figure 4A and 4C). In contrast, results showed marked differences between pre- and post-cryopreserved PBMCs related to the surface marker expressions of CD14 (24.2% for pre- and 44.2% for post-cryopreservation; P < 0.01), CD16 (25.7% for pre- and 30.8% for post-cryopreservation; P < 0.05), CD19 (13.6% for pre- and 17.1% for post-cryopreservation; P < 0.05), and HLA-DR (46.9% for pre- and 56.7% for post-cryopreservation; P < 0.05) (Figure 4B and 4D). The cell population that expressed CD14, CD16, CD19, and HLA-DR was enriched after cryopreservation, which suggests changes to surface marker expressions directly related to cryopreservation or incomplete recovery of all cell types within PBMCs following cryopreservation in which the loss of cells was not captured by the employed phenotyping panel. Overall, post-cryopreserved cells maintained surface marker expression as did fresh cells (pre-cryopreservation). Using the Finia system along with a controlled rate freezer can maintain surface marker expression for cells cultured as both adherent cells and suspension cells. The process can generate high-quality cells to ensure successful cell production.
Figure 4. Results of flow cytometry analysis of phenotypes pre- and post-cryopreservation. Quantitative comparison of surface marker expressions of mesenchymal stromal cells (MSCs) (A) and peripheral blood mononuclear cells (PBMCs) (B). Representative histograms of MSCs (C) and PBMCs (D) showing positive gate applied to each surface marker based on FMO controls. Statistical comparison of pre- and post-cryopreserved samples was performed using a paired t test. *, P < 0.05; **, P < 0.01; ***P < 0.001.
Validation of protocol
The protocol offers a robust and reproducible workflow for the processing and cryopreservation of adherent and suspension cells, which was demonstrated with MSCs and PBMCs. The following results are provided as validation of the protocol performed on the different cell types to compare measurable parameters representing the quality of the cell products before and after cryopreservation (Tables 2 and 3). Additionally, examples of the expected results generated from the CPA of the Finia system for each cell type are shown (Tables 4 and 5).
Table 2. Validation results for MSCs
Sample Cells/mL Viability (%) Diameter (μm) Total cells (× 106)
Sample 1
Fresh cell at harvest (pre-cryopreservation) 6.39 × 105 98.3 14.7 19.1 (in 30 mL)
Thawed cells pre-spin (QC vial; post-cryopreservation) 6.91 × 105 96.0 15.4 13.8 (in 20 mL)
Thawed cell post-spin (QC vial; post-cryopreservation) 6.67 × 105 93.6 15.4 13.3 (in 20 mL)
Thawed cell post-spin (product bag; post- cryopreservation) 5.65 × 105 90.4 15.8 5.65 (in 10 mL)
Sample 2
Fresh cell at harvest (pre-cryopreservation) 1.02 × 106 93.9 15.7 30.6 (in 30 mL)
Thawed cell pre-spin (QC vial; post-cryopreservation) 1.16 × 106 92.4 16.3 24.4 (21 mL)
Thawed cell post-spin (QC vial; post-cryopreservation) 1.14 × 106 90.8 16.1 23.9 (in 21 mL)
Table 3. Validation results for PBMCs
Sample: PBMCs Cells/mL Viability (%) Diameter (μm) Total cells (× 106)
Fresh cell at harvest (pre-cryopreservation) 3.36 × 106 99.0 9.9 137.8 (in 41 mL)
Thawed cell pre-spin (QC vial; post-cryopreservation) 2.99 × 106 97.1 10.0 101.7 (in 34 mL)
Thawed cell post-spin (QC vial; post-cryopreservation) 2.6 × 106 96.7 9.9 88.4 (in 34 mL)
In summary:
Both MSC runs resulted in >90% viabilities of the thawed cells post-cryopreservation with similar cell counts.
The Finia system allows for overage in the QC bag. Prior to sealing, the system pumps in all residual volume from the mixing bag to the QC bag.
In both cases, the Finia system run was 10 min from connecting the transfer bag to the end of the filling procedure.
Table 4. Example of results from the FINIA device run report for MSCs
Sample Planned (CPA protocol) Actual (CPA report)
Total volume for cell suspension in Cryostor CS10 26 mL 26 mL
Bag 1 MSC product 10 mL 9.9
QC bag 10 mL 11.8
Residual volume in tubing 6 mL -
Total end product (include QC vials) 20 mL 21.7
Note: Confirm that the transfer bag (materials) contains enough volume for the Finia system to prepare the mixing bag.
Table 5. Example of results from the FINIA device run report for PBMCs
Sample: PBMCs Planned (CPA protocol) Actual (CPA report)
Total volume for cell suspension in Cryostor CS10 36 mL 35.1 mL
Bag 1 product 20 mL 19.8
QC bag 10 mL 14.3
Residual volume in tubing 6 mL -
Total end product (include QC vials) 30 mL 34.1 mL
Note: Confirm that the transfer bag (materials) contains enough volume for the Finia system to prepare the mixing bag.
In summary:
The viability of the thawed cells post-cryopreservation was >90% with some cell loss from centrifugation.
The Finia system run was 10 min from connecting the transfer bag to the end of the filling procedure.
General notes and troubleshooting
General notes
Optimal thawing solutions (e.g., media), time, and centrifugation speeds need to be considered and may be cell type–specific for thawing cryopreserved cells.
Automated cell counters and manual counting methods may result in discrepancies to cell counts and viability assessments. Selecting the appropriate cell counting method, instruments, and associated ranges of detection needs to be considered and may be cell type–specific when obtaining cell counts and viability measurements.
Selection of culture media, enzyme-based digestion solutions for detachment of cells, proliferation rates, and culture conditions and duration are cell type–specific.
When estimating confluence of adherent cells cultured in HYPERFlask culture vessels, observing the bottom-most layer assumes an even distribution among the ten layers.
A density gradient centrifugation procedure is typically followed for the isolation of PBMCs from a leukopak or apheresis-derived products. Consideration of density gradient solutions, temperature, and centrifugation time and conditions are necessary to obtain a sufficient buffy coat comprised of PBMCs.
Retaining both the pellet and the supernatant from the first wash after Lymphoprep separation is important, as some PBMCs will remain in the supernatant.
The CPA can be adjusted for the number of materials used (i.e., cell suspension only or cell suspension plus cryopreservation solution), the volume of each material, the temperature, and the desired end product volume in each bag. The specific gravity information is important, as that is used by the Finia system to determine the volume in the bags. For PBMCs (1.077) and MSCs (1.056–1.075) and Cryostor CS10 (1.065–1.069), a specific gravity of 1.07 g/mL at RT was used.
The QC bag does not have air removal, and the allowable volume range on the QC bag is 10–40 mL.
An amount equaling 15% of the volume of cells plus cryopreservation solution was added to the transfer bag to account for dead volume in the transfer bag. Material left in the transfer bag (not pumped into the Finia system) is not recorded in the Finia run report. If an alarm is triggered due to insufficient volume being drawn from the transfer bag, it is usually necessary to transfer the bag contents to a new transfer bag and start the process again.
Start the welding process only when directed by the Finia system. Welding prior to loading the disposable set could cause unintended flow of material out of the transfer bag and trigger alarms.
Measure the height of the first canister insert in the frame to determine if the bag will sit in the liquid or vapor phase of LN2. If needed, add a platform riser to raise up the frame.
As described above, Finia has a user-configurable flexible protocol option. In this procedure, the authors utilized a manual method of cryoprotectant addition prior to loading into the mixing bag. The Finia system is designed to mix up to three materials and can automatically add a cryoprotectant solution to a cell suspension if desired. The system allows the user to establish notification flags to monitor the amount of time the cell suspension has been in contact with the cryoprotectant solution.
Troubleshooting
The Finia system must be restarted to receive a new protocol configured from the CPA.
Unloading prior to the end of the run should be avoided and only be used as a last resort, as this requires the whole process to start over, i.e., remove, recalculate based on the lost volume in the lines, and start again.
Finia procedures can be performed in an offline mode if the system loses connection to the network. The Finia system will send a run report once it re-establishes connection. The device may need to be restarted to send the run report. Updates from the CPA (e.g., protocol, users) cannot be added to the Finia device when the system is offline.
Always perform an action before reporting that an action has been taken, as the confirmation will trigger the next step in the process, and you cannot go back to the previous screen.
Considerations for current good laboratory practices (cGLP)
cGLP is a set of standards and guidelines that ensure the quality and reliability of non-clinical laboratory studies (Tarlengco, 2023). The FDA’s Code of Federal Regulations (21 CFR 58) outlines GLPs, which include considerations for Organization and Personnel, Testing Facilities Operation, and Test and Control Articles. The methods performed within this manuscript adhere to these cGLP guidelines. Within our facility, personnel are trained in donning appropriate PPE (hairnet, safety glasses, face mask, gown, shoe covers, and gloves) and aseptic technique is practiced, as with all cell culture studies, to ensure the correct preparation of cells in a sterile environment (Siddiquee, 2017). Along with the creation and adherence to standard operating procedures, reagents and solutions are properly labeled to be in accordance with cGLP guidelines. All fill-finish work was carried out in clean room space capable of supporting open and closed manufacturing processes for phase I good manufacturing practices evaluation. Environmental monitoring of non-viable and viable particle counts ensured that these counts remained under specified limits. For cryopreservation of samples, each container was labeled with initials, code/batch number, and expiration date and was properly handled and stored. All samples within this study were barcoded to distinguish each batch filled with the Finia system. Once the system had completed a run, the bags were collected and run through a controlled-rate freezing program for proper handling and storage of the samples. By following the GLP considerations, our laboratory ensured the reliability, reproducibility, and quality of the data generated.
Considerations for current good manufacturing practice (cGMP)
cGMP is a set of quality management principles and guidelines that govern the production, distribution, and supply of a health product (Gouveia et al., 2015). The World Health Organization (WHO) defines quality metrics for production and quality control (World Health Organization, 2023). Considerations and key aspects of GMP include Quality Management, Facility and Equipment, Personnel, Raw Materials, Process Control, Validation and Qualification, and Documentation. The FDA outlines the GMP guidelines within the Code of Federal Regulations (21 CFR 211). There are five Ps of GMP: people, products, processes, procedures, and premises (Tarlengco, 2023). Personnel are fully trained in their roles and duties and all products have clear specifications at every phase of production. All products must undergo testing and quality assurance before being distributed to customer s. The Finia system and the controlled rate freezer can be operated in GLP space such that potential cGMP applications can be executed.
Acknowledgments
This study was supported by the Billi and Bernie Marcus Foundation and Terumo Blood and Cell Technologies.
Competing interests
Authors A.R.H. and S.L.G. are affiliated with Terumo Blood Cell and Technologies.
Ethical considerations
No human subjects were used for any experiments included in this study. All procedures using MSCs derived from umbilical cord tissue supplied by collaborators at Duke University were in accordance with the ethical standards and approved by the ethics committee of the institutional review boards at Georgia Institute of Technology and Duke University (IRB Protocol No. H17348).
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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4,901 | https://bio-protocol.org/en/bpdetail?id=4901&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Correlative Light and Electron Cryo-Microscopy Workflow Combining Micropatterning, Ice Shield, and an In-Chamber Fluorescence Light Microscope
SB Sabrina Berkamp §
DD Deniz Daviran
MS Marit Smeets
AC Alexane Caignard
RJ Riddhi A. Jani
PS Pia Sundermeyer
CJ Caspar Jonker
SG Sven Gerlach
BH Bernd Hoffmann
KL Katherine Lau
CS Carsten Sachse
(§ Technical contact)
Published: Vol 13, Iss 24, Dec 20, 2023
DOI: 10.21769/BioProtoc.4901 Views: 2282
Reviewed by: Abhilash PadavannilLeeya EngelBeatrice LiVamseedhar Rayaprolu
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Abstract
In situ cryo-electron tomography (cryo-ET) is the most current, state-of-the-art technique to study cell machinery in its hydrated near-native state. The method provides ultrastructural details at sub-nanometer resolution for many components within the cellular context. Making use of recent advances in sample preparation techniques and combining this method with correlative light and electron microscopy (CLEM) approaches have enabled targeted molecular visualization. Nevertheless, the implementation has also added to the complexity of the workflow and introduced new obstacles in the way of streamlining and achieving high throughput, sample yield, and sample quality. Here, we report a detailed protocol by combining multiple newly available technologies to establish an integrated, high-throughput, optimized, and streamlined cryo-CLEM workflow for improved sample yield.
Key features
• PRIMO micropatterning allows precise cell positioning and maximum number of cell targets amenable to thinning with cryo focused-ion-beam–scanning electron microscopy.
• CERES ice shield ensures that the lamellae remain free of ice contamination during the batch milling process.
• METEOR in-chamber fluorescence microscope facilitates the targeted cryo focused-ion-beam (cryo FIB) milling of these targets.
• Combining the three technologies into one cryo-CLEM workflow maximizes sample yield, throughput, and efficiency.
Graphical overview
Keywords: CLEM cryo-EM Tomography Cryo FIB–SEM Fluorescence microscopy Maskless micropatterning
Background
In the conventional in-situ cryo-electron tomography (cryo-ET) workflow, initially cultured cells expressing a fluorescent marker gene are required to adhere to an electron microscopy grid before they are cryo-fixed by plunge freezing (Iancu et al., 2007; Rigort et al., 2012; Mahamid et al., 2016). To navigate through the complex environment of the frozen-hydrated cell, correlative light and electron microscopy (CLEM) approaches were introduced into the workflow to localize the regions of interest (ROIs) (van Driel et al., 2009; Schorb and Briggs, 2014; Klein, Wachsmuth-Melm, et al., 2021). To this end, the grids are typically transferred to a cryo-fluorescence microscope to acquire spatial information on various features of interest using fluorescence (Arnold et al., 2016; Chakraborty et al., 2020). Following a second cryo-transfer step to a cryo focused-ion-beam–scanning electron microscopy (cryo FIB–SEM), the cells are thinned down to ~150 nm-thick lamellae using a focused ion beam, making them amenable to cryo-ET or cryo-EM imaging (Marko et al., 2007; Rigort et al., 2012; Villa et al., 2013; Lucas et al. 2021).
The workflow provides natively preserved insight into the structure-dependent functionality of cellular components at molecular or even close-to-atomic resolution (Xing et al., 2023). However, the complexity of the workflow poses challenges that frequently compromise the throughput and success rate (Hylton and Swulius, 2021). To name a few: i) conventional cell culture procedures lack proper control over cell shape and cell positioning on the grids. This limitation often results in cell clustering and cells positioned on grid bars rather than in the center of the grid squares, both of which make them unsuitable for FIB milling (Wagner et al., 2020). ii) The time-consuming nature of cryo FIB milling requires long residence time of the lamellae within the vacuum chamber. Consequently, the risk of in-chamber ice contamination significantly increases during batch milling sessions, making it impossible to mill multiple lamellae in an overnight session (Buckley et al., 2020; Tacke et al., 2021). iii) Combining CLEM approaches with cryo FIB milling allows for targeted preparation of lamellae at each intended region of interest (ROI). However, the cryo-transfer steps to and from the cryo-fluorescence microscope significantly increase the chance of ice contamination and devitrification of the sample. iv) The conventional way of lamella preparation and target determination is time consuming and complicated. Therefore, the error-prone procedure prior to lamella milling can easily lead to a mismatch in correlation and result in the loss of the ROI (Fukuda et al., 2014; Klein, Wimmer, et al., 2021).
In this protocol, we describe the details of a combined workflow with several newly available technological upgrades. i) Micropatterning the electron microscopy (EM) grids prior to cell culture using PRIMO: the customized patterns result in improved control over cell shape and spatial distribution over the entire grid (Engel et al., 2019; Swistak et al., 2021). By covering the EM grid in an anti-fouling layer and selectively removing it through UV light, the cell can only grow on selected areas of the grid. Provided the applied mask of the UV light is not too big, the cell will adapt to the mask shape, allowing one to reposition neuronal axons, study stress fibers in a specific area of the cell periphery, or adjust the cell shape in some other way. Consequently, more millable sites will be available per grid, making an overnight milling session worthwhile (Toro-Nahuelpan et al., 2020; Sibert et al., 2021). ii) Cryo FIB milling in the presence of the CERES ice shield: the CERES ice shield eliminates the parasitic growth of an amorphous ice layer on top of the lamellae. Without it, lamella can be rendered unusable due to excessive ice contamination at the end of an overnight batch milling session (Tacke et al., 2021; Lau et al., 2022). iii) Continuous, in-chamber fluorescence detection using METEOR: by integrating the METEOR cryo-fluorescence microscope into the vacuum chamber of the cryo FIB–SEM, one major cryo-transfer step is eliminated from the workflow, which significantly reduces the chances of sample devitrification, contamination, and mishandling (Smeets et al., 2021). iv) The fluorescence integration in the FIB–SEM further allows for continuous monitoring of the ROI within the lamella during the milling process, thus eliminating the need for separate signal correlation to find back the ROI. This protocol has been proven to substantially increase the number of produced usable lamellae per milling session and hence surpass the conventional achievable success rate and throughput for high-resolution imaging in a cryo-transmission electron microscope (cryo-TEM). One can normally expect to produce 3–7 lamella in a fully manual milling session, and an estimated 25 lamella in automated session using micropatterned grids and the technological improvements described in this protocol, which are summarized in Table 1.
Table 1. Estimation of the effect of different workflow improvements on the output
Estimated number of amenable milling sites per grid Estimated number of final lamellae per grid
Manual milling, no micropatterning 8 ~4
Manual milling + micropatterning 40 ~8
Manual milling + micropatterning + CERES ice shield 40 ~10
Automated milling + micropatterning + CERES ice shield + METEOR 40 ~25
Materials and reagents
Biological materials
Human immortalized RPE-1 cells (ATCC, hTERT RPE-1, CRL-4000), stably expressing human mCherry-p62
Reagents
200 mesh gold Quantifoil grids R2/1 with SiO2 support layer (Quantifoil, catalog number: N1-S15nAu20-01, 100/pk)
PLL-g-PEG [SuSoS, catalog name: PLL(20)-g[3.5]- PEG(5)]
Poly L-Lysine (Sigma-Aldrich, catalog number: P8920)
mPEG-Succinimidyl Valerate MW 5,000 (mPEG-SVA) (Laysan Bio, Inc. catalog number MPEG-SVA-5000)
14.5 mg/mL PLPP (1 mL PLPP solution) [Alvéole, catalog number: PLPP (for liquid) or PLPP_Gel (for gel)]
Fibronectin (human native fibronectin) (Gibco, catalog number: PHE0023)
Trypan blue (0.4% trypan blue stain) (Invitrogen, catalog number: T10282)
Trypsin (0.05% Trypsin-EDTA solution with phenol red) (Gibco, catalog number: 25300054)
Pen/strep (penicillin-streptomycin, 10.000 U/mL) (Gibco, catalog number: 15140122)
FCS (Fetal Bovine Serum) (Sigma-Aldrich, catalog number: F7524)
Phosphate buffered saline (PBS), pH 7.4 (Gibco, catalog number: 10010056)
DMEM (DMEM/F-12, GlutaMAX supplement) (Gibco, catalog number: 31331028)
Fluorobrite (FluoroBrite DMEM) (Gibco, catalog number: A1896701)
Laboratory supplies
Tweezers (Biological Tweezers, High Alloy DX Style 5) (Electron Microscopy Sciences, catalog number: 78325-5DX)
Parafilm (Parafilm M) (Sigma-Aldrich, catalog number: P7793)
Petri dish (900 mm Petri dish) (VWR, catalog number: 391-2016)
Primo stencil (PDMS stencil with 4 wells) (Alvéole, catalog number: PDMS_STENCILS_4)
30 mm glass-bottom dishes (Cellview cell culture dishes, 35/10 mm, glass bottom) (Greiner Bio-One, catalog number: 627860)
T75 tissue culture flasks (T75 tissue culture flasks) (TPP, catalog number: Z707503-100EA)
Fluorescent marker (Staedtler, Textsurfer classic 364)
Black marker (black laboratory marker) (SecurLine, catalog number: 3083.2)
Cell counting slides (Countess Cell Counting Chamber Slides) (Invitrogen, catalog number: C10228)
Cell strainer (PluriStrainer mini 40 μm) (PluriSelect, catalog number: 43-10040-40)
15 mL falcon tubes (15 mL conical centrifuge tubes) (Falcon, catalog number: 352196)
Eppendorf tubes (Safe-Lock 1.5 mL micro test tubes) (Eppendorf, catalog number: EP0030123328)
1 μm FluoSpheres (FluoSpheres, carboxylate-modified microspheres) (Invitrogen, catalog number: F8816)
Ethane (Air Liquide, catalog number: P0500S10R0A001) or ethane/propane mix (37% ethane/63% propane, Nippon gas, catalog number: 611961)
Grid box for unclipped grids (Jena Biosciences, catalog number: X-CEM-201)
Grid box for clipped grids (Autogrid Compatible Cryo Grid Box) (MiTeGen, catalog number: M-CEM-7AGB)
Autogrid rings with cutout for FIB–SEM (Thermo Fisher Scientific, catalog number: 1205101)
C-ring (Thermo Fisher Scientific, catalog number: 103171)
Equipment
Pelco easiGlow (Pelco easiGLOW Glow discharge Unit) (Ted Pella Inc. catalog number: 91000)
Pelco TEM grid holder block (Ted Pella inc., catalog number: 16820-25)
Fridge (Samsung, model: RL30J3005SA)
Inverted confocal or epifluorescence microscope (Nikon Eclipse Ti2 is used in this protocol)
PRIMO photopatterning device [Alvéole, Photopatterning Device consisting of a (DMD + laser 375 nm, 75 mW) Modulus, Driving Electronics, IHM software LEONARDO]
Mammalian cell incubator (Binder, catalog number: 9040-0112)
Laminar flow hood (Lamsystems, Uniflow, catalog number: 2E-B.003-15)
Water Bath (Julaba Pura 10, catalog number: 9 550 510)
Cell counter (Countess II automated cell counter) (Invitrogen, catalog number: AMQAX1000)
Widefield microscope (Zeiss Axio Vert.A1 FL)
Centrifuge (Eppendorf, model: 5702RH, catalog number: 5704000010)
Vitrobot Mark IV (Thermo Fisher Scientific)
Heating plate (CultureTemp 37 °C, Bel-art Products, catalog number: CHC7.1)
Clipping station (Thermo Fisher Scientific, catalog number: 1130697 or SubAngstrom product code CSA01)
Aquilos2 (Thermo Fisher Scientific)
CERES ice shield (Delmic B.V., catalog number: 2708-999-0010-1)
METEOR system (Delmic B.V., catalog number: 2707-999-0014-2)
High-end cryo TEM with direct electron detector for tomogram acquisition. Here, a 200 kV Talos Arctica G2 (Thermo Fisher Scientific) equipped with a Bioquantum GIF (Gatan) and K3 direct election detector (Gatan) was used
Software and datasets
Inkscape or Adobe Illustrator (2020, version 24.1 used here)
Leonardo (version 4.15, https://www.alveolelab.com/our-products/leonardo-photopatterning-software/) (Access date, September 2023)
xT Microscope Control (version 20.1.1) (Access date, September 2023)
MAPS (version 3.19, https://www.thermofisher.com/order/catalog/product/de/en/MAPS2) (Access date, September 2023)
AutoTEM (version 2.4.1, https://www.thermofisher.com/order/catalog/product/AUTOTEM5?SID=srch-srp-AUTOTEM5) (Access date, September 2023
ODEMIS (version 3.2.2, https://github.com/delmic/odemis) (Access date, September 2023)
Procedure
Micropatterning of electron microscopy grids (Figure 1)
Figure 1. Overview of the micropatterning process. The electron microscopy grids are glow discharged to render them hydrophilic. (B) The grids are incubated in drops with PLL-g-PEG that acts as an anti-fouling layer. (C) The fluorescence microscope equipped with an Alvéole PRIMO device is focused and the DMD mirrors are calibrated. (D) The grids are incubated in PLPP, and the PLL-g-PEG layer is removed locally with a UV laser. (E) The grids are incubated with fibronectin or another extracellular matrix (ECM) protein. (F) Mammalian cells are seeded on the micropatterned grids.
Grid passivation: application of anti-fouling layer on the grids. In this step of the process, the entire grid surface is covered in PLL-g-PEG, which prevents cell adhesion.
Take several 200 mesh gold Quantifoil R2/1 grids (SiO2 support layer).
Glow discharge both sides of the grid using a Pelco easiGlow plasma cleaner. Grids are placed in a holder or on a glass slide covered in parafilm for easier pick up and glow discharged at vacuum: 15 mA, 0.4 mBar, 60–90 s with 10 s hold. Notice the purple light indicating glow discharging is taking place. Flip over the grids with a tweezer to glow discharge the other side using the same settings. Glow discharging will allow the grid to be properly treated in the next steps, without the grid floating to the surface of droplets or sticking to the dishes used (Figure 1A).
Place the grids on a sheet of parafilm in a Petri dish with 30 μL drops of PLL-g-PEG, ~0.5 mg/mL in PBS, and incubate them overnight at 4 °C (Figure 1B).
Wash the grids by dipping them vertically in PBS several times.
Store the grids temporarily in a dish with PBS. The grids can be stored for several days at 4 °C.
Micropatterning of grids using PRIMO photopatterning device and PLPP.
Turn on the fluorescence microscope and PC.
Draw the pattern for patterning in Inkscape or Adobe Illustrator. Save it as a .png file with 125 DPI.
Place an empty glass-bottom dish onto the microscope for calibration purposes. Mark the dish with a fluorescent marker and draw a small cross with a black marker.
Insert a 20× aperture (20× S Plan Fluor ELWD, 0.45 NA, Air is used in this protocol).
Focus the microscope on the black cross and set the perfect focus system or other autofocus system at this z-height (Figure 1C).
Move the dish slightly to an area with fluorescent marker, open the Leonardo software, and click on Calibrate tube lens.
Choose a calibration pattern from the options supplied in the Leonardo software. You can choose any pattern to calibrate the tube lens, but we recommend a pattern with features going down to 1 µm. Additionally, we recommend choosing a pattern that will resemble the pattern you will actually use for your experiment. If you will pattern lines, then choose a pattern with lines. If you will pattern dots or some exotic shapes, then choose to calibrate with dots. The Leonardo software contains basic calibration patterns to choose from.
Turn down the laser power and exposure time until you clearly see the smallest sub-structures in the pattern.
When the pattern is out of focus, turn the DMD wheel on the side of the PRIMO device.
On the next screen, the pixel ratio and the laser power are displayed. Confirm these are within expected values.
Stick a PRIMO stencil with 4 mm circular holes in a new 30 mm glass-bottom dish.
Pipette 2 μL of PLPP in a hole and ensure it spreads to cover the entire surface of the hole.
Take a grid and blot away excess PBS.
Critical: Do not blot completely dry.
Place the grid in the PLPP droplet and replace the lid of the dish to prevent the grid from drying out and place the dish on the microscope. The orientation of the grid does not matter (Figure 1D).
Center the grid and focus on the Quantifoil film using the reflected light mode on the microscope.
Open your pattern file in the Leonardo software. The pattern contains a grid of white masks on a black background and determines the appropriate diameter of the shape for your cell type. The cell needs to have sufficient space to grow and spread, while ideally restricting the area to one cell per masked area (Figure 2). Move and rotate the pattern so that it is placed in the center of the grid squares (if desired) and at the center of the EM grid near the asymmetric mark. You may need to access the advanced features to enable pattern rotation. It is advisable to pattern only the squares in the middle of the grid for easier access with FIB milling and tomography.
Figure 2. Examples of patterns of employed masks used on EM grids. A. An 8 × 8 grid of 20 μm circular masks used in the micropatterning procedure. B. An 8 × 8 grid of 30 μm circular masks used in the micropatterning procedure.
Set the dose to 900 mJ/mm2 and enable stitching mode.
Switch the fluorescence microscope to the correct optical path, compatible with the PRIMO device.
Lock the pattern in place and press start to begin the micropatterning procedure. You should see small spots of light reflecting off the grid surface. This process should take 3–10 min, depending on the size of the pattern and the number of squares that you want to pattern.
Immediately remove the grid after the micropatterning procedure is done to avoid excess PLL-g-PEG layer degradation and wash the grid several times in fresh PBS.
Place the grids in a fresh 30 mm glass-bottom dish filled with PBS.
Wash the glass-bottom dish with the stencil used for micropatterning before repeating the protocol for the other grids.
Pause point: The micropatterned grids can be stored in PBS at 4 °C for up to four weeks. Critical: Be careful and avoid grids drying out.
Side protocol—micropatterning of grids using PRIMO and PLPP gel:
Alternatively, the micropatterning of grids with PRIMO can be done using PLPP gel instead of PLPP (Engel et al., 2021). Indeed, the PLPP photoinitiator from Alvéole is also available in a gel form, which speeds up the photopatterning 30 times in comparison to the liquid form. Another advantage is that you can flip the sample upside down to be able to print on the EM grid bars if needed. The protocol using PLPP gel requires several drying steps of the grid, and the passivation is made with polylysine (PLL) and then mPEG-SVA:
• As in the regular PLPP procedure described above, glow discharge and place a PDMS stencil on your grid.
• Grid passivation: incubate your grid with a PLL solution (100 μg/mL for 30 min), rinse three times with 20 mM HEPES at pH = 8.5, and incubate your grid again with a solution of mPEG-SVA (10–20 μL for 1 h). Alternatively, an incubation with PLL-g-PEG can be done as described above.
• Micropatterning with PRIMO: a drop of PLPP gel (1 μL) is diluted in deionized water (2 μL) and spread on the grid. After the complete drying of the solution, the grid is placed upside down on the microscope deck holder.
• The steps using the Leonardo software are then similar to the PLPP protocol described above, with significantly lower UV doses (20–40 mJ/mm2).
Incubation of grids with fibronectin.
Prepare a Petri dish with a small square of parafilm.
Place 30 μL drops of fibronectin (50 μg/mL) on the parafilm and place a grid on each drop. Incubate at room temperature for 20–30 min. The fibronectin solution was prepared by diluting the fibronectin stock solution in PBS.
Wash the grids twice in PBS for 5 min and once with DMEM before use.
Critical: Be careful and avoid grids drying out.
Pause point: The grids can be stored for up to seven days in PBS at 4 °C. Critical: Be careful and avoid grids drying out.
Seeding mammalian cells on micropatterned grids
The next steps are all performed under sterile conditions in a laminar flow hood.
Preparing the cells for seeding.
Check the cells under a standard microscope for viability and cell health. Our cells were maintained in T75 flasks in 12 mL of DMEM media supplemented with Pen/strep and FCS.
Aspirate the media and briefly wash the cells with warm PBS.
Add 2 mL of 0.05% trypsin and incubate for several minutes at 37 °C with 5% CO2 until the cells detach from the dish.
Add 5 mL of DMEM, resuspend the cells, and pipette them in a 15 mL Falcon tube.
Centrifuge the cells at 500× g for 5 min at 37 °C.
Aspirate the supernatant and resuspend the cells carefully in a small amount of fresh, warm DMEM.
Pass the entire cell suspension through a 40 μm cell strainer into a new 15 mL Falcon tube to obtain single cells.
Mix 10 μL of cell suspension with 10 μL of Trypan Blue and pipette 10 μL of the mixture on a counting slide. Use the cell counter to determine the concentration of living cells in number of cells/mL.
Place the micropatterned grids in ~1 mL of DMEM supplemented with Pen/strep and FCS in a fresh 30 mm glass-bottom dish and carefully pipette 200,000 cells dropwise on top of the grids. Ensure the grids are on the bottom of the dish, Quantifoil film up, evenly spaced and a short distance away from the edges of the dish. Incubate the dish with the grids and the cells in the cell incubator at 37 °C with 5% CO2. Save the remainder of the cells in a warm place (Figure 1E).
Critical: Optimize the cell density and incubation time for your cell line. Adding too many cells will lead to cells growing also on areas of the grid covered with an anti-fouling layer. Adding too few cells will lead to reduced throughput and loss of enough sites suitable for FIB milling.
Assessment of the grids.
After 30–60 min, remove the grids from the incubator and look at them under a light microscope.
The cells should be attaching themselves to the areas of the grid that were micropatterned and to the areas of the dish surrounding the grids. When the micropatterned areas do not have a lot of cells, add additional cells dropwise and incubate again.
Incubate the cells on the grid 4–5 h before plunge freezing at 37 °C with 5% CO2.
Vitrification of mammalian cells
Turn the vitrobot on and warm the chamber to 37 °C with 85% humidity.
Pre-warm some Fluorobrite medium on a small heating plate of the incubator and, when needed, make a suspension of 1 μm FluoSphere fiducials in Fluorobrite medium (1:30).
Assemble and cool down the nitrogen container with liquid nitrogen. Once it is cold, fill the ethane container with ethane or ethane/propane mix. The ethane will turn cloudy and freeze on the sides of the cup when the cryogen is at the correct temperature.
Place the grids, Fluorobrite medium for washing, and the fiducial suspension on a 37 °C hot plate or small incubator.
Place the blotting paper on one blotting pad and a ring of parafilm on the other blotting pad for backside blotting.
For each grid, carefully pick the grid up with the vitrobot tweezers by the outer ring, trying not to disturb the Quantifoil film. Attach the tweezers to the vitrobot with the top of the grid and the cells facing towards the pad with the parafilm. Briefly dip the grids in Fluorobrite to remove the autofluorescent DMEM medium.
Add 3.6 μL of 1:30 FluorSpheres fiducial suspension right before plunge-freezing. Ensure the suspension is properly mixed by vortexing.
Blot the grids for 8.5 s with 10 blot force and 1 drain time. As vitrobot machines are not entirely reproducible, the settings may need to be adjusted when needed to increase or reduce ice thickness. Yearly instrument alignment performed by Thermo Fisher is advisable to help with reproducibility. The blot force can be measured according to the protocol in Sader et al. (2020).
Pause point: After plunge-freezing, store the grids in a grid box for unclipped grids in a liquid nitrogen dewar.
Thinning of the cells to ~150 nm using cryo FIB–SEM
Assessment of ice contamination rate in the FIB–SEM chamber.
Notes: Chamber vacuum can affect the contamination rate. Installation of the Delmic CERES ice shield can reduce the contamination rate significantly. Assessment of the contamination rate under different conditions can help determine how long a user can mill the grid without the lamellae deteriorating to an unusable thickness (Figure 3).
Figure 3. Micropatterning of EM grids can be used to greatly improve grid quality for cryo focused-ion-beam–scanning electron microscopy (FIB–SEM). A. Top: schematic of a grid pattern. Bottom: a typical EM grid that was incubated with human mesenchymal stem cells. B. Top: grid pattern with 20 μm diameter circles in the center of the grid square used for PRIMO photopatterning before seeding cells on the grid. Bottom: photopatterned EM grid with centrally adhered RPE-1 cells. C. Top: two touching 20 μm diameter circles were patterned in the center of all grid squares to align cell adherence to allow for milling from different angles. Bottom: an EM grid that was photopatterned with the PRIMO system. Possible milling sites are indicated with red arrows.
Load an empty, clipped grid into the FIB–SEM.
Navigate to an area in the center of the grid, focus, and determine eucentric height. Use a stage rotation of 110° and a stage tilt of 30°. For the full procedure, see details below.
Set the magnification to cover approximately 4 μm horizontal field width (HFW) and image with the electron beam. Focus on the film next to the holes. Use a scan rotation of 0°.
Pick two holes in the support film close to the top of the grid and measure the thickness of the film on the edge of the holes. Repeat this for two holes in the center of the grid and two holes close to the bottom of the grid.
Wait a fixed amount of time and repeat the procedure. Imaging the same area multiple times could shrink the built-up ice layer. To avoid this, after each timepoint, use a different set of holes to do the measurement.
Plot the data in a graph and calculate the slope to obtain the contamination rate.
Selection and preparation of sites appropriate for milling.
Preparing the cryo FIB–SEM:
i. Wake up the system in the xT microscope Control software.
ii. In the Beam Control tab, select Sputter Vacuum and purge the lines five times. This should take approximately 20 min. Return to High Vacuum when done.
iii. In the Cryo TEM Preparation tab, purge the GIS lines for 30 s.
iv. Fill the dewar with liquid nitrogen and place the heat exchanger inside to cool down the system.
Clipping the grids:
i. Follow Thermo Fisher’s procedure to clip the grids.
Critical: Work carefully to avoid buildup of ice contamination on the grids.
ii. Use special Auto-grid rings with a cutout to improve grid access. Mark the edges of the ring with a black or blue marker to help position the grids in the autoloader of the TEM microscope.
iii. Save the grids in a grid box for AutoGrids under liquid nitrogen.
Loading the grids into the FIB–SEM:
i. Cool down the loading station and insert a pre-tilted shuttle.
ii. Once it is cooled down, place the grid boxes into the station and load two grids into the shuttle. Ensure that the Autogrid cutout is positioned perfectly at the top. Work carefully to avoid buildup of ice contamination on the grids.
iii. Quickly load the shuttle into the FIB–SEM chamber and check proper position using the in-chamber camera before disconnecting the transfer rod.
Atlas acquisition:
i. Move the shuttle to the mapping position in the chamber using the xT Microscope Control software. It may help to set the scan rotation for the ion and electron beams to 180° for easier visualization.
ii. Using the SEM beam and the Everhart-Thornley detector (ETD), focus the beam on the grid and correct for astigmatism. Use a 2 kV beam at 25 pA for samples with thin ice and a 5 kV beam for samples with thicker ice.
iii. Open MAPS software and take a snapshot of the whole grid.
iv. Place a tile set on the center of the grid, 5 × 7 tiles with a tile height of 600 μm HFW and a total HFW of 2.76 mm. Set the resolution to 3,072 × 2,048, the pixel size to 195 nm, and the dwell time to 1 us. Start the acquisition.
Eucentric height and milling angle determination:
i. Mark the potential milling sites in the MAPS software by clicking Add Lamella Site Here. Ideally, these are single cells that are positioned in the middle of the grid squares, without any ice contamination on top. Only squares close to the center of the grid should ideally be selected, as the sites close to the Autogrid ring will yield small lamellae due to a high milling angle.
ii. Navigate to each site using the Drive To Mapping Position button, increase the magnification of the SEM beam, and focus the beam on the carbon film. Use 2–5 kV, 13 pA, and a 300 ns dwell time. While in live mode, link z.
iii. Determine the eucentric height of each milling site using a manual approach; the prompts are available in MAPS or the automated procedure is available in AutoTEM. Turn on the ion beam and use the ETD detector to image each site. Focus the beam. Use 30 kV, 30 pA, and 100 ns dwell time. Tilt the stage to 20° and image the site. If the edge of the Autogrid ring is in view, increase the tilt angle. When it is not in view, decrease the angle. From the point where the edge of the Autogrid ring is in view, tilt up 1–2° to get sufficient clearance for the ion beam not to be deflected. Mark this milling angle in MAPS software by clicking Store Angle. The icon next to the name of the lamella site will change from orange to green.
iv. Mark each site at the eucentric height and the milling angle in the FIB–SEM GUI for easier navigation later.
Assessment of milling sites by the METEOR system.
Make sure the z-height of the stage is linked in the microscope GUI.
Open the ODEMIS software on the METEOR microscope PC and make a new project folder. Press FM imaging to move the stage to the METEOR position and insert the objective lens (LMPLFLN 50×/NA = 0.5/WD = 10.6 mm).
Add imaging streams for each fluorophore, select the power for each channel, and set the exposure time to ~300 μs. Usually, 200–400 mW is sufficient, depending on the fluorophore, the abundance of the ROI, and the camera on the system. Focus the image using the autofluorescence signal in the green channel by slowly approaching the sample with the objective lens until the signal inside the target cell is sharp or until you can see the holes in the Quantifoil support film due to its autofluorescent properties. Critical: Do not insert the objective lens too far, as it can crash into the shuttle and damage the lens.
Acquire an image of each lamella site to confirm the presence of your ROI in the cells. A good start for camera settings is: bin 1, resolution 1,190 × 928 px, a gain of 16-bit, and a readout rate of 310 MHz. Adjust the exposure time and the power in the rest of the channels to a value that will yield a clear signal but not an oversaturated histogram.
Save these imaging positions in the FIB–SEM xT Microscope Control software for easier navigation later.
Switch back to the milling site in the chamber by pressing SEM imaging. Wait for the objective lens to retract and the shuttle to move back to the milling position in the chamber.
Navigate to the next milling position using MAPS or the saved positions in the xT Microscope Control software and repeat Steps D3d–D3f.
Setup of batch milling protocol.
Open AutoTEM. It should read in all the positions from the MAPS software.
Apply a template with the following parameters (Table 2) to all sites.
Table 2. Milling parameters used in AutoTEM. Description of all parameters that were imported in AutoTEM and used in this protocol. Sample-dependent optimization may be required.
Preparation
• Ion HFW Oversize: 80 μm
• Eucentric Tilt: disabled
• Artificial Features: disabled
• Milling Angle: disabled
• Image Acquisition: disabled
• Lamella Placement: enabled
- Ion HFW Oversize: 150%
- Minimal Ion HFW: 10 nm
Milling
• Lamella Size: 17 μm × 3 μm
• Correction Factor: 0.60
• Delay: disabled
• Reference Definition: enabled
• Electron Reference Definition: disabled
• Stress Relief Cuts: test if cuts help prevent lamella bending in your sample
• Reference Redefinition 1: disabled if no Stress Relief Cuts were made, otherwise enable
• Rough Milling:
- Pattern Offset: 1 μm
- Front Pattern Height: 7.2 μm
- Rear Pattern Height: 6.3 μm
- Depth Correction: 30%
- Front Width Overlap: 1.5 μm × 1.5 μm
- Rear Width Overlap: 1.5 μm × 1.5 μm
- Milling Current: 0.5 nA
- Pattern Type: Rectangle
- DCM Rescan Interval: 120 s
- Show Graphics: enabled
• Rough Milling - Electron Image:
- 1,536 × 1,024, 3 μs
- Enable ACB
- Enable Auto Focus
- HFW: 70 μm
• Reference Redefinition 2: enabled
• Medium Milling:
- Pattern Offset: 600 nm
- Front Width Overlap: 650 nm × 650 nm
- Rear Width Overlap: 650 nm × 650 nm
- Overtilt: 0°
- Depth Correction: 160%
- Milling Current: 0.3 nA
- DCM Rescan Interval: 90 s
- Pattern Overlap: 400%
- Pattern Type: CleaningCrossSection
• Medium Milling - Electron Image:
- 1,536 × 1,024, 3 μs
- Enable ACB
- Disable Auto Focus
- HFW: 70 μm
• Fine Milling:
- Pattern Offset: 300 nm
- Front Width Overlap: 350 nm × 350 nm
- Rear Width Overlap: 350 nm × 350 nm
- Overtilt 0°
- Depth Correction: 160%
- Milling Current: 0.1 nA
- DCM Rescan Interval: 60 s
- Pattern Overlap: 200%
- Pattern Type: CleaningCrossSection
• Fine Milling - Electron Image: disabled
• Finer Milling:
- Pattern Offset: 200 nm
- Front Width Overlap: 50 nm × 50 nm
- Rear Width Overlap: 50 nm × 50 nm
- Overtilt 0°
- Depth Correction: 140%
- Milling Current: 50 pA
- DCM Rescan Interval: 30s
- Pattern Overlap: 200%
- Pattern Type: CleaningCrossSection
• Finer Milling - Electron Image: disabled
Thinning • Disabled
Navigate to each milling position by clicking the saved position in the xT Microscope Control software, focus both beams, and click Update in the AutoTEM GUI. The software will take an image and store the stage position and focus parameters for later use.
Enable the sites in the Milling List one by one. For each site, click Run, enable only the Preparation section of the protocol, and click Ok. AutoTEM will take an image and place a milling box in the center of the image according to the specifications in the template. Adjust the size and shape of the milling box and place them in the middle of the cell, with the center positioned where your ROI is. Note that some part of the cell should remain on either side to maintain the lamella. Press Ok to confirm the site and save it in the program. Navigate to the next site in the xT Microscope Control software and repeat the steps D4c–D4d.
Once all sites are marked and appear green in AutoTEM, apply a GIS layer on the grid for 8 s in a FIB–SEM without lift-out system or 90 s for a system with a lift-out system. The layer deposition is initiated by navigating to the Cryo TEM preparation tab in the xT Microscope Control software, selecting the grids you want the GIS to be deposited on, setting the Flow Duration, and clicking Start.
Fill the dewar of the FIB–SEM to ensure sufficient liquid nitrogen is present for the automated milling process to finish.
Click Run in AutoTEM. In the pop-up window, only enable the Rough Milling tab and make sure all sites are clicked. Disable Fine Milling when you plan on doing the lamella polishing manually. Set the program to mill step-wise and start the program.
Once you start the automated rough milling procedure, the CERES ice shield should automatically insert and stay inserted during all milling steps.
Assessment of lamella quality with METEOR system.
Move the stage to the imaging position and insert the objective lens (LMPLFLN 50× /NA = 0.5/WD = 10.6 mm).
Reuse the same imaging streams for each fluorophore as desired and set the exposure time to ~300 μs. Focus the image using the autofluorescence signal in the green channel by slowly inserting the objective lens.
Take an image of each lamella site to confirm presence of your ROI in the lamellae using an exposure time that will yield a clear signal, but not resulting in an oversaturated histogram.
By using the saved imaging positions (step D4c) in the xT Microscope Control software, switch directly to the next imaging site. Make sure to retract the objective lens 5–10 mm before executing the stage movement.
Once all lamellae are imaged, move the shuttle back by pressing SEM imaging in the ODEMIS software.
Polishing the lamella.
In AutoTEM, navigate to each site individually and manually inspect to confirm the presence of a good lamella. Focus the beam.
Polish the lamella step-wise by using a 30 kV ion beam at 30 pA using two rectangular boxes placed ~300 nm apart in the xT Microscope Control software. Regularly monitor the lamella and look for cracks, holes, bending, and a disappearing GIS layer on the leading edge of the lamella. It can be helpful to enable iSPI in xT Microscope Control and set it to automatically take an SEM image every 2–8 s. More SEM images will damage the lamella, so be conservative.
Move the boxes in closer to ~200 nm and repeat the process.
When the lamella remains intact and stable, lower the beam current to 10 pA, bring the boxes closer to ~150 nm or less, and repeat the thinning process.
Critical: If the leading GIS layer starts degrading, it can be helpful to continue thinning using an overtilt. Disable the lower milling box and tilt the stage an additional +0.5°–1°. Mill only from the top at a 10 pA current. When the back of the lamella starts degrading or the GIS disappears, stop the milling process.
Appropriate lamella thickness can be estimated by taking two images with the SEM. Take an image at 5 kV and one at 2 kV. At around 250–300 nm lamella thickness, this lamella will appear dark gray in the 5 kV image but white or light gray in the 2 kV image. When the lamella thickness approaches 150–200 nm, the lamella will appear black in the 5 kV image and dark gray in the 2 kV image.
Final image acquisition with METEOR system.
Move the stage to the imaging position and insert the objective lens (LMPLFLN 50× /NA = 0.5/WD = 10.6 mm).
Reuse the same imaging streams for each fluorophore as desired and set the exposure time to ~300 μs. Focus the image using the autofluorescence signal in the green channel by slowly inserting the objective lens.
Take an image of each lamella site to confirm presence of your ROI in the final lamella using an exposure time that will yield a clear signal, but not resulting in an oversaturated histogram. This may require an exposure time of up to 30 s in case of a small ROI or a small amount of fluorophore present.
By using the saved imaging positions in the xT Microscope Control software, switch directly to the next imaging site. Make sure to manually retract the objective lens 5–10 mm before executing the stage movement.
Once all lamellae are imaged, export all images as “print-ready tiffs” after adjusting the histogram outliers to get the final image.
Sputter coating the lamella.
A thin platinum layer is added on top of all lamellae to increase lamella stability and reduce sample charging in the TEM microscope. Click Prepare For Sputtering in the xT Microscope Control software in the Cryo TEM Preparation tab.
Once the chamber pressure reaches 0.1 mbar, the Sputter button will appear and a sputter chamber will cover the shuttle, which has moved to a different chamber position.
Sputter for 5–7 s at 7 mA. Confirm sputtering is taking place by looking out for a glowing white light on the in-chamber camera.
Immediately after the sputtering is done, press Recover From Sputtering to return to high vacuum.
Unload the grids.
Cool down the loading station.
Quickly remove the shuttle from the FIB–SEM chamber and place it in the loading station using the transfer rod. Take care not to slam the shuttle on the bottom of the loading station.
Flip the shuttle 90° and move the grids very carefully to a grid box for clipped grids. Work carefully to avoid buildup of ice contamination on the grids.
Store the grid box under clean liquid nitrogen until ready for TEM imaging.
Warm up the FIB–SEM chamber to room temperature by removing the heat exchanger from the liquid nitrogen dewar and “sleep” the system.
Data analysis
FM/SEM image correlation
Correlation between FM and SEM images was performed using MATLAB R2021b (Natick, Massachusetts: The MathWorks Inc.) Briefly, sets of control pairs were chosen on both images based on recognizable features, and a similarity transformation matrix was determined using this point-to-point correspondence between FM and SEM images. The best transformation was determined using least squares minimization of a suitable cost function.
Quantification of imageable areas within lamellae
Quantification of imageable areas was performed using ImageJ software [US National Institutes of Health, Bethesda, Maryland, USA (Schindelin et al., 2012)]. Briefly, the TEM overview images of the lamellae were initially binned by a factor of 3 to allow for better visual differentiation of clean and contaminated regions within the lamellae. An initial global thresholding was performed to outline the major regions contaminated with thickest ice crystals (lowest gray values). The minor ice-contaminated regions were then manually selected using the free selection tool and the area corresponding to each region was calculated.
Reconstruction of the tomograms
Tilt series were motion corrected and CTF corrected using WARP (Tegunova and Cramer, 2019). Next, the tomogram was reconstructed using AreTomo (Zheng et al., 2022) and deconvolved and denoised using IsoNet (Liu et al., 2021). Visualization of the data was done using IMOD (Mastronarde and Hel, 2017).
Validation of protocol
Micropatterning of electron microscopy grids improves cell positioning and cell shape
Figure 3 displays the effect of photopatterning with the PRIMO system and the associated benefit on grid quality. Figure 3A displays a traditional, non-micropatterned grid. Note that the cells clump together leading to poor vitrification. Many cells are also positioned on top of the grid bars, making them inaccessible for thinning with the cryo FIB–SEM. Figure 3B shows an EM grid that was photopatterned. Cells are perfectly positioned in the center of the grid squares, leading to a significant increase in possible FIB milling sites, as indicated with the red arrows. Figure 3C reveals how cells can be deliberately aligned to cover all possible milling orientations. A modified pattern can be used to help reposition ROIs or organelles to an area of choice.
Micropatterning of electron microscopy grids can aid in ROI positioning to the center of the grid squares
In many cases, the cellular feature of interest will be scarce and may be located far away from the cell nucleus, which complicates reliable positioning of the ROI at the periphery of the cell.
Figure 4 illustrates how micropatterning can be used to help re-direct the ROI to the center of the grid squares, improving the ability to target these cellular features. In Figure 4A, a grid that did not undergo micropatterning is shown. Note that there are several cells in this grid square, with the largest portion of the cell body positioned on or near the grid bars. Most of the ROIs are not accessible due to their proximity to the edges of the grid squares or due to their proximity to the thinner regions of the cell near the plasma membrane. In Figure 4B, a simple, circular pattern was used to help re-position the cells, but not the organelles of interest. Here, the cell is positioned away from the grid square edges. The ROI that was labeled with mCherry mainly localizes to the cell periphery, making it hard to capture in the lamella, as indicated with a white, dotted line. In Figure 4C, a modified pattern was used to increase the number of ROIs that could be captured in a lamella. Most of the cells had these features in the cell periphery. Therefore, the interface between the two cells was directed towards the center of the grid square by the photopattern. The lamella could now be positioned at the cell periphery, increasing the likelihood that the ROI (in red dots) could be captured in a single lamella. Other patterns can be designed to specifically position other organelles or cellular features.
Figure 4. Micropatterning enables organelle positioning for ROIs located at the periphery of the cell. A. A typical EM grid that was incubated with RPE-1 cells. B. EM grid that was photopatterned with 20 μm diameter circles in the center of all grid squares before seeding RPE-1 cells on the grid. C. An EM grid that was photopatterned with two touching 20 μm diameter circular patterns in the center of the grid square. Possible milling sites are indicated with a white dotted line and the photopattern used is indicated in the upper left side of each image.
CERES ice shield reduces the ice contamination growth on the sample
Figure 5A and 5B display representative SEM images illustrating the ice thickness estimation procedure described in Section D1. The results of these measurements were used to quantify the rate of in-chamber ice growth. The graphs in Figure 5C and 5D display the average thickness of the ice layer at each measured time point determined in triplicates before and after installation of the CERES ice shield, respectively. Before installation, the measured ice layer thickness after 3 h was approximately 8.5 ± 4.2 nm. By contrast, extending the residence time of the grids inside the chamber after installation and measuring the ice layer thickness after 22 h yielded an approximate ice layer thickness of 10.8 ± 2.0 nm. Based on these results, the average ice growth rate was estimated as 3.0 nm/h and 0.6 nm/h before and after installation of the CERES ice shield, respectively. The differences correspond to a >5-fold reduction in the ice contamination in the presence of the CERES ice shield. Please note that this Aquilos2 cryo FIB–SEM instrument already underwent chamber vacuum improvements leading to a lower contamination rate prior to the installation of the CERES ice shield. Older, similar devices showed an ice growth rate around 50 nm/h, which will drop below 2.0 nm/h after installation of the CERES ice shield (Tacke et al., 2021)
Figure 5. Determination of in-chamber ice growth rates before and after the installation of the CERES ice shield. A/B. Representative SEM images of Quantifoil holes on support used to measure the thickness of the ice layer: top row images depict the first time point (T = 0 h) and bottom row images depict the last time point (T = 3 h pre-installation, T = 22 h post-installation). Top right magnified inset shows the measurement of layer thickness on the image. To determine ice thickness, the layer thickness difference to T = 0 h was used. C/D. Corresponding diagrams of ice thickness at each measured time point before and after installation of the CERES ice shield. Measurements were done in triplicate on different grid regions to generate an average. The average ice growth rate was estimated by linear fitting at 3.0 and 0.6 nm/h before and after installation.
The improved workflow using CERES ice shield and the METEOR system reduces ice contamination on lamellae and thereby greatly enhances sample throughput
Figure 6 depicts the impact of ROI confirmation with METEOR compared to using a stand-alone cryo-fluorescence microscope. Without METEOR, the batch-milled lamellae were transferred to a cryo-confocal microscope to confirm the presence of ROIs. We used a Zeiss LSM 800 upright confocal microscope equipped with a Linkam CMS196 cryo-stage for this work, but any cryo-capable fluorescence microscope can be used to image the lamella. The contamination build-up rate will vary across different microscope and cryo-stage configurations, as well as handling speed and room humidity. As shown in the overview TEM image as well as the higher magnification close-ups in Figure 6A, this extra transfer step leads to significant crystalline ice contamination in the form of many large (up to 1 μm) opaque ice crystals across the lamella, obscuring the potential candidate regions for tomogram acquisition. The additional transfer puts the sample in further danger of devitrification. However, assessing lamella quality using METEOR in the FIB chamber, as described in Section 5, eliminates this extra transfer step and thereby results in clean, frost-free lamellae as displayed in Figure 6B. To quantify the impact of METEOR on reducing crystalline ice contamination, TEM overview images of six lamellae without use of METEOR and nine lamellae with use of METEOR were screened for detecting the crystalline ice particles. Without using the integrated METEOR in the workflow, 30.2% of obscured area was determined in 668 μm2 of total screened area. With METEOR, 2.8% of obscured area was detected in 2200 μm2 of screened area. The presence of the CERES ice shield and the METEOR system enabled more intact lamellae to be milled (+329% increase in total surface area) and led to more usable area per session (+458% increase).
Figure 6. Extent of frost contamination on lamellae with and without in-chamber fluorescence detection. A. Overview TEM image of a batch-milled lamella where the confirmation of the ROI was performed with a separate cryo-confocal microscope. Due to the additional transfer step to the cryo-confocal microscope, ice crystals with different sizes and shapes (marked in white) have covered 25.4% of the shown lamella area. In the close-ups, the red arrow points toward an obscured potential ROI that otherwise could have been a candidate for tomogram acquisition. B. Overview TEM image of a batch-milled lamella where the presence of the ROI was confirmed within the vacuum chamber using METEOR. Eliminating the extra final transfer step to a separate fluorescence microscope led to clean lamella with 99.2% of imageable area (inset scale bars: 500 nm).
To validate the effectiveness of the upgraded workflow, the described procedure was tested on stable, immortalized human RPE-1 cells expressing mCherry-p62. p62/SQSTM1 (hereafter, p62) is a classic autophagy cargo receptor that binds poly-ubiquitinated cargo at its C-terminus and polymerizes into filaments at its N-terminus (Ciuffa et al. 2015; Jakobi et al. 2020). This polymerization results in phase separation showing droplets within the cell, thereby facilitating autophagic cargo concentration and segregation (Turco et al., 2021). The droplets are subsequently confined in a double membrane forming an autophagosome. Finally, the autophagosomes are transported to the lysosomes.
Figure 7 depicts the step-by-step targeted lamella milling process described in Section 3 of the procedure. Once the cell of interest was chosen based on the in-situ fluorescence signal provided by METEOR, as presented in Figure 7A–7C, the milling process was initiated. To avoid milling away the suitable ROIs, multiple checks were performed during the milling process. In Figure 7D–7F the checkpoint at 500 nm lamella thickness is shown as it confirmed the presence of potential ROIs, allowing for further thinning of the lamella. Once the lamella reached the desired thickness, a final check was performed by METEOR, as indicated in Figure 7G–7I, to ascertain that the lamella is a suitable candidate for tilt series acquisition. Correlations between SEM and FM images at different stages of cryo FIB milling are superimposed to monitor progress as displayed in Figure 8A–8C. This final fluorescence image was additionally correlated with a TEM overview image of the final lamella to accurately pinpoint the ROIs as shown in Figure 8D. The image correlation was used to record tilt series on the sites of interest in the cryo-TEM. Subsequently, the tilt series were reconstructed to yield tomograms and the presence of the ROI was confirmed by correlating the METEOR image to the final tomogram (Figure 9).
Figure 7. Targeted lamella milling on human RPE-1 cells using METEOR. A–C. Pre-milling status of the cell of interest imaged using SEM, FIB, and METEOR, respectively. Red arrows in SEM and FIB images point toward the same cell. D–F. SEM, FIB, and METEOR images corresponding to a checkpoint at 500 nm lamella thickness, respectively. The white arrowheads point towards fluorescence emitted from potential ROIs. G–I. SEM, FIB, and METEOR images corresponding to the final checkpoint, respectively. White boxes in (I) outline the fluorescent areas in the final lamella.
Figure 8. In-situ correlative cryo FIB milling on human RPE-1 cells. Superimposed images after correlation between (A) SEM and METEOR images prior to milling, (B) at 500 nm lamella thickness, and (C) at final lamella thickness. (D) The final fluorescence image of the lamella was additionally correlated with a TEM overview image to pinpoint the suitable locations for tomogram acquisition (white boxes).
Figure 9. Tomographic slice correlated to METEOR image. A. Image slice from a tomogram showing a lysosome. B. The corresponding fluorescent microscopy image from the METEOR system overlaid transparently with the tomographic slice (red: mCherry-p62; green: autofluorescent background signal, which often accompanies lysosomes).
General notes and troubleshooting
General notes
By combining PRIMO (Alvéole) with the CERES ice shield (Delmic) and the METEOR (Delmic), the productivity of our cryo-ET workflow improved by an order of magnitude compared to the traditional workflow (Table 1). First, by enhancing the grid quality using the PRIMO photopatterning system, we were able to improve the cell positioning by confining the cells to the center of the grid squares, thereby increasing the number of amenable cells for milling by at least 4-fold. Micropatterning also provides control over the cell shape and hence allows for repositioning of cellular features of interest, e.g., when they are located in the periphery of the cell. Subsequent addition of the CERES ice shield drastically reduced the rate of surface ice contamination build-up in the cryo FIB–SEM chamber. This contamination trap allowed for longer milling sessions, resulting in increased throughput. Finally, by including the METEOR in-chamber fluorescence microscope, we reduced the number of sample transfer steps between instruments, which are the main source of ice contamination. Additionally, the METEOR system allowed for frequent checking of the lamella to make sure that it still contained the ROI. At each intermediate checkpoint, when the ROI was no longer present in the lamella and was milled away, the polishing step on this lamella could be skipped and valuable microscope time could be saved for more suitable sites. Together, the three described integrated additions to the standard FIB–SEM instrumentation are of highly complementary value and improve the quality as well as the efficiency of lamella generation for in situ structural biology approaches.
Troubleshooting
Problem 1: No patterns observed and cells growing everywhere on the grid.
Solution 1: Glow discharging may have been inefficient. Increase the glow discharging duration or current. Make sure that you see liquid spreading out on the surface of the grids and not rounding up and rolling off the sides.
Solution 2: The anti-fouling layer/passivation was not carried out under the right conditions as described above. Increase PLL-g-PEG concentration or incubation duration.
Solution 3: Ensure the grid does not dry out while handling it. Drying the grids will cause the anti-fouling layer to collapse.
Solution 4. Reduce incubation time of the cells on the grid. When overgrowing cells on the grid, the cells will continue dividing and eventually spread to non-patterned areas.
Solution 5. Cells could be growing under the grid and not on top of the grid; move the grid to a clean dish with media and check cell distribution on the grid. Always ensure the grids are flat on the bottom of the dish before seeding the cells on top.
Solution 6: After micropatterning, remove the grids immediately from the PLPP photoinitiator solution and wash thoroughly. Extensive incubation after patterning can lead to formation of free radicals that can degrade the PLL-g-PEG layer.
Problem 2: No patterns observed and cells not growing anywhere on the grid.
Solution 1: Increase fibronectin concentration or incubation duration before seeding the cells.
Solution 2: Switch to another extracellular matrix protein such as vitronectin and see if cell growth is not observed.
Solution 3: Ensure the grid material is not toxic to your cells. For instance, copper grids leak ions in the media leading to cell death.
Problem 3: Extensive ice contamination is seen on the lamella despite the CERES ice shield.
Solution 1: Take extreme care while unloading the grid from the cryo FIB–SEM and loading the grid in the TEM. Use only clean liquid nitrogen, work in a low humidity room, work fast but accurately, and, finally, consider wearing a face mask while handling the grid.
Solution 2: Check the cryo FIB–SEM chamber vacuum. Normal wear and tear can lead to leaky tubing and degraded chamber vacuum values.
Problem 4: Extensive devitrification is seen on the lamellae imaged with the METEOR system.
Solution 1: Reduce the excitation light power and/or exposure time.
Solution 2: Always ensure all tools are fully cooled to liquid nitrogen temperatures before use, especially tweezers that are used to handle the grid directly.
Problem 5: No fluorescence is seen in the lamellae using the METEOR system despite the target being present in the lamella.
Solution 1: Increase the excitation power and/or exposure time and use camera binning.
Solution 2: Proteins with low abundance or faint tags may be hard to detect in the final ~150 nm thick lamella. Switch to a newer generation of brighter fluorophores, encode a triple tag, i.e., 3× eGFP instead of eGFP, or consider tagging another more abundant protein that will also localize to your ROI.
Acknowledgments
The authors gratefully acknowledge the electron microscopy training, imaging, and access time granted by the life science EM facility of the Ernst-Ruska Centre at Forschungszentrum Jülich. The authors gratefully acknowledge the computing time granted by the JARA Vergabegremium and provided on the JARA Partition part of the supercomputer JURECA at Forschungszentrum Jülich. Sabrina Berkamp was funded by the fellowship for postdoctoral researchers of the Alexander von Humboldt Foundation. The authors gratefully acknowledge Andreas Brech and Sebastian Schultz from Oslo University Hospital, Oslo, Norway for the cell line used in this Protocol. Parts of Figure 1 were created with BioRender (www.BioiRender.com). The micropatterning protocol was adapted from Toro-Nahuelpan et al. (2020).
Competing interests
M.S., C.J., and D.D. are employees of Delmic BV. A.C. and R.A.J. are employees of Alvéole. S. Berkamp. P.S., S.G., B.H., and C.S. declare no competing interests.
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4,902 | https://bio-protocol.org/en/bpdetail?id=4902&type=0 | # Bio-Protocol Content
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Headplate Installation and Craniotomy for Awake In Vivo Electrophysiological Recordings or Two-Photon Imaging of the Mouse Inferior Colliculus
BK Blom Kraakman
SS Sofja Solovjova
JB J. Gerard G. Borst
AW Aaron Benson Wong
Published: Vol 13, Iss 24, Dec 20, 2023
DOI: 10.21769/BioProtoc.4902 Views: 611
Reviewed by: Geoffrey C. Y. LauXiaoliang Zhao Anonymous reviewer(s)
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Bio-protocol journal peer-reviewed
Dec 20, 2023 | This version
Preprint
Sep 28, 2021
Original Research Article:
The authors used this protocol in eLIFE Oct 2019
Abstract
The inferior colliculus (IC) is an important processing center in the auditory system, which also receives non-auditory sensory input. The IC consists of several subnuclei whose functional role in (non-) auditory processing and plastic response properties are best approached by studying awake animals, preferably in a longitudinal fashion. The increasing use of mice in auditory research, the availability of genetic models, and the superficial location of the IC in the mouse have made it an attractive species for studying IC function. Here, we describe a protocol for exposing the mouse IC for up to a few weeks for in vivo imaging or electrophysiology in a stable manner. This method allows for a broader sampling of the IC while maintaining the brain surface in good quality and without reopening the craniotomy. Moreover, as it is adaptable for both electrophysiological recordings of the entire IC and imaging of the dorsal IC surface, it can be applied to answer a multitude of questions.
Key features
• A surgical protocol for long-term physiological recordings from the same or separate neuronal populations in the inferior colliculus.
• Optimized for awake in vivo experiments in the house mouse (Mus musculus).
Keywords: Mouse Inferior colliculus Multielectrode recordings Two-photon imaging Long-term recording Awake Surgery
Background
The inferior colliculus (IC) is an auditory midbrain nucleus consisting of several sub-nuclei (Oliver, 2005). It receives both auditory and non-auditory input from several ascending and descending pathways (Lesicko et al., 2020). Research into the functional role of the IC in auditory processing would benefit from studies in awake animals. Moreover, responses of IC neurons have shown plasticity from stimulus exposure (Cruces-Solís et al., 2018) and activation of corticofugal projections (Suga, 2012). Longitudinal measurements would aid in the study of the underlying mechanism of these plastic changes. Although the mouse IC lies superficially, the thickness of the overlying interparietal bone and the height of the surrounding structures (cerebellum, cerebral cortex) make it difficult to establish stable access for long-term physiology. Here, we describe a protocol optimized for exposing the IC for stable, multi-day recordings by means of a chronic cranial window. This method can be used to either expose the dorsal surface for in vivo two-photon imaging (Wong and Borst, 2019) or make the entire IC readily accessible for repeated electrophysiological recordings (van den Berg et al., 2023).
The stability provided by this protocol allows a larger sampling of neurons within the same animal, repeated measurements of the same neurons over days (imaging), or a study of long-term changes in the timespan of days and weeks. This protocol was utilized to show that the cortical IC contains a tonotopic map, albeit different in organization than the central nucleus of the IC, characterized by a reversal of the characteristic frequency gradient in the rostromedial–caudolateral direction (Wong and Borst, 2019). Moreover, the same study confirmed non-auditory (in the form of spontaneous movement related) signal processing in the IC cortex. Another adaptation of the protocol showed that mice are able to distinguish amplitude modulations (AM) up to 512 Hz of an auditory stimulus. The set of IC neurons with the largest change in firing rate in response to this AM change are sufficient to explain the behavioral detection threshold (van den Berg et al., 2023).
The main advantage of this method is the possibility to gather data at multiple time points without having to reopen the craniotomy. This gives the opportunity to do multiple recordings in the same region over an extended period of time without implanting a recording probe or reopening the craniotomy, as well as recording at multiple sites to get a completer sample of the IC neuronal population. By sealing the craniotomy with a low viscosity silicone gel (Jackson and Muthuswamy, 2008), the risk of infection and amount of tissue regrowth are reduced. As a result, the brain surface is kept in better quality for measurements. A limitation is that a well is required to hold the silicone gel in place, which limits the angle of approach of a recording probe. The cranial window construct described here for chronic imaging provides the depth needed for mechanical stability while limiting the amount of glass in the light path. An inherent limitation of two-photon imaging is the limited depth of the imaging plane (Helmchen and Denk, 2005). Given the anatomical shape of the mouse IC, the imaging will be limited to the dorsocaudal part of dorsal and lateral cortices (Wong and Borst, 2019).
Materials and reagents
Reagents
Superglue (e.g., UHU, catalog number: 62687)
Dental etching gel Etch Rite (38% phosphoric acid) (Pulpdent, catalog number: 185120)
Dental adhesive primers OptiBond FL 1 Prime (Kerr, catalog number: 25881E)
Dental adhesive OptiBond FL 2 Adhesive (Kerr, catalog number: 25882E)
Standard dental composite Charisma A1 (Heraeus, catalog number: 66056076)
Flowable dental composite Charisma Flow A1 (Heraeus, catalog number: 66015495)
Isoflurane
70% ethanol
0.9% NaCl (Baxter, catalog number: TKF7124)
Eye ointment (e.g., Alcon, Duratears Z, catalog number: 10308)
Silicone gel sealant (Dura-Gel, Cambridge NeuroTech, 2 components, catalog number: Dura-Gel or equivalent: DOWSILTM 3–4680 Silicone Gel Kit, DOW, catalog number: 3-4680)
Lidocaine spray, 100 mg/mL (e.g., Aspen, Xylocaine spray)
Buprenorphine (e.g., Dechra, Bupredine Multidose 0.3 mg/mL)
Carprofen (e.g., ASTfarma, Carporal 50 mg/mL)
15% Mannitol (Sigma, catalog number: M4125-100g)
Laboratory supplies
Drill head (Ø 0.3 mm) (Komet Dental, catalog number: H71 104 003)
Fine disposable micro applicators (FHS Dental, catalog number:18-902T)
Sugi Sponge Points (Kettenbach GmbH & Co. KG, catalog number: 31603)
Cotton swabs
Haemostatic gelatin sponge (Spongostan Standard, Ferrosan Medical Devices, catalog number: MS0006)
Paper points (Henry Schein, catalog number: 9001217)
Clean round headplate (manufactured in-house, Figure 1A) with the following dimensions: 1 mm thick, 10 mm diameter outer ring, 7 mm diameter inner ring, 18 mm wing to wing. Total weight: 500 mg
Craniotomy cover (manufactured manually, Figure 1B) with roughly the following dimensions: ~10 mm × 15 mm for the metal wire looped around the headplate, covered by either tape or hard plastic of ~70 mm2
Glass window construct
Position custom steel rings for the cranial window upside down.
Using a paper point, apply a small amount of optical adhesive (NOA 68, Norland Products, catalog number: 25882E) to the bottom rim of the steel ring.
Place a cover glass (CS-3R-0 Ø 3 mm #0 thickness, Warner Instruments, catalog number: 64-0726) on the bottom side of the steel ring (Figure 1C).
Apply a small pressure with the forceps and make sure the optical adhesive completely connects the steel ring and the cover glass, i.e., without air gaps.
Cure the optical adhesive under a UV lamp according to manufacturer’s specifications (3 min under 100 W).
Ground pin construct
Cut the silver wire (Advent, catalog number: AG549311) to ~3 cm and strip off the Teflon coating.
Remove the plastic from the PCB pin and trim the end.
For extra mechanical stability, the silver wire could be looped with both ends of the wire soldered together onto the pin.
Chlorinate the tip of the wire with bleach for a few minutes and then rinse the bleach off with demi water (Figure 1D).
Figure 1. In-house made materials. Shown are the (A) headplate, (B) craniotomy cover, (C) glass window construct, and (D) ground wire construct. Scale bars correspond to 5 mm.
Equipment
Scalpel holder with #10 scalpel blade
Coarse forceps
Fine forceps (e.g., Fine Science Tools, catalog number: 11254-20)
Fine scissors (e.g., Fine Science Tools, catalog number: 15024-10)
Micro curette (Fine Science Tools, catalog number: 10082-15)
Composite instrument (e.g., Hu-Friedy, catalog number: CI0135)
Coverslip forceps (Fine Science Tools, catalog number: 11251-33)
Headplate holder (manufactured in-house)
Heat pad (e.g., FHC, Bowdoinham, catalog number: 40-90-8C)
Dental drill (e.g., Foredom Micromotor, H.MH-170)
LED composite curer (e.g., GDT, catalog number: 077340)
Procedure
Headplate installation
Anaesthetize the mouse with isoflurane (~0.8 L/min, 5%).
Once unconscious, remove mouse from incubation chamber and put under face mask and on a heating pad.
Switch to ~0.8 L/min, 2% isoflurane, and adjust as necessary throughout the surgery.
Apply eye ointment.
Inject carprofen (5 mg/kg) and buprenorphine (0.05 mg/kg) subcutaneously.
When placing a cranial window: Inject 15% D-mannitol (2,000 mg/kg) intraperitoneally.
Straighten the head by supporting the neck with a roll of paper tissue or by using ear bars.
Wet the fur on the scalp using lidocaine spray and shave the scalp with a scalpel.
Note: Be careful to keep the lidocaine from flowing to the nose of the animal, which will block its airway.
Use 70% ethanol to sterilize skin, spray lidocaine on clean scalpel, and cut open the skin from 1–2 mm caudal to the occipital ridge to the middle of two eyes.
Spray additional lidocaine on the incision site and exposed periosteum.
Pull skin laterally to expose the skull; cut the periosteum between the skin and the skull following the perimeter of the incision (see Figure 2A).
Locate the anatomical landmarks Bregma and Lambda on the exposed skull.
Make a small incision caudally along the midline to separate the thin membranous muscles at the dorsal neck region. Push away the muscles laterally to expose the caudal edge of the skull (occipital ridge) and the attached muscles (splenius capitis and semispinalis capitis muscles).
Detach the muscles from the occipital bone by cutting with the fine scissors or scrapping with forceps (see Figure 2B). Avoid damaging the muscle by keeping the incisions as close to the occipital bone as possible.
Figure 2. Headplate installation. Images show procedure steps corresponding to (A) exposure of the skull, (B) detachment of the muscles, and (C) preparation of the skull surface for (D) headplate placement roughly over lambda.
Scratch the bones with a curette or the tip of a scalpel to remove periosteum and residual attached muscle. The exposed area should cover the dorsal surface of the skull from the rostral side to Bregma and the caudal edge of the skull and extend laterally.
Clean the surface with 70% ethanol. Let the bone air dry briefly.
Insert four paper points between the occipital bone and skin and underneath the muscle on the interparietal bone. This makes the area accessible for etching and prevents damage to skin and muscles (see Figure 2C).
Apply etching gel on the skull and let it sit for 15–30 s until the bone surface becomes lighter and turns matte. Wipe away the etching gel with a cotton swab wet with saline until no etching gel is left on the surface of the skull.
Apply bonding primer (OptiBond prime) to the skull using a micro applicator and let it air dry.
Note: Make sure no excess amount of primer remains on the surface, i.e., it should not look too watery but can be a bit shiny. However, a desiccated over-dried surface will also hamper proper bonding in the next steps.
Apply bonding adhesive (OptiBond adhesive) to the entire exposed skull with a micro applicator or a paper point in a scratching motion while applying some pressure.
Note: The scratching action ensures that the adhesive gets into small pores of the bone and prevents these pores from closing up.
Cure adhesive with UV light (see General note 1).
Apply bonding adhesive (OptiBond adhesive) to the headplate using the same micro applicator.
Position the headplate on the skull, so that the opening is slightly caudal to Lambda. Make sure the headplate is as close to the skull as possible. Too much adhesive between the headplate and the skull may prevent short working distance objectives from reaching the brain.
Note: Positioning depends on the dimensions of the headplate and the purpose of experiments. A well-centered (lateral) headplate eases head fixation for chronic measurements and helps with standardization of position.
Cure the headplate to the skull with UV light. Ensure that the headplate does not change position while curing.
Secure the headplate with dental composite (see General notes 2 and 3). Make sure all gaps between the headplate and the skull are filled (see Figure 2D).
Ensure the dental composite has solid contact with the back part of the skull (occipital bone) in addition to the top parts of the skull (parietal and interparietal bones). This gives extra strength to the skull, prevents accidental breaking during head fixation, and helps reduce relative motion between the skull and the brain during recordings.
Fill up the rostral part of the skull with dental composite, with a bit grabbing onto the headplate. Be careful not to overdo it as this will increase the imaging distance from the objective.
Close up the opening by attaching any loose skin to the caudal side of the headplate with the superglue.
Pause point: If desired, the craniotomy steps (sections B–D) can be done in a second surgery at a later date. In that case, apply and cure a thin layer of flowable dental composite on the exposed skull to protect the bone and then proceed to section E for recovery.
Craniotomy
Use landmarks or atlas coordinates to locate the skull region underneath which the IC is expected to be.
Using a fine drill, sequentially thin away the skull (see General notes 4 and 5). Remove any bone debris.
Once the vascularized middle layer has been excavated, apply a bit of saline to the bone. At this point, the bone should be transparent and mildly flexible once pressured with forceps or the drill tip.
At this point, the IC is roughly visible. Use the cerebellum and transverse sinus as landmarks. The cerebellum tends to have small blood vessels running in the sagittal direction.
For cranial window installation (e.g., steel ring constructs): check the position of the IC and extend the size of the craniotomy around it to accommodate the window. This is best done by first marking the intended extent of the craniotomy by drilling with the aid of a glass coverslip glued to a paper point. Because of the skull thickness, add an extra margin of approximately 0.3 mm at the skull surface to ensure the bottom of the craniotomy will fit the cranial window.
Continue to thin the bone above the IC until it becomes membranous. The created funnel should extend rostro-laterally towards the sinus and a bit caudo-medially towards the cerebellum.
Once the intended area is thinned, remove the debris and rinse the surface with saline. Make sure that at this point any bleeding has been stopped to ensure a blood-free brain surface when the bone is removed.
Apply saline on the bone. Using a pair of fine forceps, very carefully chip away the bone above the brain starting from the cerebellum side (see General note 6).
The IC should be rather white and have a shiny surface, compared to the more gray/dark tone of the cerebellum. The dura mater should be intact at this point, forming a continuous surface on top of the IC and the cerebellum.
Note: Continue to either section C or D; the protocol is not compatible to combine both.
Variant 1: cranial window for in vivo two-photon calcium imaging
For pre-prepared cranial window constructs (e.g., steel ring constructs), the craniotomy should already fit the size of the cranial window. If necessary, expand the craniotomy and avoid debris and bone shards on the brain surface.
Fit the cranial window within the craniotomy (see General note 7).
Apply pressure and hold the window in place. Apply a tiny drop of superglue with the tip of a paper point to 2–3 contact points between the rim of the window and the bone.
Proceed to secure the cranial window with dental adhesive or cement (see Figure 3). Applying pressure to the cranial window is important to prevent growth of connective tissue between the glass window and the brain. Rule of thumb: if you think you have pressed too much, you are good. Alternatively, the whole window can be fastened with superglue for a removable window.
Figure 3. Cranial window for in vivo two-photon calcium imaging. Illustrated is (A) the two-photon calcium imaging setup previously used and (B) the final result of the installed cranial window over the left inferior colliculus (IC). SC, superior colliculus; Cb, cerebellum. Figure reproduced from Wong and Borst (2019) under a CC-BY license.
When applicable: Close the lid of the headplate once the glue has dried.
Variant 2: craniotomy for in vivo electrophysiological recordings
Apply bonding primer to the edges of the craniotomy using a paper point and let it briefly air dry.
Apply bonding adhesive to the craniotomy funnel using a paper point and cure it with UV.
Create a well of dental composite around the craniotomy and cure with UV light.
Secure the ground pin outside or on the edge of the headplate with Charisma. Gently slide the wire in-between the brain and the skull (see General note 9).
Fill the well with the silicone gel. Remove bubbles and debris from the gel using fine forceps.
Allow the silicone gel to set for 30 min.
Secure the ground construct with dental composite, leaving the pin opening exposed. Form a sheet of dental composite that covers the ground construct and the rest of the exposed skull (see General note 2, Figure 4A).
Make a cover by looping wire around the wings of the headplate (see Figure 4B).
Figure 4.Craniotomy for in vivo electrophysiology recordings. (A, B) Craniotomy over the inferior colliculus (IC) with silicone gel sealant. (C) Schematic overview of a vertical section through the preparation illustrating the Charisma barrier necessary to hold the silicone gel sealant and the position of the ground pin. The secured chlorinated silver wire is placed at the edge of the craniotomy and slipped in between the skull and the brain. Scale bars correspond to 1 mm.
Recovery from surgery
When using D-mannitol, inject saline intraperitoneally at a volume equal to previously injected D-mannitol.
Stop isoflurane and allow the mouse to wake up.
Wait for the first sign of spontaneous movements and transfer the animal to its cage under a heat lamp or on a heating pad.
For electrophysiological recordings, allow animals to recover for 3–4 days before performing subsequent awake experiments (Figure 5). For in vivo two-photon calcium imaging, imaging can start after 3–4 days of recovery. However, for stable, repeated long-term recordings, it is advisable to wait for two weeks after the surgery before imaging to allow the brain tissue to adapt and stabilize mechanically.
Figure 5. Head fixation and awake electrophysiological recordings. During electrophysiological recordings, mice are head fixed (A) before the recording electrode can be lowered into the craniotomy and the ground pin connected (B). To protect the gel sealant, the craniotomy is covered (C).
Validation of protocol
The protocol as described was used in previous works; see van den Berg et al. (2023) for its implementation in in vivo electrophysiology experiments and Wong and Borst (2019) for in vivo imaging application.
In van den Berg et al. (2023), we performed electrophysiological recordings using a high-density acute silicon probe single-unit recordings (see Figure 1B and sections “Surgery” and “Electrophysiological Recordings” from the materials and methods section in the cited article). The probe was mounted on a 3-axis motorized micromanipulator (Luigs-Neumann Mini-25 controlled by a SM-8 controller) and inserted perpendicular to the brain surface. The probe was connected to a head stage (RHD 64-channel headstage, part C3315, Intan Technologies) and acquisition board (RHD USB-interface board, part C3100, Intan Technologies, Los Angeles, CA). The electrophysiological signal was sampled at 30 kHz using the supplied recording GUI (RHD data acquisition GUI, Intan Technologies). At the end of the experiment, the probe was slowly retracted from the craniotomy and cleaned by placing the tip in protease-containing detergent (e.g., 1% Tergazyme) for 1 h followed by 1 h in ddH2O. This protocol was used for the awake recording data set in van den Berg et al. (2023), which amounted to 99 well-isolated single units recorded from four animals across recording sessions spanning between 4 and 13 days.
In Wong and Borst (2019), we performed in vivo two-photon imaging using a custom-built two-photon microscope with a 20× water-immersion objective (LUMPlanFI/IR, 20×, NA: 0.95; Olympus Corporation) (see Figure 1, “surgery” and “Two-photon imaging” from the materials and methods section in the cited article). Excitation light (920 nm) was provided by a MaiTai Ti:Sapphire laser (Spectra Physics Lasers, Mountain View). Images (256 × 128 pixels) were collected at 9 Hz (2 ms/pixel; 1–2 mm/pixel). In addition, this protocol has been used in our lab to measure neuronal activity in the IC for an extended period of time, as shown by pilot data in Figure 6 performed in a GCaMP6s-expressing mouse line (GP4.3; Chen et al., 2013).
Figure 6. Long-term imaging of the same inferior colliculus (IC) neurons. (A) Average fluorescence images of a region of interest in the dorsal IC. (B) Frequency response areas of the neuron marked by the yellow arrow in the corresponding session. Each subplot shows the average ΔF/Fb to a stimulus of the specified frequency and intensity. Background color shows average Pearson’s correlation among repetitions, indicating consistency of response (see Geis et al., 2011). Vertical scale bars in the right panel indicate 1 Fb. Horizontal scale bar is 1 s.
General notes and troubleshooting
General notes
Illumination from the surgical light may also cure the adhesive and dental composite (Charisma). Turn down the light intensity or block it with orange (UV) filters if it tends to harden too quickly while handling.
The dental composite used comes in two viscosities; use standard (paste-like) version for larger surfaces and flowable, low-viscosity version to fill up any small gaps left. Cure with UV light in between layers.
Clean tools with 70% ethanol before handling dental composite to prevent it from sticking too strongly to the tools. Preferably, use smooth, unscratched parts of the tools.
The bone above the IC (interparietal bone) consists of three layers: the middle layer is more vascularized, while superficial and deep layers are relatively vessel-free.
Small bleedings from the bone may be stopped by briefly drilling towards the source; otherwise, applying slight pressure helps to stop bleeding. Stop larger bleedings by applying a small piece of pre-wet Spongostan for a brief period (one to a few minutes) and then remove it slowly and gently with plenty of saline.
When removing the bone, working in solution helps prevent damage to the dura mater. For larger pieces that adhere strongly to dura, use two pairs of fine forceps: pick up a piece of bone on one side with the bent forceps and very slowly lift it up bit by bit like opening a rooftop trapdoor. The other fine forceps can be used to gently detach the bone from the connective tissue in a way similar to blunt dissection techniques.
For prepared cranial windows constructs (e.g., double glass, steel rings), make sure the cranial window fits into the opening before actually opening up the bone. This will avoid having to drill or chip away more bone at the edge later on, which may add bone shards on the brain surface and interfere with imaging.
A blood-free brain surface is highly beneficial to imaging. Try to prevent the formation of blood clots by promptly rinsing away any blood on IC surface. Try to remove any clotting blood with forceps before putting the glass window in.
During electrophysiological recordings, standard measures should be taken to reduce electrical noise such as proper grounding and shielding from electrical interference. For this preparation, the proximity of the ground wire to the recording site reduces impedance in general. Securely cementing the ground wire helps to reduce motion artefacts. The use of Ag/AgCl electrodes reduces offset drifts and makes this protocol also suitable for patch-clamp type recordings.
Daily maintenance of the imaging glass window involves rinsing the glass surface with saline and removing it with a Sugi or suction. In case of dirt or dried up fluid that is difficult to remove, a gentle scratch with fine forceps in saline can help dislodge it. A deterioration in the quality of view under the window, i.e., between the glass and brain, is mostly due to a (re-)growth of tissue between the glass and the brain. There is no good, practical way to reverse this process once happened. Thus, it is important to minimize the chance by making sure that no bone shards remain on the brain surface and applying enough pressure while fixing the cranial window. See also problems 2 and 3 under Troubleshooting.
Troubleshooting
Problem 1: Headplate detaches from the skull after a few days or upon head fixation.
Possible cause #1: Skull became weak due to immune response.
Solution: Make sure the procedure is performed as cleanly as possible. Sterilize all tools and disinfect skin and fur of the animal around the incision. If desired, dexamethasone can be given to the animal at/after the surgery to reduce immune reaction.
Possible cause #2: Adhesive did not cure properly.
Solution: Make sure the skull surface is dry. Make sure the adhesive and primer have not expired. Make sure the adhesive or primer were not contaminated by other chemicals (e.g., ethanol).
Possible cause #3: Poor contact between the dental composite and the skull.
Solution: To prevent this, make sure that, when placing the headplate, the skull surface is dry and any tissue is removed. A headplate with small holes in the top can help to fill up the space between the headplate and the skull with the low viscosity composite. When securing the headplate with dental composite, pay extra attention to securing the back of the skull.
Problem 2: Significant (re-)growth of soft tissue between brain surface and cranial window, impairing imaging.
Possible cause #1: Not enough pressure between glass window and brain surface.
Solution: Make sure sufficient pressure was applied when securing the window. If necessary, thin the bone where the outer rim of the steel ring will sit to allow a deeper insertion of the window construct.
Possible cause #2: Bone growth between glass window and brain surface.
Solution: See Problem 3.
Problem 3: Growth of bone between brain surface and cranial window.
Possible cause: Small bone shards between cranial window and brain surface began to grow and enlarge.
Solution: Clear the brain surface of any tiny bone shards before installing cranial window.
Problem 4: Detachment of ground pin from the animal.
Possible cause: The animal could have bumped into something in the home cage that made the pin detach. Any liquid or air pockets make the attachment of the dental composite less secure.
Solution: In general, take care to position the ground pin as to not protrude out of the headplate too much/at all. Make sure a sufficient amount of dental composite surrounds the ground pin. To remedy an animal with a detached pin, check the state of the craniotomy and confirm that brain was not damaged by the event. In that case, attach a new pin and electrically connect it to the end of the remaining silver wire using conductive paint such as RS PRO Conductive Lacquer.
Problem 5: Cloudy glass window.
Possible cause: Vapor of superglue will condense on the clean surface of the glass window and make it foggy.
Solution: This is a problem that occurs often. The vapor is very easily removed by gentle scratching with forceps once the window is fixed. The removal can be done (preferably) at the first imaging session.
Acknowledgments
This research was funded by the EU Marie Skłodowska-Curie Innovative Training Network LISTEN (H2020-MSCA-ITN #722098) to JGGB, a ZonMW TOP grant (#91218033) to JGGB; and an EU Marie Skłodowska-Curie Individual Fellowship (660157-OPTIMAPIC) to ABW and a Dutch Research Council (NWO) VIDI project (VI.Vidi.213.198) to ABW. This protocol was used in Wong and Borst (2019; DOI: 10.7554/elife.49091) and van den Berg et al. (2023; DOI: 10.1152/jn.00048.2023).
Competing interests
The authors declare no competing interests.
Ethical considerations
All animal experiments were performed in accordance with institutional, national, and European ethical guidelines and legislation for laboratory animals as overseen by the Animal Welfare Board of the Erasmus MC. The studies carried out were approved by the animal ethical committee of the Erasmus MC (Instantie voor Dierenwelzijn; IvD) and the national authority (Centrale Commissie Dierproeven, The Hague, The Netherlands) as required by Dutch law.
References
Cruces-Solís, H., Jing, Z., Babaev, O., Rubin, J., Gür, B., Krueger-Burg, D., Strenzke, N. and de Hoz, L. (2018). Auditory midbrain coding of statistical learning that results from discontinuous sensory stimulation. PLoS Biol. 16(7): e2005114.
Chen, T.-W., Wardill, T. J., Sun, Y., Pulver, S. R., Renninger, S. L., Baohan, A., Schreiter, E. R., Kerr, R. A., Orger, M. B., Jayaraman, V., et al. (2013). Ultrasensitive fluorescent proteins for imaging neuronal activity. Nature 499: 295–300.
Geis, H. A. P., van der Heijden, M. and Borst, J. G. G. (2011). Subcortical input heterogeneity in the mouse inferior colliculus. J. Physiol. 589(16): 3955–3967.
Helmchen, F. and Denk, W. (2005). Deep tissue two-photon microscopy. Nat. Methods 2(12): 932–940.
Jackson, N. and Muthuswamy, J. (2008). Artificial dural sealant that allows multiple penetrations of implantable brain probes. J. Neurosci. Methods 171(1): 147–152.
Lesicko, A. M., Sons, S. K. and Llano, D. A. (2020). Circuit Mechanisms Underlying the Segregation and Integration of Parallel Processing Streams in the Inferior Colliculus. J. Neurosci. 40(33): 6328–6344.
Oliver, D. L. (2005). Neuronal Organization in the Inferior Colliculus. The Inferior Colliculus (pp. 69–114). In: Winer, J. A. and Schreiner, C. E. (Eds.). New York: Springer-Verlag.
Suga, N. (2012). Tuning shifts of the auditory system by corticocortical and corticofugal projections and conditioning. Neurosci. Biobehav. Rev. 36(2): 969–988.
van den Berg, M. M., Busscher, E., Borst, J. G. G. and Wong, A. B. (2023). Neuronal responses in mouse inferior colliculus correlate with behavioral detection of amplitude-modulated sound. J. Neurophysiol. 130(3): 524–546.
Wong, A. B. and Borst, J. G. G. (2019). Tonotopic and non-auditory organization of the mouse dorsal inferior colliculus revealed by two-photon imaging. eLife 8: e49091.
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4,903 | https://bio-protocol.org/en/bpdetail?id=4903&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
In vitro Assessment of Efferocytic Capacity of Human Macrophages Using Flow Cytometry
AS Ana C.G. Salina *
MF Marlon Fortes-Rocha *
LC Larissa D. Cunha
(*contributed equally to this work)
Published: Vol 13, Iss 24, Dec 20, 2023
DOI: 10.21769/BioProtoc.4903 Views: 2819
Reviewed by: Dipak Kumar PoriaNavnita Dutta Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Jun 2022
Abstract
Clearance of dying cells, named efferocytosis, is a pivotal function of professional phagocytes that impedes the accumulation of cell debris. Efferocytosis can be experimentally assessed by differentially tagging the target cells and professional phagocytes and analyzing by cell imaging or flow cytometry. Here, we describe an assay to evaluate the uptake of apoptotic cells (ACs) by human macrophages in vitro by labeling the different cells with commercially available dyes and analysis by flow cytometry. We detail the methods to prepare and label human macrophages and apoptotic lymphocytes and the in vitro approach to determine AC uptake. This protocol is based on previously published literature and allows for in vitro modeling of the efficiency of AC engulfment during continual efferocytosis process. Also, it can be modified to evaluate the clearance of different cell types by diverse professional phagocytes.
Graphical overview
Keywords: THP-1-derived macrophage PBMC-derived macrophages Efferocytosis Apoptotic cells Flow cytometry
Background
The uptake of apoptotic cells (ACs) is a signal that modulates macrophage metabolism and gene expression and ultimately shapes their functional programming (Boada-Romero et al., 2020; Schilperoort et al., 2023). Efferocytic activity is coupled to other environmental cues, such as the cytokine milieu and the presence of pathogens, to promote macrophage inflammatory or anti-inflammatory responses and their capacity to induce efficient tissue repair (Torchinsky et al., 2009; Bosurgi et al., 2017; McCubbrey et al., 2022; Salina et al., 2022). Of note, the latter requires efficient continuous uptake of apoptotic cells; thus, one of the consequences of efferocytosis is to promote a positive feedback loop for sequential corpse engulfment (Miyanishi et al., 2007; Park et al., 2011; Yurdagul et al., 2020). The uptake of a single AC or their continual uptake can be assessed by fluorescent cell labeling and analyzed by fluorescence microscopy or flow cytometry (Yurdagul et al., 2020; Gerlach et al., 2021; Salina et al., 2022; Wang et al., 2023). Here, we provide a detailed protocol on how to prepare human macrophages (either from THP-1 cell lines or peripheral blood samples) and evaluate the uptake of apoptotic Jurkat cells by macrophages in vitro using differential fluorescent labeling of cells and data acquisition by flow cytometry. We label each cell population of interest (phagocytes, AC target cells) and show how to identify engulfing populations. This protocol provides a simple approach to probe AC uptake and allows robust quantitative evaluation of continual efferocytic capacity. Of note, variations in cell culture conditions or donor samples may influence macrophage behavior and affect assay results. While designed to assess single and continual efferocytosis in vitro, this protocol could be adapted to facilitate probing AC uptake in challenging models in vivo by pre-labeling target cells and surface staining specific phagocytic populations of interest.
Materials and reagents
150 cm2 cell culture flasks (Corning, catalog number: 430825)
Jurkat cells, clone E6-1 (ATCC, catalog number: TIB-152)
5 mL sterile serological pipette (Falcon, Corning, catalog number: 356543)
10 mL sterile serological pipette (Falcon, Corning, catalog number: 357551)
25 mL sterile serological pipette (Falcon, Corning, catalog number: 357525)
50 mL sterile polypropylene conical tube (Corning, catalog number: SCT-50ML-25-S)
24-well tissue culture plate (Corning, Costar, catalog number: 3524)
5 mL syringe (BD Bioscience, catalog number: 990317)
Hypodermic needle 23 G × 1 (BD Bioscience, catalog number: 300388)
Disposable Pasteur pipette
28 mm diameter syringe filter, 0.2 µm pore PES membrane (Corning, catalog number: 431229)
Cell scraper (Corning, Falcon, catalog number: 353085)
5 mL non-sterile round-bottom polystyrene test tube (FACS tube) (Falcon, Corning, catalog number: 352008)
THP-1 cells (ATCC, catalog number: TIB-202)
Roswell Park Memorial Institute (RPMI) 1640 medium (Thermo Fisher Scientific, Gibco, catalog number: 22400-089)
Fetal bovine serum (FBS) (Thermo Fisher Scientific, Gibco, catalog number: 12657-029)
GlutaMAXTM supplement (Thermo Fisher Scientific, Gibco, catalog number: 35050-061)
Penicillin-streptomycin (10,000 U/mL) (Thermo Fisher Scientific, Gibco, catalog number: 15140-122)
Phosphate buffer saline (PBS) (Thermo Fisher Scientific, Gibco, catalog number: 10010-023)
CellTraceTM Far Red (Thermo Fisher Scientific, Invitrogen, catalog number: C34572)
pHrodoTM Red AM intracellular pH indicator dye (Thermo Fisher Scientific, Invitrogen, catalog number: P35372)
Tissue culture dish (100 mm × 20 mm) (Corning, catalog number: 430591)
Phorbol 12-myristate 13-acetate (PMA) (InvivoGen, catalog number: Tlrl-pma)
Leukocyte reduction system (LRS) cone
Histopaque®-1077 (Sigma-Aldrich, catalog number: 10771- 500ML)
Ammonium chloride (NH4Cl) (CAS 12125-02-9) (Mallinckrodt, catalog number: 3348)
Potassium bicarbonate (KHCO3) (CAS 298-14-6) (Sigma-Aldrich, catalog number: 237205)
Ethylenediaminetetraacetic acid (EDTA) (CAS 60-00-4) (Amresco, catalog number: 0322-500G)
Paraformaldehyde 20% aqueous solution EM grade (Electron Microscopy Sciences, catalog number: 15713)
CellTraceTM CFSE (Invitrogen, Thermo Fisher Scientific, catalog number: C34554)
CellTraceTM Violet (Invitrogen, Thermo Fisher Scientific, catalog number: C34571)
Human serum from human male AB plasma (human serum), USA origin, sterile-filtered (Sigma-Aldrich, catalog number: H4522-100ml)
Zombie NIR Fixable Viability (BioLegend, catalog number: 423106)
ACK lysis buffer (see Recipes)
Recipes
ACK lysis Buffer
0.15 M NH4Cl
10 mM KHCO3
0.1 mM EDTA
Solvent: deionized H2O
Adjust the pH solution to 7.3.
Filter the solution using a 0.2 μm sterile filter and keep it at 4 °C.
Equipment
Hemostatic forceps
Surgical scissor
Centrifuge (Thermo Scientific, Heraeus Megafuge 40R, catalog number: 50119920)
UV Crosslinker (Fisher Scientific, model: 234100, catalog number: 13-245-221)
Automated cell counter (Life Technologies, model: Countless II FL, catalog number: AMQAF1000) or cell counting chamber
Flow cytometer (BD Biosciences, FACS Verse, catalog number: 651153)
Software
FlowJo v10.8.0 (FlowJo, LLC, www.flowjo.com)
Prism 8 (GraphPad Software, Inc., www.graphpad.com)
Procedure
Note: Suggested conditions are sufficient to prepare one 24-well tissue culture plate of macrophages and AC for the efferocytosis assay, including flow cytometer compensation controls and fluorescence-minus-one controls for cell gating at Data analysis.
THP-1-derived macrophage differentiation
Note: THP-1 cells are maintained in 175 cm2 cell culture flasks in RPMI culture medium supplemented with 10% of FBS, 1% GlutaMAXTM supplement, and 1% penicillin-streptomycin medium (RPMIc), at 37 °C and 5% CO2 atmosphere. The cell culture is split, and the medium is replenished every other day. Whenever mentioned below, cell transferring and resuspension were performed using disposable serological pipettes. All media and buffers were pre-warmed prior to use.
Collect the cell suspension from the flask, transfer to a 50 mL conical tube, and pellet them by centrifugation (400× g for 5 min at room temperature).
Dump the medium, resuspend the pelleted cells in 5 mL of PBS, and count them.
Transfer 2.4 × 107 THP-1 cells to a clear 50 mL conical tube (Tube A) and centrifuge them.
Transfer at least 4.0 × 106 THP-1 cells to another clear 50 mL tube (Tube B). Centrifuge them, dump the media, and resuspend the cell pellet in fresh RPMIc at 4.0 × 106 cells/mL. These cells will be spared to prepare compensation controls (unstained and viability dye single-color controls) and should be kept at 37 °C and 5% CO2 until seeding.
Dump the medium in tube A and resuspend the THP-1 cells in 2.4 mL of PBS (1.0 × 107 cells/mL).
Proceed to cell labeling with CellTraceTM Violet or CFSE (working solution concentration: 5 μM) following manufacturer's instructions (Note 1).
Add the labeling reagent at 1:1,000 to obtain adequate working concentration, mix by flicking the tube, and incubate for 20 min protected from light in a 37 °C water bath.
Add five times the original staining volume of fresh pre-warmed cell media to the tube and incubate for 5 min at 37 °C.
Centrifuge the tube (400× g for 5 min).
Dump the medium and rinse the cell pellet with 10 mL of fresh pre-warmed cell media.
Following the washout step, resuspend the THP-1 cell pellet in 6 mL of fresh RPMIc (4.0 × 106 cells/mL).
Seed 250 µL (1.0 × 106 cells) of the cell suspension in tubes A and B per well of a 24-well cell culture plate.
Prepare a solution of PMA at 100 ng/mL using RPMIc as diluent.
Add 250 µL of PMA solution to each well (final volume 500 μL, final concentration 50 ng/mL).
Incubate the plate for 24 h at 37 °C and 5% CO2.
Aspirate all the medium, replenish with 500 μL of fresh RPMIc, and incubate the plate for another 24 h at 37 °C and 5% CO2.
Macrophages are ready for efferocytosis assay.
PBMC-derived macrophage differentiation
Note: This protocol uses peripheral blood mononuclear cells (PBMC) purified from leukocyte reduction system (LRS) cones obtained from the apheresis of donated blood samples. Monocytes are sorted by adherence to generate primary macrophages.
Collect the LSR cone and preserve it at 4 °C until use.
Lock one of the LRS cone accesses with hemostatic forceps.
Use a sterile surgical scissor to cut the opposite LRS cone access.
Insert a 23 G hypodermic needle attached to a 5 mL syringe into the cone, collect the cells, and transfer them to a 50 mL conical tube.
Transfer up to 2.5 mL of the collected sample to a 50 mL conical tube. Split the sample into two conical tubes if necessary.
Prepare a cell suspension by adding 35 mL of cold PBS to the sample and thoroughly mix by up-and-down pipetting.
Transfer 13 mL of Histopaque®-1077 solution to a clear 50 mL conical tube.
Overlay the cell suspension onto the top of Histopaque®-1077 solution without mixing by carefully dropping the cell suspension on the tube wall using a sterile disposable Pasteur pipette.
Set up the centrifuge to medium acceleration and no break. Centrifuge the gradient at 640× g for 30 min at 4 °C. If successful, the formation of a gradient with four separate layers should be clear (Figure 1). Carefully manipulate the tube to avoid mixing them up.
Figure 1. Schematics of the gradient obtained by centrifugation to isolate peripheral blood mononuclear cells (PBMCs)
Using a disposable Pasteur pipette, discard the top layer (remaining plasma and thrombocytes).
Using a new disposable Pasteur pipette, collect the intermediate layer containing PBMCs and transfer to a clear 50 mL conical tube.
Resuspend the PBMC layer in 25 mL of cold PBS and centrifuge the cells (400× g for 5 min).
Dump the PBS and resuspend the PBMCs in 5 mL of cold ACK lysis buffer (see Recipes).
Incubate the PBMC solution on ice for 5 min.
Add 25 mL of PBS to the tube, mix by pipetting, and centrifuge the cells.
Dump the PBS, resuspend the PBMCs in 5 mL of pre-warmed PBS, and count the PBMCs.
Transfer 2.4 × 108 PMBCs to a clear 50 mL conical tube (Tube A) and centrifuge them.
Transfer 4.0 × 107 PBMCs to a clear 50 mL conical tube (Tube B) for compensation controls as described in step A2b. Centrifuge the tube, dump the PBS, and resuspend the PBMC pellet in 2 mL of fresh pre-warmed RPMI without serum (2.0 × 107 cells/mL). The cells should be kept at 37 °C and 5% CO2 until seeding.
Dump the medium in tube A and resuspend the PBMC in 2.4 mL (108 cells/mL) of pre-warmed PBS.
Proceed to cell labeling with CellTraceTM Violet or CFSE (working solution concentration: 5 μM) following steps A4a–A4d.
Following the washout step, resuspend the PBMC pellet in 12 mL of fresh pre-warmed RPMI without serum (2.0 × 107 cells/mL).
Seed 500 µL (1.0 × 107 cells) of the PBMC suspension in tubes A and B per well of a 24-well cell culture plate and gently swirl the plate to evenly distribute the cells. Spare 2–3 of the wells seeded with labeled PBMCs (from tube A) to confirm the amount of differentiated macrophages immediately before proceeding to the efferocytosis assay.
Incubate the plate for 1 h at 37 °C and 5% CO2.
Thoroughly rinse each well thrice with pre-warmed PBS to remove unattached leucocytes. Monocyte sorting by adherence should give on average 1.0 × 106 cells/well.
Aspirate all the medium and replenish with 1 mL of pre-warmed RPMI supplemented with 10% of human serum (RPMIh) per well.
Incubate the plate for 72 h at 37 °C and 5% CO2.
Add 1 mL of fresh pre-warmed RPMIh per well and incubate the plate for 72 h at 37 °C and 5% CO2.
Lift the macrophages of 2–3 wells using a cell scraper and count cells to confirm cell confluence (Note 2).
Macrophages are ready for efferocytosis assay.
Generation of stained apoptotic Jurkat cells
Note: Jurkat cells are maintained in 175 cm2 cell culture flasks in RPMIc at 37 °C and 5% CO2. The medium is replaced every other day. Whenever mentioned below, cell transferring and resuspension were performed using disposable serological pipettes. All media and buffers were pre-warmed prior to use.
Collect the cell suspension from the flask, transfer to a 50 mL conical tube, and pellet them by centrifugation (400× g for 5 min at room temperature).
Dump the medium, resuspend the pelleted cells in 5 mL of fresh RPMIc, and count them.
Transfer 3.0 × 107 Jurkat cells to a 50 mL conical tube and centrifuge them.
Dump the medium and resuspend the pelleted cells in 3 mL of fresh RPMIc.
Transfer the cells to a tissue culture dish (100 mm × 20 mm).
Using the UV crosslinker, irradiate the tissue culture dish at 50 mJ/cm2 (Notes 3 and 4).
Collect the irradiated cells and transfer them to a clear 50 mL conical tube (Tube A).
Rinse the tissue culture dish to collect remaining cells with 9 mL of PBS and transfer the suspension to Tube A. Transfer 2.0 mL of the cell suspension (5.0 × 106 cells) from Tube A to another clear 50 mL tube (Tube B). Centrifuge them, dump the media, and resuspend the pelleted cells in 10 mL of fresh RPMIc. These cells will be spared to prepare experimental control tubes FMO AC 1 and FMO AC 2 (Figure 1).
Centrifuge the remaining cells in tube A, discard the medium, and resuspend the cells in 3 mL of PBS (1.0 × 107 cell/ mL).
Proceed to cell labeling with CellTraceTM Far Red (working solution concentration: 1 μM) or pHrodoTM Red (working solution concentration: 5 nM), following steps A4a–A4d (Notes 5 and 6).
Following the washout step, resuspend labeled ACs in 10 mL of fresh RPMIc.
Transfer the cell suspensions in Tube A and B to separate tissue culture dishes.
Incubate them for 4 h at 37 °C and 5% CO2 to allow apoptosis to proceed.
Collect the ACs and transfer them to 50 mL conical tubes.
Rinse the tissue culture dish to collect remaining cells with 10 mL of pre-warmed PBS and transfer the suspension to Tube A.
Centrifuge the AC tubes, discard the medium, resuspend the cells in 5 mL of RPMIc or RPMIh (if using THP1- or PBMC-derived macrophages assays, respectively), and count them.
Adjust the cell concentration to 5.0 × 106 AC/mL with RPMIc or RPMIh and proceed to efferocytosis assay.
Efferocytosis assay and sample preparation for data acquisition
Rinse the plate of CFSE-labeled macrophages with 500 μL of pre-warmed PBS per well.
Transfer 200 µL of the labeled AC 1 preparation in tube A to each well of the experimental group for single uptake or first round of continual uptake. This should give a 1:1 macrophage to AC ratio (cell ratio may vary according to experimental settings).
Transfer 200 µL of the unlabeled AC 1 preparation in tube B to each well of the FMO AC 1 control group (Figure 1).
Gently swirl the plate to evenly distribute the ACs.
Incubate the plate for 2 h at 37 °C and 5% CO2.
If performing single uptake, proceed to step D11.
If performing continual efferocytosis assay, incubate the plate for 18 h instead at 37 °C and 5% CO2. Proceed to step D6 (Note 7).
Remove the cell medium in each well and rinse the plate thrice using pre-warmed 500 μL of PBS per well.
Transfer 200 µL of the labeled AC 2 preparation in tube A to each well of the experimental group fed with labeled AC 1.
Transfer 200 µL of the unlabeled AC 2 preparation in tube B to each well of the FMO AC 2 control group fed with labeled AC 1 (Figure 1).
Gently swirl the plate to evenly distribute the ACs.
Incubate the plate for 2 h at 37 °C and 5% CO2.
Rinse the plate thrice with pre-warmed 500 μL of PBS per well.
Add 200 μL of PBS to each well.
Lift the cells using a cell scraper. Rinse or replace the cell scraper between each group.
Transfer the samples to FACS tubes.
Prepare a solution by diluting Zombie NIR in PBS with enough volume for all samples (final dilution 1:400, final staining volume per tube: 50 μL).
Centrifuge the samples (400× g for 5 min).
Resuspend the pellets in 50 μL of Zombie NIR solution.
Incubate the samples on ice for 10 min in the dark.
Centrifuge the samples.
Dump the solution and rinse the samples twice with 500 μL of PBS per tube. If necessary, resuspend the cells in 250 μL of a PBS solution with 2% paraformaldehyde and incubate for 15 min on ice for cell fixation.
Resuspend the pellet in 250 μL of PBS and proceed to data acquisition with a flow cytometer.
Data analysis
Note: Gating strategy to quantify AC uptake in single and continual efferocytosis is shown in Figure 1. We recommend acquiring at least 50,000 total events.
Export acquired data to .fcs file format and analyze using the software FlowJo.
Apply morphological criteria to gate the macrophage cluster and exclude debris based on forward (FSC) and side scatter (SSC) profiles.
Exclude doublets creating a gate through the diagonal area on FSC-A vs. FSC-H plot.
Exclude dead cells based on the staining for Zombie NIR viability dye.
Select the macrophages based on the positive staining for CFSE.
In the CFSE+ macrophage subpopulation, create a gate for the first round of AC uptake (AC1) using FMO AC1 as the reference tube. The percentage of positive cells in this gate represents the rate of the single uptake or first round of the continual uptake.
If performing continual uptake, create a gate for the second AC uptake in the CFSE+ AC-pHrodo+ macrophage subpopulation, using FMO AC2 as the reference tube. The percentage of positive cells in this gate represents the percentage of labeled macrophages that engulfed AC1 and were also efficient in the uptake of a second AC.
Validation of protocol
An example of the percentage of apoptotic cell uptake in the first (AC1) and second (AC2) rounds of efferocytosis by THP-1 cells following the steps described in the Procedure and Data analysis section is shown in Figure 2B.
Figure 2. Flow cytometric analysis to determine the uptake of apoptotic cells by the macrophages. THP-1-derived macrophages (CTV-labeled) were incubated with UV-irradiated apoptotic Jurkat cells for 18 h (first round, pHRodo-labeled AC1), and subsequently incubated with a second batch of apoptotic Jurkat cells for 2 h (second round, CTFR-labeled AC2), following the schematics described on the Graphic Abstract. (A) Representative flow plot of gating strategy using FMO controls. Macrophages are initially selected, and debris are excluded based on morphology profile on FSC-A vs. SSC-A plot. Next, doublet exclusion is performed on FSC-A vs. FSC-H plot. Next, dead cells were excluded based on staining for viability dye (Zombie-NIR). Next, labeled macrophages (CFSE+) were selected. Gating for labeled macrophages that uptake AC on the single uptake or the first round (AC1) is performed using FMO AC1 sample as reference control. Gating for labeled macrophages that uptake AC on the second round (AC2) is performed using FMO AC2. (B) Percentage of macrophages with internalized AC1 and AC2. Boxes represent the mean of four biological replicates and error bars are ± S.E.M. Each biological replicate is shown as a circle. Significance was calculated by Student’s t-test.
This protocol or parts of it has been used and validated in the following research article:
Salina et al. (2022). Efferocytosis of SARS-CoV-2-infected dying cells impairs macrophage anti-inflammatory functions and clearance of apoptotic cells. eLife, 11, e74443.
Data of single and continual uptake of AC by PBMC and THP-1-derived macrophages is presented in Figure 1 and supplement Figure 1 and Figure 4 of the aforementioned article.
Notes
It is possible to use cell surface staining following the efferocytosis assay to label the macrophages instead of using fluorescent probes. However, in our experience and as reported by others (Forrester et al., 2018), we found that THP-1-derived macrophages may express low levels of common surface markers used for phenotyping.
We recommend confirming macrophage differentiation by flow cytometry phenotyping.
Induction of apoptosis by UV irradiation may need to be adjusted according to different equipment.
We recommend the evaluation of the efficiency of apoptosis induction using standard apoptosis assay using flow cytometry (i.e., using fluorescent annexin V and viability dye co-staining). If inducing cell death with other methodologies (i.e., pharmacological drugs or by genetic manipulation), labeling may need to be performed prior to induction.
The amount of Jurkat cells for labeling can be scaled but avoid altering cell concentration. Ideal cell concentration for optimum labeling may vary for different cell types used to prepare the AC.
There is contradicting data on the literature about CFSE leakage from apoptotic cells. For that reason, we do not recommend its use to label the ACs. Other commercially available fluorescent probes may be used to label target ACs and assess their uptake by flow cytometry, such as CypHer5E, DAPI, and PKH26.
Incubation time for the first round of continual uptake may vary according to experimental settings. Labeling with pHrodoTM Red is associated with phagolysosome acidification and its specificity should be taken into consideration for ideal experimental design.
Acknowledgments
The authors thank Sra. Denise B. Ferraz (FMRP-USP) for technical assistance. Funding: This work was supported by grants from Fundacao de Amparo a Pesquisa do Estado de Sao Paulo (FAPESP) grants 2018/25559–4 and 2020/05288–6; Conselho Nacional de Desenvolvimento Cientifico e Tecnologico (CNPq) grant 434538/2018–3; Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) grant 88887.507253/2020-00. The schematics in this article were created with Biorender (www.BioRender.com). The protocol described here has been performed in Salina et al. (2022).
Competing interests
The authors declare no competing financial interests.
Ethical considerations
The procedures followed in the study were approved by the Research Ethics Committee of Hospital das Clínicas de Ribeirão Preto (CEP-FMRP/USP) and by the National Ethics Committee, Brazil (Comissão Nacional de Ética em Pesquisa (CONEP), protocols 30248420.9.0000.5440 and 39722020.9.0000.5440. Written informed consent was obtained from recruited donors.
References
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© 2023 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Immunology > Immune cell function > Macrophage
Cell Biology > Cell-based analysis > Flow cytometry
Cell Biology > Cell-based analysis > Endocytosis
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4,904 | https://bio-protocol.org/en/bpdetail?id=4904&type=0 | # Bio-Protocol Content
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Identifying Antigenic Switching by Clonal Cell Barcoding and Nanopore Sequencing in Trypanosoma brucei
AT Abdoulie O. Touray
TS Tamara Sternlieb
TI Tony Isebe
IC Igor Cestari
Published: Vol 13, Iss 24, Dec 20, 2023
DOI: 10.21769/BioProtoc.4904 Views: 890
Reviewed by: Marcelo S. da Silva Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Nov 2023
Abstract
Many organisms alternate the expression of genes from large gene sets or gene families to adapt to environmental cues or immune pressure. The single-celled protozoan pathogen Trypanosoma brucei spp. periodically changes its homogeneous surface coat of variant surface glycoproteins (VSGs) to evade host antibodies during infection. This pathogen expresses one out of ~2,500 VSG genes at a time from telomeric expression sites (ESs) and periodically changes their expression by transcriptional switching or recombination. Attempts to track VSG switching have previously relied on genetic modifications of ES sequences with drug-selectable markers or genes encoding fluorescent proteins. However, genetic modifications of the ESs can interfere with the binding of proteins that control VSG transcription and/or recombination, thus affecting VSG expression and switching. Other approaches include Illumina sequencing of the VSG repertoire, which shows VSGs expressed in the population rather than cell switching; the Illumina short reads often limit the distinction of the large set of VSG genes. Here, we describe a methodology to study antigenic switching without modifications of the ES sequences. Our protocol enables the detection of VSG switching at nucleotide resolution using multiplexed clonal cell barcoding to track cells and nanopore sequencing to identify cell-specific VSG expression. We also developed a computational pipeline that takes DNA sequences and outputs VSGs expressed by cell clones. This protocol can be adapted to study clonal cell expression of large gene families in prokaryotes or eukaryotes.
Key features
• This protocol enables the analysis of variant surface glycoproteins (VSG) switching in T. brucei without modifying the expression site sequences.
• It uses a streamlined computational pipeline that takes fastq DNA sequences and outputs expressed VSG genes by each parasite clone.
• The protocol leverages the long reads sequencing capacity of the Oxford nanopore sequencing technology, which enables accurate identification of the expressed VSGs.
• The protocol requires approximately eight to nine days to complete.
Graphical overview
Keywords: Oxford nanopore sequencing Trypanosomes Antigenic switching DNA barcode VSG-seq Variant surface glycoproteins Antigenic variation
Background
Trypanosoma brucei is a single-celled protozoan parasite that causes African trypanosomiasis and evades the host antibody response by changing its surface coat by antigenic variation (Cestari and Stuart, 2018). T. brucei expresses a single variant surface glycoprotein (VSG) gene from one of the 20 telomeric expression sites (ESs) and periodically switches to a different VSG via transcriptional switching between ESs or by VSG gene recombination. T. brucei has an extensive repertoire of over 2,500 VSG genes and pseudogenes located in telomeric and sub-telomeric regions of large chromosomes. VSG genes are also found on dozens of mini chromosomes often used for VSG gene recombination. VSG genes are approximately 2 kb in length with conserved C-terminus sequences. VSG recombination can occur by gene or segmental gene conversion, resulting in new mosaic VSG sequences. The mechanisms controlling VSG monogenic expression and switching likely entail multiple processes, including controlling VSG repression and expression via proteins associated with telomeric ESs. Several proteins associate with the telomeric repeats or ESs to regulate VSG gene expression and/or switching, such as the repressor activator protein 1 (Touray et al., 2023), phosphatidylinositol phosphate 5-phosphatase (PIP5Pase) (Cestari et al., 2019), telomeric repeat-binding factor (Jehi et al., 2014), VSG exclusion protein 2 (Faria et al., 2019), and ES body 1 protein (López-Escobar et al., 2022); for a review on additional proteins controlling VSG expression and switching, see (Cestari and Stuart, 2018).
Approaches used to study VSG switching rely on genetic modifications that disrupt the ES DNA sequences by incorporating drug-selectable markers or fluorescent proteins downstream of the promoter sequence and upstream of the VSG gene (Rudenko, 1998; Ulbert et al., 2002; Aitcheson et al., 2005) or by adding exogenous endonuclease sites resulting in DNA breaks (Boothroyd et al., 2009; Glover et al., 2013). However, the ES modifications might disrupt protein binding sites and thus affect VSG switching rates; as an example, RAP1 binds to 70 bp and telomeric repeats flanking ES VSG genes and represses their transcription, and disruption of its binding dramatically increases VSG switching rates (Touray et al., 2023). In addition, the genetic modifications of ESs are laborious and restrict the use of drug-selectable markers available for other genetic changes, such as gene knockout or expression of mutant variants in the cell. Other studies used Illumina RNA-seq to investigate VSG expression at a population level (Mugnier et al., 2015). Although this approach helps to identify expressed VSG genes, many short reads fail to align uniquely to the genome and unambiguously identify and distinguish VSG genes expressed from the extensive repertoire of VSG genes/pseudogenes. Moreover, it does not identify switching cells but expressed VSG genes in the population.
Hence, we sought to develop an approach to track cell-specific antigenic switching without genetically modifying ES sequences in an adaptable high- or medium-throughput fashion. We devised a method to detect VSG switching at nucleotide resolution using clonal cell barcoding and nanopore sequencing. We combine DNA barcoding to identify parasite cell clones and thus track switching and non-switching cells and a broad-spectrum VSG primer for cDNA synthesis to capture all transcribed VSGs in parasite clones. Barcoded samples are multiplexed, and VSG cDNAs are sequenced using Oxford nanopore sequencing, followed by sequencing data analysis using a streamlined VSG-BarSeq pipeline to identify switchers. We performed VSG-BarSeq after a temporary knockdown of PIP5Pase, an enzyme that regulates VSG switching in T. brucei (Touray et al., 2023). We found that 99% of the clones switched VSG genes, whereas no switching was detected in the control no-knockdown cell line. The long nanopore reads helped to identify complete VSG sequences and thus track modes of switching. We detected VSG switching by transcriptional and recombination mechanisms, indicating the approach's usefulness in studying antigenic variation. This protocol will enable the efficient study of VSG switching without genetic alterations of the ES. The protocol may be easily adapted to other organisms to study antigenic switching (e.g., Plasmodium, Giardia) or clonal expression analysis of large gene families in prokaryotes or eukaryotic cells.
Materials and reagents
Biological materials
Trypanosoma brucei 427 strain bloodstream forms or conditional null for the gene PIP5Pase derived from the 427 strain (Cestari and Stuart, 2015)
Reagents
M-MuLV reverse transcriptase (New England Biolabs Ltd, catalog number: M02535)
M-MuLV reverse transcriptase 10× buffer (New England Biolabs Ltd, catalog number: B02535)
MgCl2 (Invitrogen, catalog number: R0971)
NucleoMag NGS Clean-up and Size Selection beads (Takara, catalog number: 744970.5)
NEBNext end repair module (New England Biolabs Ltd, catalog number: E6050S)
Native Barcoding Expansion 1-12 (Oxford Nanopore Technologies, catalog number: EXP-PBC001)
10 M Sodium hydroxide solution (NaOH) (Sigma, catalog number: 1310-73-2)
NEBNext Quick Ligation Module (New England Biolabs Ltd, catalog number: E6056S)
NEBNext FFPE DNA Repair Mix 24 reactions (New England Biolabs Ltd, catalog number: M6630S)
Taq DNA polymerase with ThermoPol buffer (New England Biolabs Ltd, catalog number: M0267S)
Deoxyribonucleotide triphosphate (dNTPs) mixture 10 mM (Biobasic, catalog number: DD0056)
Agarose (Bioshop Canada Inc., catalog number: AGA002.250)
Iscove's modified Dulbecco's medium (IMDM) powder (Life Technologies, catalog number: 12200069)
Sodium bicarbonate (Fisher Scientific, catalog number: S233-3)
Hypoxanthine (Millipore, catalog number: 4010CBC-25GM)
Sodium pyruvate (Wisent Inc., catalog number: 600-110-EL)
Bathocuproin sulfonate disodium salt hydrate (Thermo Scientific, catalog number: B22550.MD)
L-cysteine hydrochloride monohydrate (Thermo Scientific, catalog number: J14035-22)
2-mercaptoethanol (Sigma, catalog number: M722)
Penicillin streptomycin (MP Biomedicals, catalog number: 1670049)
Heat-inactivated fetal bovine serum (FBS) (Life Technologies, catalog number:12484-028)
Neomycin (G418) (Sigma, catalog number: N6386)
Tetracycline hydrochloride (Fisher Scientific, catalog number: BP912-100)
Sodium phosphate dibasic anhydrous (Na2HPO4) (Fisher Scientific, catalog number: S374-500)
Sodium phosphate monobasic anhydrous (NaH2PO4) (Sigma, catalog number: S-0751)
Sodium chloride (NaCl) (Fisher Scientific, catalog number: S-271-3)
Glucose (Sigma, catalog number: G5400-250G)
Sodium citrate (dihydrate) (Fisher Scientific, catalog number: S279-500)
Bovine serum albumin (BSA) (Biobasic, catalog number: A500023-0100)
Glycerol (Fisher Bioreagents, catalog number: BP229-1)
Tris base (Fisher Scientific, catalog number: BP152-5)
Glacial acetic acid (Fisher Chemical, catalog number: A38-212)
Ethylenediaminetetraacetic acid (EDTA) (Fisher Scientific, catalog number: BP120-500)
Trypan Blue solution, 0.4% (Amresco, catalog number: K940-100ML)
Ethyl alcohol anhydrous solution (Commercial Alcohols, catalog number: P016EAAN)
96-well plate bacterial total RNA mini-prep super kit (Biobasic, catalog number: BS585-5)
Oxford Nanopore Ligation Sequencing Kit (Oxford Nanopore Technologies, catalog number: SQK-LSK109)
AMPure XP beads (Beckman Coulter Inc., catalog number: A63880)
Phleomycin (Bioshop Canada Inc., catalog number: PEO422.10)
Solutions
HMI-9 cell culture medium (see Recipes)
Drug selectable markers (see Recipes)
Neomycin (G-418) stock solution (20 mg/mL)
Tetracycline stock solution (5 mg/mL)
Phleomycin stock solution (1 mg/mL)
Phosphate-buffered saline-glucose (PBS-G) (see Recipes)
Bloodstream stabilate freezing solution (see Recipes)
50× Tris acetate EDTA solution (TAE) (see Recipes)
80% Ethanol (see Recipes)
Recipes
HMI-9 cell culture medium (1,000 mL)
Reagent Final concentration Quantity or Volume
IMDM n/a 17.7 g
Sodium bicarbonate 0.035 M 3 g
Hypoxanthine 10 mM 10 mL
Sodium pyruvate 100 mM 10 mL
Bathocuproine sulfonate disodium salt hydrate 49.95 µM 28.2 mg
L-cysteine hydrochloride monohydrate 1.04 mM 182 mg
2-mercaptoethanol 14.3 M 14 µL
Nanopure MilliQ water n/a 879.9 mL
Total volume n/a 900 mL
Dissolve the IMDM and salts in 500 mL of water and stir to mix thoroughly. Add 10 mL of hypoxanthine, 10 mL of sodium pyruvate, and 14 µL of 2-mercaptoethanol. Adjust water volume to 900 mL. Filter the medium using a 0.22 µm filter system of 250 mL. Add 100 units of sterile penicillin and 100 µg/mL sterile streptomycin (optional). Complete the media by adding 10% heat-inactivated FBS. Store at 4 °C. The shelf life of the medium is four months.
Drug-selectable markers
Neomycin (G-418) stock solution (20 mg/mL)
Reagent Final concentration Quantity or Volume
Neomycin (G-418) n/a 1 g
Nanopure MilliQ water n/a 50 mL
Total n/a 50 mL
Tetracycline stock solution (5 mg/mL)
Reagent Final concentration Quantity or Volume
Tetracycline hydrochloride n/a 0.5 g
Nanopure MilliQ water n/a 10 mL
Total n/a 10 mL
Phleomycin stock solution (1 mg/mL)
Reagent Final concentration Quantity or Volume
Phleomycin n/a 0.05 g
Nanopure MilliQ water n/a 50 mL
Total n/a 50 mL
Dissolve the drugs in water. Filter the dissolved drug solutions using a 0.22 µm syringe filter. Prepare 1 mL aliquots into 1.5 mL Eppendorf tubes in the hood and store the aliquots at -20 °C freezer. Aliquots are stable for at least one year when stored at -20 °C.
Phosphate-buffered saline-glucose (PBS-G) (1,000 mL)
Reagent Final concentration Quantity or Volume
Sodium phosphate dibasic anhydrous 0.01 M 1.42 g
Sodium phosphate monobasic anhydrous 0.01 M 1.20 g
Sodium chloride 0.145 M 8.5 g
D-Glucose 0.006 M 1.081 g
Nanopure MilliQ water n/a 1,000 mL
Total n/a 1,000 mL
Dissolve salts in 900 mL of water. Add D-Glucose and stir thoroughly to dissolve it. Adjust the pH to 7.0 and filter sterilize using a 0.22 µm filter. Store at 4 °C. This buffer is stable for three months at 4 °C.
Bloodstream stabilate freezing solution (500 mL)
Reagent Final concentration Quantity or Volume
D-Glucose 0.052 M 9.3 g
Sodium chloride 0.036 M 2.1 g
Sodium citrate 0.003 M 0.75 g
BSA n/a 0.5 g
Glycerol n/a 75.0 g
Nanopure MilliQ water n/a 500 mL
Total n/a 500 mL
Dissolve the salts in 300 mL of water. Add D-glucose and stir thoroughly to dissolve it. Weigh the volume of glycerol equivalent to 75 g in a separate container and add it to the solution. Adjust the volume of the solution to 500 mL and stir thoroughly. Filter using a 0.22 µm filter. Store it at 4 °C. The solution is stable for six months.
50× Tris Acetate EDTA solution (TAE) (500 mL)
Reagent Final concentration Quantity or Volume
Tris base 1 M 121 g
Glacial acetic acid 1 M 28.6 mL
EDTA 0.05 M 50 mL
Nanopure MilliQ water n/a 421.4 mL
Total volume n/a 500 mL
Prepare a stock solution of 0.5 M EDTA in a separate tube by dissolving 18.6 g of EDTA disodium salt in 80 mL of water. Adjust the pH to 8.0 with 10 M NaOH solution. Adjust the volume to 100 mL. Dissolve the Tris base in 300 mL of water and add 28.6 mL of 17.4 M glacial acetic acid and 50 mL of 0.5 M EDTA. Adjust the volume of the solution to 500 mL with water. Autoclave the solution. Store it at room temperature (RT). The solution is stable for at least six months.
80% Ethanol
Reagent Final concentration Quantity or Volume
Ethyl alcohol anhydrous solution (100% v/v) 80 % 80 mL
Nanopure MilliQ water n/a 20 mL
Total volume n/a 100 mL
Prepare 80% v/v ethanol by transferring 80 mL of 100% Ethyl alcohol solution to a 100 mL graduated cylinder. Then, add 20 mL of Nanopure MilliQ water. Store at 4 °C. The solution is stable for one month.
Laboratory supplies
Tissue culture flasks
25 cm2 TC-treated T-flask with filter cap (Biobasic, catalog number: SP81136)
75 cm2 TC-treated T-flask with filter cap (Biobasic, catalog number: SP81186)
Cell culture plates
96-well cell culture plates (WUXI NEST Biotechnology Co., catalog number: 101722BL01)
24-well TC plates, treated (Biobasic, catalog number: SP41135)
Sterile pipette tips
1,000 µL pre-sterile barrier tips (Neptune, catalog number: BT100.96)
200 µL pre-sterile barrier tips (Neptune, catalog number: BT200)
10 µL pre-sterile barrier tips (Neptune, catalog number: BT10)
96-well 2 mL deep plate, natural, edge filled, 10 plates/bag (Biobasic, catalog number: BR581-96NS)
96-well deep collection plates (Biobasic, catalog number: 107-E627LA2221)
Plate seals (Biobasic, catalog number: BS585-5)
96-well PCR plate (Life Technologies, catalog number: 4346907)
Foil sealing film, non-sterile (Celltreat Scientific Products, catalog number: 501535152)
Multichannel pipette (Eppendorf, catalog number: 4056991)
Heating block (Eppendorf, catalog number: 535028642)
Filter system 250 mL 0.2 µm PES (Fisher Scientific, catalog number: FB12566502)
2.0 mL microtubes (UltiDent Scientific, catalog number: 48-C200-CS)
1.5 mL Eppendorf tubes (Eppendorf, catalog number: 0030123611)
100 mL graduated cylinder (Grainger, catalog number: GGS5PTJ5)
100 mL reagent bottle (UltiDent Scientific, catalog number: 170-14170100)
Equipment
Ultrafocused sonicator (Covaris, M220, catalog number: 006168)
Magnetic rack for 1.5 mL tubes (Promega, catalog number: PR-Z5342)
T100 thermal cycler (Bio-Rad, catalog number: 1861096)
Avanti J-E centrifuge (Beckman Coulter, catalog number: JSE02M13)
AllegraTM 25R centrifuge, TJ-25 rotor (Beckman Coulter, catalog number: AJC024001)
Nanodrop (Nanodrop Spectrophotometer, catalog number: ND-1000)
Microcentrifuge (Eppendorf, catalog number: EP5401000137)
Water system ultrapure (Nanopure MilliQ water) (Millipore Synergy, catalog number: F1CA45528 A)
Electrophoresis unit: Thermo Scientific Power Supply 400 mA 300 V (Fisher, catalog number: S65533Q)
Biological safety cabinet (NuAire Biological Safety Cabinet Class II Type AIB3, catalog number: NU-425-300)
CellDropTM automated cell counter (DeNovix Cell Drop FL Fluorescence Cell Counter)
HEPA CO2 incubator (Thermo Electron Corporation; Forma Series II, Water Jacketed, catalog number: 308606-30529)
Microscope (Nikon, model: Eclipse TS100, catalog number: 302115)
-80 °C freezer (New Brunswick Ultra-Low Temperature Freezer U101 Inova, catalog number: U9420-0000)
MinION Mk1C (Oxford Nanopore Technologies, catalog number: MIN-101C)
Flongle Flow Cell (R9.4.1) (Oxford Nanopore Technologies, catalog number: FLO-FLG001)
Rotator (ManSci Inc., catalog number: A706514)
Milli-Q IQ 7000 purification system (Millipore Sigma, catalog number: ZIQ7000T0C)
Software and datasets
Minimap2, version 2.24 (Li, 2018) (https://github.com/lh3/minimap2)
Samtools, version 1.17 (Danecek et al., 2021) (https://github.com/samtools/samtools)
DeepTools, version 2.0 (Ramírez et al., 2016) (https://deeptools.readthedocs.io/en/develop/index.html)
Subread, version 2.0.3 (Liao et al., 2014) (http://subread.sourceforge.net/featureCounts.html)
Rcgrep, version 0.1 (https://github.com/dib-lab/rcgrep)
Integrative Genomics Viewer (IGV), version 2.16.2 (Robinson et al., 2011) (https://software.broadinstitute.org/software/igv/)
VSG-BarSeq (this work), version 1.0.1 (https://github.com/cestari-lab/VSG-Bar-seq)
Procedure
Parasite treatment and cloning
We recommend determining cell treatment conditions before starting this protocol. The conditions used in this protocol were optimized for T. brucei bloodstream forms of the 427 strain or conditional null (CN) cells derived from the single-marker 427 strain (Cestari and Stuart, 2015). The treatment described here is the knockdown for 24 h of the T. brucei gene encoding PIP5Pase, which results in high rates of VSG switching (Touray et al., 2023). The PIP5Pase CN cell line is grown in G418 and phleomycin to maintain the selection of the tetracycline-inducible system. Tetracycline is added to induce expression of the PIP5Pase gene under the control of a procyclin promoter and a tetracycline operator.
Grow 5 mL of T. brucei cells seeded at 1.0 × 104 cells/mL in HMI-9 medium supplemented with 2 µg/mL G418, 2.5 µg/mL phleomycin, and 500 ng/mL tetracycline in a 37 °C incubator with 5% CO2 for 24 h or until it reaches mid-log growth (~1.0 × 106 cells/mL). Throughout this protocol, cell growth will be as described above unless otherwise stated. Cells’ doubling time should be approximately 5.5–6 h and viability approximately 90%–95%. Avoid overgrowing cell culture (>1.5 × 106 cells/mL) because it will affect cell viability.
Transfer the 5 mL cell culture to a 15 mL Falcon tube and centrifuge at 3,500× g for 5 min at RT. Discard the supernatant.
Resuspend the pellet in 10 mL of PBS-G pre-warmed at 37 °C and then centrifuge the cells as in step A2. Discard the supernatant.
Repeat step A3 three times to ensure complete removal of tetracycline.
Split the cells into two 5 mL cell culture flasks (treated and non-treated groups), seeding each culture at 1.0 × 104 cells/mL in HMI-9 medium with 2 µg/mL G418 and 2.5 µg/mL phleomycin.
Add 500 ng/mL tetracycline to the non-treated flask (Tet +, control) and no tetracycline to the treatment flask (Tet -, knockdown) and grow the cells for 24 h.
Add 500 ng/mL tetracycline to the treatment flask (Tet -, knockdown). No additional tetracycline is required for the non-treated flask (Tet +, control). Quantify the cell concentration and viability of both the treatment and control groups by mixing 10 µL of cell culture and 10 µL of 0.4% Trypan blue staining. Add 10 µL to the CellDropTM cell counter to obtain viability and cell concentration. Cell concentration should be approximately 1.0 × 105 cells/mL, and viability should be >90%.
Add 9 mL of HMI-9 medium supplemented with 500 ng/mL tetracycline to a 50 mL Falcon tube. Repeat the procedure to have three flasks for the treatment group and three for the non-treatment group.
From a starting cell concentration of 1.0 × 105 cells/mL (from A7 above), gently mix the cells by flicking the flasks five times and transfer 1 mL of the culture to the Falcon tube 1 containing 9 mL of HMI-9 media (1 in 10 dilutions) to obtain a cell concentration of 1 × 104 cells/mL.
From the Falcon tube 1 (1 × 104 cells/mL) culture, transfer 1 mL to Falcon tube 2 to obtain a cell concentration of 1 × 103 cells/mL (1 in 10 dilutions). Repeat the same for Falcon tube 3 to obtain a cell concentration of 1 × 102 cells/mL. Perform the procedure for control and treated groups.
Transfer 7 mL of the culture from Falcon tube 3 to a 75 cm2 cell culture flask containing 63 mL of HMI-9 medium supplemented with 2 µg/mL G418, 2.5 µg/mL phleomycin, and 500 ng/mL tetracycline, to obtain a cell concentration of 10 cells/mL (1 in 10 dilutions).
Transfer 60 mL of the diluted cell culture from step A11 to a 75 cm2 cell culture flask and add 140 mL of HMI-9 medium supplemented with 2 µg/mL G418, 2.5 µg/mL phleomycin, and 500 ng/mL tetracycline (3 in 10 dilutions) to obtain a final cell concentration of 3 cells/mL.
Aliquot the diluted parasite culture (3 cells/mL) onto ten 96-well cell culture plates (10 plates per treatment group) using a multichannel pipette. Transfer 200 µL to each well so that the probability of obtaining one single cell per well (200 µL) is approximately 30%, i.e., one cell per well for a third of the wells of a 96-well cell culture plate. Ensure the cells are well mixed by gently swirling the parasite cultures while pipetting.
Grow the cells in 96-well cell culture plates in an incubator for 5–7 days.
Check the 96-well cell culture plates under a microscope to identify parasite clones. Approximately 30% of the wells should contain parasites (see Note 1). We recommend checking each well for parasite clones starting from day 5 up until day 7 post seeding.
Transfer 200 µL of parasite clones onto 24-well TC plates and add 1.8 mL of fresh HMI-9 with 2 µg/mL G418, 2.5 µg/mL phleomycin, and 500 ng/mL tetracycline.
Grow the clones in the 24-well TC plates for 24 h to increase the number of cells for RNA extraction.
(Optional) Aliquot 600 µL of each clonal population onto new 96-well deep collection plates and add 600 µL of freezing solution. Freeze the parasites at -80 °C for short-term storage (2–3 weeks) or liquid nitrogen for long-term storage.
The remaining 1.4 mL of the clonal parasite cultures are used for RNA extraction.
RNA extraction
Transfer 1.4 mL of each clonal culture (approximately 1.4 × 106 cells) into 96-well deep collection plates, centrifuge at 3,500× g for 5 min at RT, and pour off the supernatant (see Note 2).
Add 350 µL of buffer Rlysis-BG (provided in the 96-well plate bacterial total RNA mini-prep super kit) to each well and resuspend the cells by pipetting up and down five times.
Thoroughly seal the plate with a sealing film to prevent cross-contamination of the samples and immediately mix the samples by inverting the plate three times.
Briefly spin the plate at 1,000× g for 30 s to collect the solution to the bottom of the wells.
Add 175 µL of absolute ethanol to each well, tightly seal the plate with a new sealing film, and mix thoroughly by inverting five times.
Place the EZ-10 96-well plate (filtration column plate provided with the 96-well plate bacterial total RNA mini-prep super kit) on top of a new 96-well deep collection plate and transfer the lysate from step B5 into the columns on the EZ-10 96-well plate.
Centrifuge the plate at 5,000× g for 2 min at RT and discard the flowthrough.
Place the EZ-10 96-well plate back on the deep cell collection plate and add 500 µL of universal GT solution (provided in the kit) to each column.
Centrifuge the plate at 5,000× g for 1 min at RT and discard the flowthrough.
Place the EZ-10 96-well plate back on the deep well collection plate and add 500 µL of universal NT solution (provided in the kit) to each column.
Centrifuge the plate at 5,000× g for 1 min at RT and discard the flowthrough.
Place the EZ-10 96-well plate back on the deep well collection plate and centrifuge the column at 5,000× g for 2 min at RT to ensure complete removal of the residual ethanol.
Place the EZ-10 96-well plate into a new deep-well storage plate (provided in the kit), add 30 µL of RNase-free water (supplied in the kit), and then incubate the plate at RT for 5 min.
Centrifuge the plate at 5,000× g for 1 min at RT to elute the RNA solution.
Quantify the recovered RNA by measuring 1 µL of the RNA solution at 260 nm from approximately 10 random wells using a NanoDrop. This will help estimate the isolated RNA concentrations.
Tightly seal the plate with an adhesive cover and keep it on ice (or at -80 °C) until the RNA samples are ready for cDNA synthesis.
VSG-enriched cDNA synthesis and barcoding
This step requires a combination of primers (Figure 1) to barcode each clonal cell population cDNAs with a unique eight-nucleotide sequence for their identification during sequencing analysis. It will also provide an adapter sequence for DNA sequencing library preparation. The forward Ad-SL (5′-TTTCTGTTGGTGCTGATATTGCacagtttctgtactatattg-3′) primer is universal and includes an Oxford nanopore adaptor sequence (capital letters) followed by a sequence (small letters) that hybridizes to mRNA splice leader sequence, a 39-nt sequence added to the 5′ of all trypanosomes’ mRNAs. The reverse Ad-3endVSG primer (5′-TACTTGCCTGTCGCTCTATCTTCXXXXXXXXgtgttaaaatatatc-3′) contains an Oxford nanopore adaptor sequence (capital letters) followed by eight-nucleotide long variable sequence (barcode) unique to each clone and a sequence pairing with the conserved 3′-end of VSG mRNAs, which encodes the C-terminus of VSG proteins (Mugnier et al., 2015). See Supplementary information for the complete primer list. We recommend preparing a working primer solution containing a mix of both primers at 10 µM in a 96-well plate.
Figure 1. Schematic representation of clonal cell barcode and nanopore sequencing protocol. (A) Diagram of the clonal cell barcoding, nanopore sequencing workflow showing the parasite treatment, and cloning (Step 1), RNA isolation from the individual clones, cDNA synthesis, and clonal barcoding (Step 2), and Oxford nanopore technology (ONT) library preparation, sequencing, and data analysis using VSG-BarSeq script (Step 3). (B) Scheme of the forward Adaptor Splice Leader (Ad-SL) primer and the reverse Adaptor-barcode-3′-end VSG primer (Ad-3endVSG) used for cDNA synthesis and clonal cell barcoding. (C) Diagram describing the cDNA synthesis, clonal barcoding, ONT library barcoding, and PCR amplification. (D) Sequences of the Ad-SL and Ad-3endVSG annealing primers used for cDNA synthesis and clonal cell barcoding (see the annealing regions in C). The sequences in black correspond to ~20 bp nanopore barcode adapter sequences, and the sequences in pink and green correspond to the forward Ad-SL and reverse Ad3endVSG primers, respectively. The Ad-SL pairs to the mRNA splice leader sequence, while the reverse Ad3endVSG primer pairs to the conserved 3′-end of VSG mRNAs. The 8-nucleotide long variable sequence (clonal barcode) unique to each clone is shown in orange and depicted here as X. A complete list of primers is available in Supplementary Information. PBS-G: phosphate-buffered saline (PBS)-glucose; ONT: Oxford Nanopore Technology; PIP5Pase CN: phosphatidylinositol 5-phosphatase (PIP5Pase) conditional null.
Take 4 μL of primer mix from the working primer solution plate and transfer it to each well of a new 96-well PCR plate using a multichannel pipette.
Add 1 μL of 10 mM dNTPs mix onto each well of the same 96-well PCR plate.
Thaw the RNA samples (from step B16) on ice if frozen at -80 °C and add 5 μL of the samples, keeping the same orientation in the 96-well PCR plate as the original RNA plate. Mix the solutions by gently pipetting up and down five times and spin down the plate briefly at 1,000× g for 30 s at 4 °C.
Incubate the plate at 65 °C for 5 min in a thermocycler and then transfer the plate immediately to ice.
Prepare cDNA synthesis master mix by adding 2 μL of MuLV 10× Buffer, 5 U of M-MuLV Reverse Transcriptase, and 8 U of RNase inhibitor (supplied in the MuLV reverse transcriptase kit) and adjust the reaction volume to 10 μL per reaction using nuclease-free water.
Aliquot 10 μL of the cDNA synthesis master mix onto each well of the 96-well plate containing the RNA, dNTPs, and primer mix.
Thoroughly seal the plate using an adhesive plate sealing film, spin down at 1,000× g for 30 s, and incubate at 42 °C for 2 h and then 65 °C for 20 min in a thermocycler.
Store the synthesized cDNA samples at -80 °C or proceed to library preparation.
Oxford nanopore library preparation and DNA sequencing
Combine the synthesized cDNA samples from each well (from step C8) into one 1.5 mL microcentrifuge tube and mix gently five times. Avoid forming bubbles.
Prepare 10 barcoding PCR reactions containing 5 μL of 10× ThermoPol buffer, 400 μM dNTPs mix, 500 nM barcode primer mix (Native Barcoding Expansion 1-12), 2 U of Taq DNA Polymerase, and 750 nM MgCl2. Add 3 μL of the pooled cDNA and adjust the final reaction volume to 50 μL with nuclease-free water.
Perform the PCR reaction in a thermocycler at 95 °C for 10 min, then 22 cycles at 95 °C for 1 min, 62 °C for 1 min, and 68 °C for 3:30 min, and a final extension at 68 °C for 10 min (see Notes 3 and 4).
Pool together all PCR amplicons in one 1.5 mL Eppendorf tube to obtain a final volume of 500 μL. Add 350 μL (0.65× beads to sample ratio) of NucleoMag NGS Clean-up and Size Selection beads to clean up the DNA.
Incubate beads/samples in a rotator (60 rpm) for 10 min at RT.
Spin down the tube briefly at 1,000× g for 30 s at RT, place it on a magnetic rack, and incubate for 1 min at RT. Pipette off and discard the supernatant without disturbing the beads.
Add 350 µL of freshly prepared 80% ethanol to the beads gently. Avoid disturbing the beads or taking the tube out of the magnetic rack. The volume of the 80% ethanol to add per wash should be equal to the sum of the total PCR amplicon volume and added NucleoMag NGS beads volume (e.g., 500 µL of PCR reaction + 350 µL of NucleoMag NGS magnetic beads = 850 µL of the mix, then 850 µL of 80% ethanol).
Incubate the mix for 1 min on the magnetic rack at RT. Collect and discard the 80% ethanol. Repeat steps D7 and D8.
Centrifuge the tube at 1,000× g for 30 s at RT. Place the tube back on the magnetic rack and remove any residual 80% ethanol from the tube.
Remove the tube from the magnetic rack and let the beads air dry for 10 min at RT.
Resuspend beads in 61 μL of nuclease-free water, mix gently by flicking the tube or gently pipetting up and down five times, and incubate for 5 min at RT.
Place the tube back on the magnetic rack and incubate for 1 min at RT.
Collect and transfer the clear supernatant containing the eluted DNA into a new labeled tube. Discard the tube containing the beads.
Measure and assess the concentration and purity of the eluted DNA by quantifying 1 µL of the DNA sample using NanoDrop. The DNA yield ranges from approximately 30 to 60 ng/μL with 260/280 and 260/230 absorbance ratios of approximately 1.8 and 2.0, respectively.
Prepare 1 μg of DNA in 47 μL of nuclease-free water in a 0.2 mL thin-walled PCR tube and then add the following: 3.5 μL of NEBNext FFPE DNA repair buffer and 2 μL of NEBNext FFPE DNA repair mix (from NEBNext FFPE DNA Repair Mix), 3.5 μL Ultra II End-prep reaction buffer and 3 µL Ultra II End-prep enzyme mix (from NEBNext end repair module), and 1 μL of DNA CS (provided in the Oxford Nanopore Ligation Sequencing kit).
Mix the solutions by pipetting up and down gently five times and incubate for 60 min at 20 °C in a thermocycler.
Add 39 μL of NucleoMag NGS Clean-up and Size Selection beads to the end-repaired DNAs (0.65× beads to samples ratio) to clean up and follow steps D5–D13. Elute the DNA in 61 μL of nuclease-free water.
Quantify 1 µL of the eluted DNA using NanoDrop. Approximately 90% of the DNA is expected to be recovered.
Transfer the remaining 60 µL of end-repaired DNA volume from step D18 (0.5–1 μg) to a 0.2 mL thin-walled PCR tube and then add the following components: 25 μL of ligation buffer (LNB), 10 μL of NEBNext Quick T4 DNA ligase (from NEBNext Quick Ligation Module), and 5 μL of Adapter mix F (AMX-F) (LNB and AMX-F are components of the Oxford Nanopore Ligation sequencing kit). Mix by gently pipetting the reaction up and down five times, then briefly spin it down.
Incubate the reaction for 2 h at 20 °C in a thermocycler.
Transfer the ligation reaction mix from step D20 to a new 1.5 mL Eppendorf tube and add 40 μL of well resuspended AMPure XP beads to the reaction (0.4× beads to sample ratio). Mix by gently pipetting up and down five times.
Incubate the mix on a rotator (60 rpm) for 10 min at RT.
Briefly centrifuge the sample at 1,000× g for 30 s and place the tube on a magnetic stand for 1 min at RT or until the supernatant is clear and colorless.
Remove and discard the supernatant without disturbing the beads.
Resuspend the beads in 250 μL of Short Fragment Buffer (SFB) (provided in the Oxford Nanopore Ligation Sequencing kit) and mix gently by flicking the tube five times to wash the beads.
Briefly centrifuge the tube at 1,000× g for 30 s. Transfer the tube to a magnetic stand to pellet the beads. Remove and discard the supernatant without disturbing the beads.
Repeat steps D25 and D26.
Briefly centrifuge the tube at 1,000× g for 30 s. Place it on the magnetic stand and remove any residual supernatant. Air dry the beads at RT and ensure that they do not dry to the point of cracking. Ten minutes is sufficient for the beads to dry. Extended drying of the beads for 20–30 min might cause them to crack.
Remove the tube from the magnetic stand and resuspend the beads in 12 μL of Oxford nanopore Elution Buffer (EB) (provided in the Oxford Nanopore Ligation Sequencing Kit) by gently pipetting up and down.
Briefly centrifuge the tube at 1,000× g for 30 s and incubate for 10 min at RT.
Place the tube back onto the magnetic stand for 1 min and then remove and transfer the eluate containing the DNA library into a new 1.5 mL Eppendorf tube. Dispose of the beads.
Quantify 1 µL of the eluted library DNA sample using a NanoDrop. Expect a DNA library yield of approximately 10–40 ng/μL (i.e., 200–1,500 fmol) of DNA.
Load 3–20 fmol of the prepared library into the Flongle Flow Cell (R9.4.1) and sequence the DNA according to the manufacturer's instructions.
Data analysis
Computational analysis of the sequenced library
The data analysis described here was performed using a Linux operating system (Ubuntu). The computational resources required will vary depending on the data available for analysis. The Oxford nanopore sequencer will generate fast5 files, which are basecalled to fastq files using the Guppy tool integrated in the MinKNOW software (https://nanoporetech.com/). The fastq files are the input dataset in the analysis shown here. We developed a computational pipeline for the sequencing analysis and detection of VSG switching (Figure 2). The pipeline is run via the vsg-barseq.sh script. A fastq file with 100,906 reads was analyzed using 10 threads and 4 GB of memory and completed in 5 min. The pipeline takes the DNA sequences generated by the nanopore sequencing in fastq format and splits them into subfiles according to the cell barcodes (eight mers) used for DNA-seq library preparation. The output fastq files are named by their barcodes, e.g., CCATGCAT.fastq. A split summary file (.txt) is generated and shows the number of reads per barcode file, the total count of reads analyzed, and the total amount of reads containing barcodes (see Note 5). Sequences from each file are then mapped to the organism genome (here, T. brucei 427 strain) using minimap2, outputting .sam files. The alignments are then filtered using Samtools to remove supplementary alignments and to keep alignments with a mapping quality score (mapQ) ≥ 20. The resultant .sam files are sorted and indexed with Samtools resulting in sorted.bam and bam.bai files, respectively.
The alignments are counted with featureCounts (package Subread). The top mapped reads, which correspond to the expressed VSG, are selected and compiled in a single tab-delimited file (topmapped.txt) while keeping the original output from featureCounts, which serves for analysis of other genes identified during alignment. The analysis of other identified genes is part of the quality control process. It typically shows genes with low counts reflecting low background noise from library preparation resulting primarily from sequences derived from splice-leader cDNA synthesis (Figure 3E).
The vsg-barseq.sh is open code and available on GitHub (https://github.com/cestari-lab/VSG-Bar-seq). The script takes six arguments:
1) Directory of all files (output folders will be created in this directory)
2) Directory of fastq files
3) Directory of barcode.txt files (barcode.txt file has one barcode sequence per row)
4) Directory of genome file in fasta format
5) Directory of gene transfer format file, i.e., gtf format
6) Number of threads to be passed to minimap2, samtools, and featureCounts (we recommend eight or more)
Example of barcode.txt file. Keep one barcode sequence per row.
CCGTTAGG
CCAACTAG
CAGGGCAG
CGCAGAAG
If the barcoded.txt file is generated using Windows operational system, we recommend converting from dos to Unix format before executing the analysis. The tools minimap2, samtools, subread, and rcgrep are required to run the script. After installing and/or loading the required tools, run the script as indicated below:
sh vsg-barseq.sh path/to/directory path/to/fastq path/to/barcodes path/to/genome path/to/gtf nthreads
See the example below using the structure folder exemplified (directories in bold).
sh vsg-barseq.sh \
c/user/vsg-barseq \
myfastq \
barcodes \
genomefiles/genome.fasta \
genomefiles/genome.gtf \
8
Note that T. brucei genome (.fasta) and features (.gff, general feature format) can be downloaded from TritrypDB (https://tritrypdb.org/). TriTrypDB does not provide .gtf files, but .gff files can be used to generate .gtf. We recommend using the gffread tool (https://github.com/gpertea/gffread), also available via Galaxy tools (https://usegalaxy.org/). The command below indicates how to generate a gtf file from a gff file after installing gffread tools.
gffread annotation_file.gff -T -o annotation_file.gtf
T. brucei barcodes and test fastq data files are also available for download at https://github.com/cestari-lab/VSG-Bar-seq to test the script.
Directory structure of input files required to run the script:
c/user/vsg-barseq/
----myfastq/mydata.fastq
----barcodes/barcode.txt
----genomefiles/genome.fasta, genome.gtf
After running the script, the following output directories and files are created:
c/user/vsg-barseq/
----result_split_fastq/CCGTTAGG.fastq, CCAACTAG.fastq, CAGGGCAG.fastq, CGCAGAAG.fastq, split_summary.txt
----result_mapcount/
----sam/CCGTTAGG.sam, CCAACTAG.sam, CAGGGCAG.sam, CGCAGAAG.sam,
----bam/CCGTTAGG.bam, CCAACTAG.bam, CAGGGCAG.bam, CGCAGAAG.bam
----sorted_bam/CCGTTAGG_sorted.bam, CCGTTAGG_sorted.bam.bai, etc.
----counts/CCGTTAGG.txt, CCGTTAGG.txt.summary, etc.
----topmap/topmapped.txt
The result file topmapped.txt will have information on gene id (first row, in bold below), chromosome (or contig), nucleotide position (start, end), strand (+/-), gene length in nucleotides, number of counted alignments for the gene (seventh row, in bold below), and path to the original file containing all alignment counts for the corresponding clone; the eight mers barcode identifies the cell expressing the VSG. Each row represents a cell clone, and the gene id indicates the VSG expressed by the cells in the clonal population.
Tb427_000016000 BES1_Tb427v10 75570 77000 + 1431 461 /path/to/counts/CCGTTAGG.txt
Tb427_000016000 BES1_Tb427v10 75570 77000 + 1431 363 /path/to/counts/CCAACTAG.txt
Tb427_000008000 BES12_Tb427v10 45278 46651 + 1374 310 /path/to/counts/CAGGGCAG.txt
Tb427_000284800 Chr2_5A_Tb427v10 197226 198701 + 1476 840 /path/to/counts/CGCAGAAG.txt
Figure 2. Flowchart of computational analysis using VSG-BarSeq. Multiplexed reads from clonal VSG-seq are split into clone-specific reads based on eight mers barcode. Reads are aligned to the genome using minimap2 and filtered with Samtools to remove supplementary and secondary alignments and keep alignments with mapQ ≥ 10. Then, it counts the alignments per gene and reports the top alignment per file, corresponding to the expressed variant surface glycoproteins (VSG) gene.
Anticipated results
The analysis of parasite VSG switching starts with treating the cultures to induce the switching of the VSG gene. The treatment conditions and outcomes might differ for different cell lines, so they may need to be optimized. Here, the treatment was the temporal knockdown (24 h) of the PIP5Pase gene, which results in high rates of VSG switching (Touray et al., 2023). VSG switching occurs by alternating transcription between ESs or recombining VSG genes within ESs (Figure 3A). The parasite culture was diluted to obtain approximately 30% of clones from a 96-well plate. We recommend optimizing the dilution of the cells after the treatment and checking cell viability before the VSG-BarSeq experiment. After cDNA synthesis and library amplification with ONT-barcoding primers, we recommend analysis of the product in agarose gel. We often obtain a PCR fragment smear ranging from 200 to 2,000 bp (Figure 3B).
Figure 3. Analysis of variant surface glycoproteins (VSG) switching after PIP5Pase knockdown using VSG-BarSeq. (A) Diagram of bloodstream-form expression sites (ESs) (BES) and sub-telomeric regions containing VSG genes used for recombination. The pink arrowhead represents a BES promoter. VSG genes are transcribed from BESs only. (B) Thirty-five cycles of quality-control PCR amplification of pooled clonal VSG barcoded cDNAs. PCR was amplified with ONT barcode primers. NC: negative control, M: 5 kb DNA ladder. (C) VSG genes expressed by clones of T. brucei without PIP5Pase knockdown (Tet +) and after temporary knockdown for 24 h followed by PIP5Pase re-expression and cloning for 5–7 (Tet -/+). Annotated names and corresponding chromosomes or BES identify VSG genes. The number of clones analyzed is indicated in parentheses. (D) Read coverage plots of example clones from Tet + or Tet -/+ treatment groups. The diagram on the top right summarizes the experiment and results—all clones derived from an original clone expressing VSG2 (BES 1). Graphs show read coverage of expressed VSGs on the same scale. (E) Signal-to-noise ratios of two different VSG-BarSeq libraries differing in sequencing depth. Sequencing depth is shown above plot bars as the total reads by the mean read length. Mb: megabases.
After library preparation, we usually obtain a library concentration ranging between 10 and 40 ng/μL with 260/280 and 260/230 ratios of ~1.8 and 2.0, respectively. Library concentrations below 5 ng/µL with considerable deviations from the above 260/280 and 260/230 ratios usually result in inferior quality and low throughput sequencing. Analysis of the sequencing using the vsg-barseq.sh script typically results in 70%–99% genome mapping. Figure 3C and 3D show the results of VSG switching in T. brucei after the temporary knockdown of PIP5Pase. Analysis of 117 clones from the control (no knockdown) showed that none of the cells switched VSGs (Figure 3C). However, after 24 h temporary knockdown, there were 93 switchers out of 94 clones analyzed. There was a preference for the cells to switch to VSG8 in the BES12, suggesting transcriptional switching. Moreover, switching to VSGs from sub-telomeric regions (Chr2_5A and Chr9_3A) was also detected, indicating switching by recombination (Figure 3C and D). The analysis does not require a significant amount of RNAs or sequencing throughput since the experimental setup relies on selecting VSG mRNAs, which are highly abundant (Cestari and Stuart, 2015), and on the analysis of clones rather than heterogeneous cell populations. Analysis of the expressed VSG (signal) compared to other genes (noise) showed a high signal-to-noise ratio, and increasing sequencing depth improved the signal without significantly increasing noise levels (Figure 3E). Although DNA sequencing can be costly, the amount of total RNA for cDNA synthesis per clone in this protocol is minimal (5–20 ng), and sequencing depth required for analysis can be obtained from Oxford nanopore flongle flow cells, which typically produces 500–1,000 megabases of DNA sequencing (i.e., ~100,000 reads per group), thus reducing experimental costs. The results show the method's utility in identifying VSG switching events using multiplexed clonal cell barcodes. We anticipate that, with minimal modifications of the primers used for the target sequences, the approach can be applied to other gene families or other cell types, including var genes in Plasmodium sp., variant surface proteins in Giardia sp., as well as mucins and mucin-associated proteins in Trypanosoma cruzi. Analogously, the approach can be extended to other organisms or cell lines, including mammalian cells, e.g., T-cell or B-cell repertoire analysis.
Validation of protocol
This protocol has been used and validated in the following article:
Touray et al. (2023). A PI(3,4,5)P3-dependent allosteric switch controls antigenic variation in trypanosomes. eLife 12: RP89331 (Figure 1F, and Figure 1—figure supplement 1, panel A, B, C, D).
General notes and troubleshooting
General notes
If more than 40% of the wells in a 96-well plate are positive, the parasites might not be clonal. We recommend optimizing the dilution of the parasites to obtain approximately a third of the wells from a 96-well plate containing growing parasites.
We recommend performing a quality control PCR prior to cDNA amplification. Use the same conditions indicated in step D3 but for 35 cycles. Then, analyze 30 µL the amplicons on 1% agarose/TAE electrophoresis gel with 5 µL of Ecostain at 85 Volts for 45 min and visualize using a gel imager. A smear migrating at 200–2,000 bp should be expected in the agarose gel (Figure 3B).
We recommend the volume of pooled cDNA sample added to the PCR reaction to be less than one-tenth of the total PCR reaction volume.
Reverse transcriptase enzymes are known to inhibit PCR, particularly at low template concentrations (Chandler et al., 1998). Therefore, adding more cDNA to the PCR reactions usually results in non or very little amplification.
The split summary file helps to identify the splitting of reads into multiple barcode fastq files. If the number of reads with barcodes is larger than the total number of reads, it is indicative that some reads are included in more than one barcode fastq file. Primers with longer barcodes than eight mers could be used.
Acknowledgments
This research was funded by the Canadian Institutes of Health Research (grant CIHR PJT-175222 to IC), the Natural Sciences and Engineering Research Council of Canada (grant RGPIN-2019-05271 to IC), the Fonds de Recherche du Québec - Nature et technologie (grant 2021-NC-288072 to IC), and the Canada Foundation for Innovation (grant JELF 258389 to IC). AOT is a recipient of the Islamic Development Bank Scholarship (grant 600042744). This research was enabled in part by computational resources provided by Calcul Quebec (https://www.calculquebec.ca/en/) and the Digital Research Alliance of Canada (alliancecan.ca). This protocol is derived from the original research paper by Touray et al. (2023).
Competing interests
The authors declare that they have no conflicts of interest with the contents of this article.
References
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Supplementary information
The following supplementary information can be downloaded here:
Table S1. List of primer sequences indicated in the VSG-BarSeq protocol.
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Microbiology > Microbial cell biology > Cell-based analysis
Molecular Biology > DNA > DNA sequencing
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Original Research Article:
The authors used this protocol in Frontiers in Immunology Feb 2023
Abstract
Advanced immunoassays are crucial in assessing antibody responses, serving immune surveillance goals, characterising immunological responses to evolving viral variants, and guiding subsequent vaccination initiatives. This protocol outlines an indirect ELISA protocol to detect and quantify virus-specific antibodies in plasma or serum after exposure to viral antigens. The assay enables the measurement of IgG, IgA, and IgM antibodies specific to the virus of interest, providing qualitative and quantitative optical densities and concentration data. Although this protocol refers to SARS-CoV-2, its methodology is versatile and can be modified to assess antibody responses for various viral infections and to evaluate vaccine trial outcomes.
Key features
• This protocol builds upon previously described methodology [1] explicitly tailored for SARS-CoV-2 and broadens its applicability to other viral infections.
• The protocol outlines establishing antibody responses to SARS-CoV-2 infections by determining optical densities and concentrations from blood plasma or serum.
Graphical overview
Summary of the conventional ELISA (A) vs. sensitive ELISA (B) procedures. In both A and B, wells are coated with a capture antigen, such as the spike protein, while in (C) they are coated with human Kappa and Lambda capture antibodies. For the conventional ELISA (A), wells with immobilised capture antigens receive serum/plasma containing the target antibody (A1 and B1). This is followed by an HRP-conjugated detection antibody specific to the captured antibody (A2 and B2) and then a substrate solution that reacts with the HRP, producing a colour proportional to the concentration of the antibody in the serum/plasma (A3 and B3). The reaction is halted, and absorbance is measured. In the sensitive ELISA (B), after the serum/plasma addition (A1 and B1), a Biotin-conjugated primary detection antibody is introduced (A2 and B2). Depending on the target antibody, a secondary streptavidin-HRP conjugated detection antibody is added for IgG or IgM (3a) or a poly-HRP 40 detection antibody for IgA (3b). A substrate is introduced, producing a colour change proportional to the antibody concentration (A4 and B4). The reaction is then stopped, and absorbance is measured. In Panel C, wells are coated with human Kappa and Lambda capture antibodies. Serial dilutions of a known antibody standard are introduced. After undergoing the standard ELISA steps, a detection antibody is added, specifically binding to the Ig standard antibody. Subsequently, a substrate solution causes a colour change proportional to the antibody concentration in the serum/plasma. The reaction is halted, and the absorbance of each well is measured. The resulting optical densities from the coated wells form the standard curve, plotting the absorbance against concentrations.
Keywords: ELISA SARS-CoV-2 Viral infections Optical densities Antibody concentrations
Background
The indirect enzyme-linked immunosorbent assay (ELISA) effectively quantifies and assesses antibody responses, making it invaluable for monitoring public health, evaluating vaccination programs, and testing new vaccines. This has particularly been of importance at the height of the COVID-19 pandemic, during which there was continuous emergence of antigenically distinct variants [2]. It was important to track the dynamics of antigen-specific immune responses to detect immune escape in various populations. Despite similar rates of seropositivity being seen among symptomatic and asymptomatic individuals [3], certain scenarios revealed some differences [4]. A less robust humoral response has been connected to mild COVID-19 cases, prompting concerns about a potentially accelerated decline in immunity. Conversely, severe illness has been linked to a prolonged presence of humoral immunity, enduring for up to 12 months after infection [5, 6]. It is essential to understand the evolution of antibody development within sub-Saharan African (SSA) environments, characterised by distinct disease patterns. Monitoring shifts in IgM, IgG, and IgA levels targeting S, receptor binding domain (RBD), and N antigens becomes crucial in SSA, offering insights into diagnostic approaches, public health strategies, and the immunological aspects relevant to vaccine development. We optimised and validated an in-house indirect ELISA to detect and quantify SARS-CoV-2 spike-, nucleoprotein-, and RBD-directed IgG, IgM, and IgA antibodies in a Ugandan cohort largely characterised by mild and asymptomatic COVID-19. For positive controls, 106 specimens were collected from PCR-confirmed positive study participants; 205 pre-pandemic specimens obtained between October 2012 and November 2017 were used as negative control samples to determine optimal cutoff values, maximizing assay sensitivity and prioritizing assay specificity [1, 7]. Consequently, the determined cutoff values are useful in contexts where mild and asymptomatic SARS-CoV-2 infections are the predominant clinical picture. Commercial standards of known antibody concentrations were used to establish assay limits of detection (LOD) and quantitation (LOQ). Therefore, we present parameters that can be followed to determine optical densities and antibody concentrations in different populations and in regions that had different clinical outcomes and suggest that the same can be done in investigations of antibody responses to other infectious diseases and vaccinations, for example in clinical trials of various vaccine candidates. The COVAC self-amplifying RNA vaccine trial in Uganda [8] has demonstrated the successful application of this protocol.
Materials and reagents
Biological materials
Human serum or plasma samples freshly obtained from study participants, heat inactivated at 56 °C for 30 min and centrifuged at 1,000× g for 10 min at 18–25 °C. One should expect approximately 1.8 mL of serum/plasma after centrifugation, with a pellet at the bottom and the supernatant at the top in each tube. The supernatant should be collected and aliquoted into different tubes stored at -80 °C. Once an aliquot is picked and thawed, it should be kept in a 2–8 °C fridge and used within only one week.
Reagents
Recombinant SARS-CoV-2 Nucleocapsid protein(R&D Systems, catalog number: 10474-CV-01M)
Recombinant SARS-CoV-2 Spike protein (R&D Systems, catalog number: 10549-CV-01M)
Recombinant SARS-CoV-2 Spike S2 Subunit protein (R&D Systems, catalog number: 10594-CV-100)
SARS-CoV-2 RBD (S1) S-protein obtained from a pHL-sec expression vector in HEK 293T/17 cells [obtained from the American Type Culture Collection (ATCC), catalogue number: CRL-11268]. pHL-sec expression vector was a kind donation from the Doores lab at King’s College London
Goat anti-human Kappa-UNLB antibody (Southern Biotech, catalog number: 2060-01)
Goat anti-human Lambda-UNLB antibody (Southern Biotech, catalog number: 2070-01)
Purified human IgG (Sigma, catalog number: I2511)
Purified human IgA (Sigma, catalog number: I2636)
Purified human IgM (Sigma, catalog number: I8260)
Goat anti-human IgG-HRP (Sigma, catalog number: A0170 or AP113P)
Anti-human IgM-HRP (Sigma, catalog number: A6907)
Anti-human IgA-HRP(Sigma, catalog number: A0295)
Goat anti-human Fc IgG-Biotin conjugate (Southern Biotech, catalog number: 2048-08)
Goat anti-human IgA-Biotin (Southern Biotech, catalog number: 2050-08)
Goat anti-human IgM-Biotin (Southern Biotech, catalog number: 2020-08)
PBS/Tween Sachets (PBST) (Sigma, catalog number: P3563)
Tween-20 (Sigma, catalog number: P2287)
1× Dulbecco’s phosphate buffered saline (DPBS), CaCl2 (-) MgCl2 (-) (Sigma, catalog number: D8537)
Bovine serum albumin (BSA) (Sigma, catalog number: A3803)
Goat serum (Sigma, catalog number: G9023)
Streptavidin-HRP (R&D Systems, catalog number: DY988)
Tetramethyl benzidine (TMB) substrate (SeraCare, catalog number: 5120-0080 or 52-00-02)
TMB stop solution (SeraCare, catalog number, 5150-0021 or 50-85-06)
Casein blocking buffer (Thermo Scientific, catalog number: 37582)
Poly-HRP-40 (Fitzgerald, catalog number: 65R-S104PHRP)
Solutions
PBST wash buffer (see Recipes)
BSA assay buffer/sample diluent/blocking buffer (see Recipes)
Goat serum assay buffer/sample diluent (see Recipes)
Recipes
Wash buffer
Reagent Final concentration (%) Quantity or Volume
1× DPBS pH 7.4 99.95% 999.5 mL
Tween-20 0.05% 0.5 mL
Total 100% 1,000 mL
Store in a capped bottle at room temperature up to seven days.
BSA assay buffer/sample diluent/blocking buffer
Reagent Final concentration (%) Quantity or Volume
1× DPBS pH 7.4 99.95% 499.75 mL
Tween-20 0.05% 0.25 mL
BSA 1% 5 g
Total 100% 500 mL
Filter the buffer through a 0.22 µm filter unit. Store at 2–8 °C up to seven days.
Goat serum assay buffer/sample diluent
Reagent Final concentration (%) Quantity or Volume
1× DPBS pH 7.4 94% 470 mL
Tween-20 1% 5 mL
Goat serum 5% 25 mL
Total 100% 500 mL
Filter the buffer through a 0.22 µm filter unit. Store at 2–8 °C up to seven days.
Laboratory supplies
U-bottom, 96-well dilution plates (Corning, catalog number: 3690)
Medium binding, flat-well ELISA plates (Greiner Bio-One, catalog number: 655001)
Sealing tape (Thermo Fisher Scientific, catalog number: 15036)
Reservoirs (CoStar, catalog number: 4870)
Pipettes (P2, P5, P20, P100, P200, P300-Multichannel, P1000) (at the discretion of the laboratory)
Pipette tips (10, 20, and 200 µL) (at the discretion of the laboratory)
50 mL centrifuge tubes (Sigma, catalog number: T1943)
Paper towel rolls (at the discretion of the laboratory)
2 mL cryovials (Sigma, catalog number: BR114832-1000EA)
0.22 µm syringe filters (TISCH Scientific, catalog number: SPEC16251)
Equipment
-20 °C or -80 °C freezer (at the discretion of the laboratory)
Benchtop centrifuge (at the discretion of the laboratory)
Plate washer (BioTek Washer, ELx405 UCWVSD)
Plate reader (BioTek Reader, Elx808IU)
Software and datasets
Plate reader software: Gen5TM Microplate Data Collection and Analysis Software (v3.12, 11/23/2021); license CD purchased through BioTek or provided along with the reader
Spreadsheet software: Microsoft Office Excel v16.0, 2016 or later (09/22/2015); requires Microsoft Office license.
Data analysis software: GraphPad Prism v9.3 (GraphPad, 11/15/2021); license obtained through GraphPad
Software programming language; R Version 4.3.1 (21/04/2023); free to use
Procedure
The conventional ELISA is effective for detecting and quantifying antibodies in plasma or serum after substantial antigenic exposure such as after natural infection. Conventional ELISA finds its application in scenarios where the target molecule is anticipated to exist in comparatively higher concentrations. On the other hand, sensitive ELISA proves advantageous in early detection of immune responses or diseases, especially when the levels of specific antibodies or antigens are exceedingly low. When low antibody levels are observed in clinical trials, the sensitive version of the ELISA assay is employed, incorporating the addition of a biotin–streptavidin detection step that is particularly useful for amplifying and detecting background responses to antigens that the conventional ELISA might not discern. Serum should be obtained from blood collected in serum separating tubes (SST) and plasma from blood collected in acid citric dextrose (ACD) tubes to ensure high quality samples that are free of contaminants.
Preparation of serum or plasma samples and reagents
Serum or plasma samples should be heat inactivated for 30 min at 56 °C and centrifuged at 1,000× g for 10 min at 18–25 °C to harvest the supernatant before testing. If the serum or plasma samples cannot be processed immediately, they should be aliquoted and stored at -80 °C to ensure structural integrity and long-term stability of the antibodies.
Critical: The serum/plasma is heat-inactivated to ensure the safety of the operators since this assay is run on the bench and not under a biosafety cabinet. This kills any potential pathogens present in the serum/plasma.
Unless stated otherwise, all procedures are performed at room temperature (20–25 °C) for the conventional ELISA and at 37 °C for the sensitive ELISA, as described later.
A plate plan should be drawn to record the position of samples, controls, and standards in each ELISA plate. An example is illustrated in Figure 1.
Prepare wash buffer as described in Recipe 1 or by dissolving one sachet of PBS/TWEEN in 1 L of distilled water.
Prepare BSA assay buffer for the conventional ELISA or goat serum assay buffer for the sensitive ELISA as described in Recipes 2 and 3, respectively.
All prepared and aliquoted reagents, i.e., all standards and detection antibodies, should be kept in a -20 °C freezer. Once an aliquot is picked and thawed, it should be kept in a fridge and used within only one week. It should be clearly labelled with the date of first opened/prepared.
It is crucial to utilise reagents that are new and still within their specified expiration date.
Coating of medium-binding ELISA plates
Caution: Inconsistent pipetting may undermine accuracy in processes such as sample dilution, bubble formation, or protocol deviations. Ensure proper pipetting techniques during the assay, e.g., controlled up and down pipetting to minimise bubbles. Refer to Problem 2 under the Troubleshooting section.
Coat wells A1–H2 (see Figure 1) of the 96-well medium-binding ELISA plate by adding 50 µL per well of a solution containing both anti-Human kappa and anti-Human lambda capture antibodies diluted in DPBS, at a final dilution of 1:500 at a 1:1 ratio of anti-Human kappa and anti-Human lambda capture antibodies as shown in Table 1. The coated wells will contain the purified antibodies for the standard curve.
Coat the rest of the wells of the 96-well medium-binding ELISA plate by adding 50 µL of antigen [recombinant SARS-CoV-2 Nucleocapsid protein, recombinant SARS-CoV-2 Spike protein, recombinant SARS-CoV-2 Spike S2 Subunit protein, or SARS-CoV-2 RBD (S1) S-protein] prediluted in DPBS to a working concentration of 3 µg/mL, with a separate plate for each protein. The predilution parameters for the antigen depend on the concentration of the stock solution, which could vary from lot to lot. The demonstrated dilution in Table 2 uses 500 µg/mL as the stock concentration.
Note: Medium-binding plates are more cost effective and suitable for this protocol where non-specific binding might interfere with the assay. They are generally easier to wash and are less likely to retain residual proteins. While they offer strong binding, high-binding plates may be more challenging to wash thoroughly due to their higher binding capacity.
Cover the plates with sealing tape and incubate for 2 h at 37 °C in a humidified incubator or overnight at 4 °C.
Note: During the 2 h incubation time, proceed to section C.
Caution: Do not rock the plates during incubation because rocking can interrupt the formation of bonds between the coating antigen/antibody and the surface of the plate.
After the incubation, wash the ELISA plates with five rounds of 200 µL per well of washing buffer using the plate washer, flick the plates over a container to rid it of excess buffer, and tap gently on paper towel to dry the plate. Then, proceed to section D.
Note: When using the plate washer, set up a protocol by entering the volume of washing buffer required for each well and the number of times each well should be washed on each ELISA plate. Place the plate on the plate holder and run the protocol. If an automated washer is unavailable, 5× of manual washing can be done using a multi-channel pipette set at 200 µL.
Table 1. Predilution of anti-human Lambda and Kappa capture antibodies
Anti-Human Kappa (µL) Anti-Human Lambda (µL) DPBS (µL) Total (µL)
3 3 1494 1500
Note: This 1:500 dilution is forone plate. Multiply volumes by the number of plates in the assay.
Table 2. Predilution of SARS-CoV-2 coating antigens
Capture antigen (stock concentration, µg/mL) Working concentration (µg/mL) Volume of antigen (µL) DPBS (µL) Total (µL)
Spike/Nucleoprotein/RBD (500 µg/mL) 3 36 5,964 6,000
Note: This dilution is forone plate. Multiply volumes by the number of plates in the assay.
Preparation of serum/plasma and control samples
Sample preparation should be performed on the day of the assay (diluted samples should not be stored).
Dilute samples in U-bottom dilution plates using BSA assay buffer for the conventional ELISA or goat serum assay buffer for the sensitive ELISA.
For a 1:100 dilution, add 198 µL of assay buffer to the wells of the dilution plate followed by 2 µL of test sample in duplicate wells following the plate plan in Figure 1.
For a 1:50 dilution, add 196 µL of assay buffer to the wells of the dilution plate followed by 4 µL of test sample in duplicate wells following the plate plan in Figure 1.
Prepare positive and negative control samples at the same dilution used for the test samples; if commercial, prepare according to the manufacturers’ recommendations. Positive and negative controls should be prepared using assay buffer in 2 mL cryovials.
Note: For each well of the ELISA plate, 50 µL of samples or controls will be required. Therefore, the number of wells for each control and number of plates in the experiment should be considered in preparation of the required volume of positive or negative controls.
Blocking of medium-binding ELISA plates
Add 200 µL of BSA assay buffer (conventional ELISA) or goat serum assay buffer (sensitive ELISA) to all the wells in each ELISA plate. Seal the plates and incubate for 1 h at room temperature (conventional ELISA) or at 37 °C (sensitive ELISA).
(Critical) Proceed to section E when there are 20 minutes left to the end of the blocking incubation period.
After incubation, wash the plates five times with 200 µL of wash buffer per well.
Preparation of standards
Use purified human IgG, IgA, or IgM to prepare seven serial dilutions of antibody standards in 2 mL cryovials, as shown in Table 3, Table 4, and Table 5. If more than one plate is being run at a particular time, it is more efficient to prepare the standards in cryovials and then pipette them into their respective ELISA plate wells. Using cryovials also ensures proper mixing as one performs the serial dilutions.
Note: If the stock concentrations of the purified antibodies differ from those provided in the tables, the initial dilutions will have to be adjusted to achieve a starting concentration of 1,000 ng/mL.
The IgG and IgA serial dilutions are 10-fold, while the IgM dilution is 5-fold. These parameters can be adjusted to suit particular experimental conditions.
Table 3. IgG standard serial dilution in assay buffer. Adjust the volumes depending on the plate plan and the number of plates.
Initial 1:100 dilution of human IgG
Prepare a 1:100 dilution of human IgG in the appropriate buffer prior to preparing the serial dilutions
Dilution Stock (mg/mL) Stock (µL) Assay buffer (µL) Total (µL)
Human IgG 1:100 4.52 2 198 200
Serial dilution of human IgG following the initial 1:100 dilution
Concentration (ng/mL) IgG Std (µL) Assay buffer (µL) Total (µL) Take over (µL) Left over (µL)
Std curve humanIgG (10-fold dilution)
1,000 20 884 904 90 814
100 90 810 900 90 810
10 90 810 900 90 810
1 90 810 900 90 810
0.1 90 810 900 90 810
0.01 90 810 900 90 810
0.001 90 810 900 - 900
Table 4. IgA standard serial dilution in assay buffer. Adjust the volumes depending on the plate plan and the number of plates.
Initial 1:100 dilution of human IgA
Prepare a 1:100 dilution of human IgA in the appropriate buffer prior to preparing the serial dilutions
Dilution Stock (mg/mL) Stock (µL) Assay buffer (µL) Total (µL)
Human IgA 1:100 2.0 2 198 200
Serial dilution of human IgA following the initial 1:100 dilution
Concentration(ng/mL) IgA Std (µL) Assay buffer (µL) Total (µL) Take over (µL) Left over (µL)
Std curve human IgA (10-fold dilution) 1,000 45 855 900 90 810
100 90 810 900 90 810
10 90 810 900 90 810
1 90 810 900 90 810
0.1 90 810 900 90 810
0.01 90 810 900 90 810
0.001 90 810 900 - 900
Table 5. IgM standard serial dilution in assay buffer. Adjust the volumes depending on the plate plan and the number of plates.
Initial 1:100 dilution of human IgM
Prepare a 1:100 dilution of human IgM in the appropriate buffer prior to preparing the serial dilutions
Dilution Stock (mg/mL) Stock (µL) Assay buffer (µL) Total (µL)
Human IgM 1:100 1.0 2 198 200
Serial dilution of human IgM following the initial 1:100 dilution
Concentration (ng/mL) IgM Std (µL) Assay buffer (µL) Total (µL) Take over (µL) Left over (µL)
Std curve human IgM (5-fold dilution) 1,000 90 810 900 180 720
200 180 720 900 180 720
40 180 720 900 180 720
8 180 720 900 180 720
1.6 180 720 900 180 720
0.32 180 720 900 180 720
0.006 180 720 900 - 900
Addition of samples, controls, and standards
Add 50 µL of sample/standard/control to the appropriate wells on the plate following the plate plan. Seal plates and incubate for 2 h at room temperature (conventional assay) or for 1 h at 37 °C (sensitive assay). A suggested plate plan for the arrangement of samples, controls, and standards is illustrated in Figure 1.
Figure 1. Representative plate plan to arrange standards, samples, and controls. “SPL” represents a sample, “control” represents a negative control, and “+ control” represents a positive control.
Following incubation, wash the plates five times with 200 µL of wash buffer per well.
Notes:
Some of the controls (control 1, 3, and 4) are run in more than duplicates as a way of testing operators as they run the assay. Running controls at different positions can show if there is any cross contamination or inconsistency as an operator pipettes the controls into their respective wells.
You can also choose to have different types of controls, where a plate can have two positive controls; one that is an in-house control and another that is a commercially prepared positive control, e.g., WHO seropositive serum.
In-house controls calibrate the assay specifically for the laboratory’s equipment, reagents, and techniques, while commercial controls act as a standardised reference point that can be used across different laboratories, making results comparable between different lab settings and experiments.
Addition of secondary antibody (detection antibody)
(Critical) The secondary antibody should be prepared 30 min before use. After adding the detection antibody, proceed to section H.
For the conventional assay:
For IgG plates, dilute goat α-human IgG-HRP detection antibody 1: 10,000 in assay buffer.
For IgM plates, dilute human anti-IgM-HRP detection antibody 1:1,000 in assay buffer for the Spike protein and 1:5,000 for the N-protein.
For IgA plates, dilute anti-human-IgA-HRP detection antibody 1:1,000 in assay buffer.
Add 50 µL of the particular detection antibody per well. Seal plates and incubate for 1 h at room temperature. Following incubation, wash the plates five times with PBST using 200 µL per well.
For the sensitive assay:
Add 50 µL of the primary detection antibody, prepared as described below per well. Seal plates and incubate for 1 h at 37 °C.
i. For IgG plates, dilute goat α-human IgG-Biotin detection antibody 1: 15,000 in goat assay buffer.
ii. For IgM plates, dilute goat anti-human IgM-Biotin detection antibody 1:20,000 in goat assay buffer.
iii. For IgA plates, dilute α-human IgA-Biotin detection antibody 1:20,000 in casein buffer.
Following incubation, wash the plates five times with PBST using 200 µL per well. Add 50 µL of the secondary detection antibody prepared as described below per well. Seal plates and incubate for 1 h at 37 °C.
i. For IgA plates, dilute Poly-HRP40 1:20,000 in casein buffer.
Note: For IgA plates, seal plates and incubate at 37 °C in thedarkfor40 min.
iii. For IgG and IgM plates, dilute streptavidin-HRP 1:200 in goat assay buffer.
Following incubation, wash the plates five times with PBST using 200 µL per well.
Setting up the plate reader and Gen5 software
Switch on the reader (Elx808). The reader will perform a system self-test.
Switch on the PC connected to the reader and double-click the Gen5 3.12 Task Manager icon to open. It will show a window as seen in Figure 2.
Create a protocol by clicking on Protocols > Create new as highlighted in Figure 2.
Figure 2. Open Task Manager window on the Gen5 3.12 software
Check Standard protocol and then click OK. The window shown in Figure 3 will be displayed.
Figure 3. Window that appears upon selecting the standard protocol option
Click on the icon for creating or editing the list of steps as shown in Figure 3.
After that, the window shown in Figure 4 will appear. Select the plate type, e.g., 96-well plate. Under Select wells, check At runtime and indicate if the plate you are reading will have a lid or not.
Figure 4. Procedure layout on the Gen5 software
Click on Read, check the desired wavelength (i.e., 450 nm) as shown in Figure 5, and click Ok.
Figure 5. Window where you can enter the desired wavelength for reading the ELISA plate
The selected wavelength will be displayed as seen in Figure 6. Click OK after this.
Figure 6. Selected wavelength for reading the ELISA plates
Double-click Plate layout on the tool builder as shown in Figure 7.
Figure 7. Plate Layout step on the Gen 5 software
Select the appropriate well types as shown in Figure 8 and click Next. Be critical to include the number of assay controls you want in your plate. In this protocol, there are four assay controls used. Click Next after this selection.
Figure 8. Selection of different well types according to the ELISA plate plan (i.e., Figure 1)
Update the blank as appropriate. Enter full name of BLK as BLANK, number of times the blank is repeated (e.g., 2), and the colour code of choice as shown in Figure 9. Click Next after entering this information.
Figure 9. Information to be entered for the blank wells in the plate layout window
Repeat step 11 for the assay controls and samples.
Update the standards as appropriate. As shown in Figure 10, enter full name of STD as the antibody being tested (e.g., IgG), number of times the blank is repeated (e.g., 2), and the colour code of choice. Enter the unit for the standards (e.g., ng) and the dilution factor (e.g., 10) for IgG. Enter the starting concentration in STD1 and click in the next boxes to automatically fill in the rest of the standards. Click Next after entering this information.
Figure 10. Information to be entered for the standard wells in the plate layout window
Click Ok and the plate layout will be displayed as shown in Figure 11.
Figure 11. Plate layout on which you will highlight the different wells of the ELISA plate
Allocate the different sample type onto the plate by highlighting the sample type (e.g., BLK for blank) and selecting the appropriate orientation.
Using the pointer, drag across the required wells for that sample type in the plate layout. Alternatively, click the first well in the series to automatically fill the adjacent wells with the required number of samples for that type in the selected orientation. The plate layout will appear as shown in Figure 12.
Figure 12. Plate layout window, accurately showing the positions of the different well types of the ELISA plate
After this plate layout has been correctly filled, click OK.
Double-click Data reduction on the tool builder shown in Figure 7. The screen as shown in Figure 13 will be displayed.
Figure 13. Data reduction tools in the Gen 5 software protocol
Go to Standard curve (Data In) and set the y-axis to 450. Go to Curve Fit and set it to non-linear regression (Figure 14).
Figure 14. Standard curve requirements for the ELISA plate readout
Double-click on the Report/Export builder and the window shown in Figure 15 will be displayed.
Figure 15. Report/Export Builders window
Click on New export to Excel and under name enter the name of the report output (e.g., Export 1). Check Plate, Automatic, or Custom (if you want to customise your output) and click Ok. The window in Figure 16 will appear.
Figure 16. Contents of report output, Export 1
Check all parameters under Content as shown in Figure 17.
Figure 17. Different parameters under the context subheading of Lab report
Under Workflow, check Auto-execute, Each pate in a separate workbook, and Save after export as seen in Figure 18. Click Next.
Figure 18. Workflow parameters
Under File >File name, enter file name (e.g., <PLATE_ID>). Check Folder and browse the folder where you want your files to be saved, as shown in Figure 19.
Figure 19. Fields for entering the file name and location where you would like your plate readouts to be stored
Click Next and Ok.
To create a customised protocol, click on New export to Excel, as seen in Figure 15. Under Name, enter the name of the report output (e.g., Export 2), check Plate and Custom (Figure 20), and click Next.
Figure 20. Custom parameter when creating a customised protocol
Click Edit Template and this will open an Excel worksheet.
Click on Add-Ins on the toolbar of the Excel spreadsheet. The custom toolbar is displayed as shown in Figure 21.
Figure 21. Custom toolbar in an Excel worksheet
Customise your template as appropriate by clicking on the drop-down and dragging into the worksheet.
Go to Field Group and set the category as plate information, field, and label as “plate ID.”
Go to Matrix and add plate layout.
Go to Matrix and add 450.
Go to Matrix and add concentration.
Go to Table and then Statistics and add 450.
Go to Table and then Statistics and add concentration.
Go to Graph and add standard curve.
Close the Excel and proceed with steps 20–22 above as done in the automatic builder.
Go to File and save this protocol in a folder labelled “Protocols and Experiments” for easy identification.
To create an experiment in the program, open the Gen5 Software.
The task manager box will appear as shown in Figure 22. Choose Experiments from options, then Create using an existing protocol. Select from the list the protocol that you just saved.
Figure 22. Task manager box, highlighting the Experiments section from the options provided
A new Experiment is created. In tool bar, select File > Save As and save with the appropriate name.
In Task bar select Plate, then Add Plates and add the required number of additional plates to be read.
Plates appear in the menu tree as shown in Figure 23.
Double-click Plate 1, followed by Information. Enter the appropriate plate ID and repeat this to label all the other plates.
Figure 23. Menu tree under a new experiment. The pop-up window is used to enter the plate information for each plate, e.g., in this figure, enter the Plate ID for Plate 1.
Use Microsoft Excel to create a list of the sample IDs on each plate and paste it into a notepad file.
Note: You cannot import sample ID lists from .xls files into this software.
Select Sample IDs from the drop-down menu under each plate, as shown in Figure 24.
Figure 24. Sample IDS parameter from the drop-down menu
Import sample IDs from the notepad file. Alternatively, one can enter the sample IDs manually.
Add sample ID list to each plate. Save the experiment with the included changes.
The reader and software are now set up to read the ELISA plates.
Addition of substrate and determination of absorbencies
(Critical) TMB substrate solution should be removed from the fridge at least 1 h before use.
Add 50 µL per well of TMB substrate to the plates and incubate for 3 min in the dark (conventional ELISA) or 5 min (sensitive ELISA) at room temperature in the dark.
Stop the reaction by the addition of 50 µL of stop solution per well.
Read absorbencies at 450 nm using the plate reader and Gen5 software through study customised protocols.
On your experiment, right-click on the plate you want to read. In Task bar, select Plate and then Read Plate 1.
Select all the plates to be read by clicking Select all as shown in Figure 25.
Figure 25. Window where you select the wells to be read using the ELISA reader
Place the plate on the carrier and read the plate by clicking OK, as shown in Figure 26.
Figure 26. Instructions to load the plate onto the carrier in the ELISA reader
Export the results to an Excel worksheet and store the raw data, which will include a standard curve for each plate read. Examples of these standard curves can be seen in Figure 27, Figure 28, and Figure 29.
Figure 27. Example standard curves produced during measurement of IgG optical densities (450 nm) and concentrations in serum/plasma samples. (A) IgG concentrations against the recombinant SARS-CoV-2 Nucleocapsid protein. (B) IgG concentrations against the recombinant SARS-CoV-2 Spike protein. (C) IgG concentrations against the SARS-CoV-2 receptor binding domain (RBD) (S1) S-protein. (D) IgG concentrations against the recombinant SARS-CoV-2 Spike S2 Subunit protein.
Figure 28. Example standard curves produced during measurement of IgM optical densities (450 nm) and concentrations in serum/plasma samples. (A) IgM concentrations against the recombinant SARS-CoV-2 Nucleocapsid protein. (B) IgM concentrations against the recombinant SARS-CoV-2 Spike protein. (C) IgM concentrations against the SARS-CoV-2 receptor binding domain (RBD) (S1) S-protein.
Figure 29. Example standard curves produced during measurement of IgA optical densities (450 nm) and concentrations in serum/plasma samples. (A) IgA concentrations against the recombinant SARS-CoV-2 Nucleocapsid protein. (B) IgA concentrations against the recombinant SARS-CoV-2 Spike protein. (C) IgA concentrations against the SARS-CoV-2 receptor binding domain (RBD)(S1) S-protein.
Data analysis
In this step, data can be analysed using GraphPad Prism version 9.3 or R Version 4.3.1 or higher. Net optical densities and concentrations are determined.
Copy the plate reader output onto one Excel sheet
If more than one plate is being run at a time, merge the reader data from each plate and copy onto one Excel sheet, clearly labelling each set of data with its plate information.
Calculate the net OD values
To obtain the net optical density at 450 nm, subtract the mean OD of the blank wells from the OD of each sample for each plate. This can be done using the code snippet shown in Figure 30. Alternatively, this can be specified in the Gen5 protocol to give net OD values as the raw data.
Calculate the net concentration values
To obtain the net antibody concentrations, multiply the given concentration from the plate reader data by the sample dilution factor. This can be done using the code snippet shown in Figure 30.
Figure 30. Code snippet showing how to calculate net OD values and net concentration values
Note: Because of assay LOD and LOQ, some samples will not have antibody concentrations in the plate readout and this “?????” might appear instead of their concentrations. However, all samples for each plate should have optical densities.
Using the R software mentioned above, for the samples without concentrations (i.e., those with “?????”), those whose net OD is below the assay LOD are assigned a “0” and those above the assay LOQ should be titrated to determine their end-point titer/concentration. This can be done using the code snippet shown in Figure 31.
Figure 31. Code snippet showing R script for handling samples that are below or above the limit of detection (LOD)
A detailed description to determine assay cutoff values, as well as assay LOQ and LOD, using the direct capture by coated antigen and indirect estimation by coated anti-kappa and lambda antibodies is published in Oluka et al. (2023) [1].
Validation of protocol
This protocol has been validated and used in the following research article(s):
Oluka GK, Namubiru P, Kato L, Ankunda V, Gombe B, Cotten M; COVID-19 Immunoprofiling Team; Musenero M, Kaleebu P, Fox J, Serwanga J. Optimisation and Validation of a conventional ELISA and cut-offs for detecting and quantifying anti-SARS-CoV-2 Spike, RBD, and Nucleoprotein IgG, IgM, and IgA antibodies in Uganda. Front Immunol. 2023 Mar 14; 14: 1113194. doi: 10.3389/fimmu.2023.1113194. PMID: 36999017; PMCID: PMC10045470.
Serwanga J, Ankunda V, Sembera J, Kato L, Oluka GK, Baine C, Odoch G, Kayiwa J, Auma BO, Jjuuko M, Nsereko C, Cotten M, Onyachi N, Muwanga M, Lutalo T, Fox J, Musenero M, Kaleebu P; COVID-19 Immunoprofiling Team. Rapid, early, and potent Spike-directed IgG, IgM, and IgA distinguish asymptomatic from mildly symptomatic COVID-19 in Uganda, with IgG persisting for 28 months. Front Immunol. 2023 Mar 16; 14: 1152522. doi: 10.3389/fimmu.2023.1152522. PMID: 37006272; PMCID: PMC10060567.
Nantambi H, Sembera J, Ankunda V, Ssali I, Kalyebi AW, Oluka GK, Kato L, Ubaldo B, Kibengo F, Katende JS, Gombe B, Baine C, Odoch G, Mugaba S, Sande OJ; COVID-19 Immunoprofiling Team; Kaleebu P, Serwanga J. Pre-pandemic SARS-CoV-2-specific IFN-γ and antibody responses were low in Ugandan samples and significantly reduced in HIV-positive specimens. Front Immunol. 2023 Apr 19; 14: 1148877. doi: 10.3389/fimmu.2023.1148877. PMID: 37153598; PMCID: PMC10154590.
Serwanga J, Baine C, Mugaba S, Ankunda V, Auma BO, Oluka GK, Kato L, Kitabye I, Sembera J, Odoch G, Ejou P, Nalumansi A, Gombe B, Musenero M, Kaleebu P. Seroprevalence and durability of antibody responses to AstraZeneca vaccination in Ugandans with prior mild or asymptomatic COVID-19: implications for vaccine policy. Front Immunol. 2023 May 2; 14: 1183983. doi: 10.3389/fimmu.2023.1183983. PMID: 37205095; PMCID: PMC10187141.
Reliability, reproducibility, and robustness:
In the validation of the protocol, all samples were run in duplicates and a coefficient of variation ≤25% was ensured. The standard curves conformed to a goodness of fit of R2 ≥ 0.9 (Figure 5, Table 6, Figure 7) [1].
SARS-CoV-2 specific positive control antibodies CR3022 and CR3009 as well as local plasma samples of known seropositivity were used to monitor the consistency of the assay using Levey-Jennings curves (Figure 8A and 8B). The ODs were consistently within a CV of 25% and two standard deviations of the mean, in line with recommendations of the Westgard-Sigma rules [1].
Assays were repeated on different days and by different operators to ascertain inter-assay and inter-operator precision; all tests conformed to a CV ≤ 25% (Figure 5) [1]
There was inter-site validation of the protocol between the Imperial College London and the Uganda Virus Research Institute, where the same set of samples were tested, and comparable results were obtained as shown in Figure 9 [1].
The above results from the validation of the protocol are published in the open-access article by Oluka et al. 2023 [1]. https://doi.org/10.3389/fimmu.2023.1113194.
General notes and troubleshooting
Limitations
This ELISA protocol is subject to changes in OD values due to environmental temperature fluctuations in the lab and changes in reagent batches such as the coating antigens and secondary antibodies used in the assay.
To ensure assay consistency with different reagent batches, coat ELISA plates with different batches of an antigen, e.g., a newly purchased spike antigen being compared with a spike antigen currently in use, following the procedure described above.
Run a set of both known positive and negative serum/plasma samples on each of these plates to obtain two sets of results from these plates.
Analyse the data using a scatter plot to determine the correlation coefficient and see if it is significant. One can also use a Wilkson test for matched pairs to give a p-value, which will determine if the distribution is the same in both lots. If the p-value is less than 0.05, we conclude that there is a significant difference between the two groups.
The change in the antigenic determinants of the target pathogen requires new assay validation to ensure consistency and generation of new assay quality controls for use.
Troubleshooting
Problem 1: Plates have a high background in the blank wells.
Possible cause(s): Contaminant particles in air or buffer used, plates left too long before reading, or substrate incubation temperature too high.
Solution(s): Coat plates in a biosafety cabinet. Prepare assay buffer in clean environment. Add substrate at room temperature (23 °C) and read plates promptly after adding substrate.
Problem 2: %CV for OD values of sample replicates above 25.
Possible cause(s): Inconsistent pipetting during sample dilution. Bubbles in wells when reading.
Solution(s): Use properly calibrated pipettes and avoid distractions during sample dilution. Ensure controlled up and down pipetting during sample addition. Avoid generating bubbles when adding substrate or stop solution.
Problem 3: Non-sigmoid standard curve.
Possible cause(s): Improper dilution of standards. Contamination of standards. Deterioration of standards.
Solution(s): Follow closely the dilution steps performed according to the protocol. Use appropriate diluent as blank. Minimise freeze-thaw cycles for the standards and change aliquots after a week.
Problem 4: Weak signal for standards or strong positive controls.
Possible cause(s): Deteriorated antibodies in the sample or standards.
Solution(s): Prepare aliquots that should be used within a week and stored at -20 °C.
Problem 5: Unresolved OD values for sample replicate wells.
Possible cause(s): High titres of target antibodies in the samples tested above the quantifiable range for the standard.
Solution(s): Titrate the unresolved sample in triplicates with 2-fold or 3-fold dilution series on a new plate or test unresolved samples at a higher dilution on another plate.
Problem 6: Inconsistent OD values for negative control samples above expected range.
Possible cause(s): Contamination of controls or assay buffer during preparation.
Solution(s): Prepare controls in clean vials.
Problem 7: Very low or very high signals.
Possible causes: Disregarding the incubation periods and temperature instructions at each step of the assay, contamination, cross-reactivity, poor washing of the plates.
Solution(s): If the high signal is uniform within the whole plate:
If it is a single sample and not the whole plate showing high signals, one should consider that that particular sample has a high antibody concentration indicated by the high signal.
One should ensure that the antibodies used in the assay are specific to the target and not cross-reacting with other substances in the sample.
It is advisable that this protocol be optimised multiple times in the lab to ensure consistency and reproducibility of results before it is implemented.
In case the signal is too low:
One can increase the enzyme-substrate incubation time.
One should reevaluate the wash steps to ensure that they are thorough to remove unbound substances, which could interfere with the signal.
Acknowledgments
The authors acknowledge funding by the government of Uganda through the Science, Technology, and Innovation Secretariat-Office of the President (STI-OP), grant number: MOSTI-PRESIDE-COVID-19-2020/15. Validation of the protocol was done at the MRC/UVRI and LSHTM Uganda Research Unit, which is jointly funded by the UK Medical Research Council (MRC) part of UK Research and Innovation (UKRI) and the UK Foreign, Commonwealth and Development Office (FCDO) under the MRC/FCDO Concordat agreement and is also part of the EDCTP2 programme supported by the European Union.
The project in which this protocol has been used is part of the EDCTP2 program supported by the European Union (grant number RIA2020EF-3008-COVAB). Initial specimen collections for the protocol validation were supported by the University of Glasgow GCRF COVID-19 Rapid Response Fund (Uganda COVID-19 Serological Responses UGANCOSER).
This protocol is based on research funded in part by the Bill and Melinda Gates Foundation through the GIISER Uganda Grant Agreement Investment ID INV-036306. The findings and conclusions contained within are those of the authors and do not necessarily reflect positions or policies of the Bill and Melinda Gates Foundation.
The following reagent was produced under HHSN272201400008C and obtained through BEI Resources, NIAID, NIH: Monoclonal Anti-SARS Coronavirus Recombinant Human Antibody, Clone CR3022 (produced in HEK293 cells), NR-52481. The following reagent was obtained through BEI Resources, NIAID, NIH: Monoclonal Anti-SARS Coronavirus Recombinant Human IgG1, Clone CR3022 (produced in Nicotiana benthamiana), NR-52392. The nucleoprotein mAb CR3009 (NIBSC Repository, Product No. 101011) used as a positive control was obtained from the Centre for AIDS Reagents, NIBSC, UK. The first WHO international standard for SARS-CoV-2, RN 20/136, Immunoglobulin, human, S321534 was obtained from the Centre for AIDS Reagents, NIBSC, UK. The first WHO international standard for SARS-CoV-2, Immunoglobulin, human, 20/B770 was obtained from the Centre for AIDS Reagents, NIBSC, UK. A mammalian expression plasmid for the RBD of SARS-CoV-2 was a kind donation by the Doores laboratory at King’s College London. Specimens used in the inter-site cross-validation for the protocol were obtained from Dr. Hannah Cheeseman’s Laboratory at Imperial College London, United Kingdom. 696. This protocol has been validated as outlined in a prior publication [1] and used in multiple subsequent studies [7, 9–11].
Competing interests
The authors declare that this protocol was developed in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Ethical considerations
The use of human participants to obtain samples used in the development of this protocol was reviewed and approved by the Uganda Virus Research Institute (UVRI) Research and Ethics Committee (Ref: GC/127/833) and the Uganda National Council of Science and Technology (Ref: HS637ES). The patients/participants provided their written informed consent to participate.
References
Oluka, G. K., Namubiru, P., Kato, L., Ankunda, V., Gombe, B., Cotten, M., Musenero, M., Kaleebu, P., Fox, J., Serwanga, J., et al. (2023). Optimisation and Validation of a conventional ELISA and cut-offs for detecting and quantifying anti-SARS-CoV-2 Spike, RBD, and Nucleoprotein IgG, IgM, and IgA antibodies in Uganda. Front. Immunol. 14: e1113194.
Faizo, A. A., Alandijany, T. A., Abbas, A. T., Sohrab, S. S., El-Kafrawy, S. A., Tolah, A. M., Hassan, A. M. and Azhar, E. I. (2021). A Reliable Indirect ELISA Protocol for Detection of Human Antibodies Directed to SARS-CoV-2 NP Protein. Diagnostics 11(5): 825.
Alshami, A., Al Attas, R., Anan, H., Al Maghrabi, A., Ghandorah, S., Mohammed, A., Alhalimi, A., Al-Jishi, J. and Alqahtani, H. (2021). Durability of Antibody Responses to SARS-CoV-2 Infection and Its Relationship to Disease Severity Assessed Using a Commercially Available Assay. Front. Microbiol. 12: e770727.
Marchi, S., Viviani, S., Remarque, E. J., Ruello, A., Bombardieri, E., Bollati, V., Milani, G. P., Manenti, A., Lapini, G., Rebuffat, A., et al. (2021). Characterization of antibody response in asymptomatic and symptomatic SARS-CoV-2 infection. PLoS One 16(7): e0253977.
Balachandran, H., Phetsouphanh, C., Agapiou, D., Adhikari, A., Rodrigo, C., Hammoud, M., Shrestha, L. B., Keoshkerian, E., Gupta, M., Turville, S., et al. (2022). Maintenance of broad neutralizing antibodies and memory B cells 1 year post-infection is predicted by SARS-CoV-2-specific CD4+ T cell responses. Cell Rep. 38(6): 110345.
Abraha, I., Eusebi, P., Germani, A., Pasquarelli, E., Pascolini, S., Antonietti, R., Argenti, S., Fioravanti, A., Martini, E., Aristei, L., et al. (2022). Temporal trends and differences of SARS-CoV-2-specific antibody responses in symptomatic and asymptomatic subjects: a longitudinal study from Umbria in Italy. BMJ Open 12(7): e056370.
Serwanga, J., Ankunda, V., Sembera, J., Kato, L., Oluka, G. K., Baine, C., Odoch, G., Kayiwa, J., Auma, B. O., Jjuuko, M., et al. (2023). Rapid, early, and potent Spike-directed IgG, IgM, and IgA distinguish asymptomatic from mildly symptomatic COVID-19 in Uganda, with IgG persisting for 28 months. Front. Immunol. 14: e1152522.
Kitonsa, J., Kamacooko, O., Ruzagira, E., Nambaziira, F., Abaasa, A., Serwanga, J., Gombe, B., Lunkuse, J., Naluyinda, H., Tukamwesiga, N., et al. (2023). A phase I COVID-19 vaccine trial among SARS-CoV-2 seronegative and seropositive individuals in Uganda utilizing a self-amplifying RNA vaccine platform: Screening and enrollment experiences. Hum. Vaccin. Immunother. 19(2): e2240690.
Serwanga, J., Baine, C., Mugaba, S., Ankunda, V., Auma, B. O., Oluka, G. K., Kato, L., Kitabye, I., Sembera, J., Odoch, G., et al. (2023). Seroprevalence and durability of antibody responses to AstraZeneca vaccination in Ugandans with prior mild or asymptomatic COVID-19: implications for vaccine policy. Front. Immunol. 14: e1183983.
Nantambi, H., Sembera, J., Ankunda, V., Ssali, I., Kalyebi, A. W., Oluka, G. K., Kato, L., Ubaldo, B., Kibengo, F., Katende, J. S., et al. (2023). Pre-pandemic SARS-CoV-2-specific IFN-γ and antibody responses were low in Ugandan samples and significantly reduced in HIV-positive specimens. Front. Immunol. 14: e1148877.
Ssali, I., Mugaba, S., Watelo, A. K., Bemanzi, J., Katende, J. S., Oluka, G. K., Ankunda, V., Baine, C., Kato, L., Onyachi, N., et al. (2023). Spike protein is a key target for stronger and more persistent T-cell responses—a study of mild and asymptomatic SARS-CoV-2 infection. Int. J. Infect. Dis. 136: 49–56.
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© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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4,906 | https://bio-protocol.org/en/bpdetail?id=4906&type=0 | # Bio-Protocol Content
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Dissociation and Culture of Adult Mouse Satellite Glial Cells
RT Raquel Tonello
SD Steve Davidson
TB Temugin Berta
Published: Vol 13, Iss 24, Dec 20, 2023
DOI: 10.21769/BioProtoc.4906 Views: 860
Reviewed by: Olga KopachChristian Vaegter Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Brain, Behavior, and Immunity Oct 2023
Abstract
Satellite glial cells (SGCs) are a type of glial cell population that originates from neural crest cells. They ultimately migrate to surround the cell bodies of neurons in the ganglia of the peripheral nervous system. Under physiological conditions, SGCs perform homeostatic functions by modifying the microenvironment around nearby neurons and provide nutrients, structure, and protection. In recent years, they have gained considerable attention due to their involvement in peripheral nerve regeneration and pain. Although methods for culturing neonatal or rat SGCs have long existed, a well-characterized method for dissociating and culturing adult SGCs from mouse tissues has been lacking until recently. This has impeded further studies of their function and the testing of new therapeutics. This protocol provides a detailed description of how to obtain primary cultures of adult SGCs from mouse dorsal root ganglia in approximately two weeks with over 90% cell purity. We also demonstrate cell purity of these cultures using quantitative real-time RT-PCR and their functional integrity using calcium imaging.
Key features
• Detailed and simplified protocol to dissociate and culture primary satellite glial cells (SGCs) from adult mice.
• Cells are dissociated in approximately 2–3 h and cultured for approximately two weeks.
• These SGC cultures allow both molecular and functional studies.
Graphical overview
Dissociation and culture of mouse satellite glial cells
Keywords: Satellite glial cells Glial cells Dorsal root ganglia Pain Chronic pain Cell culture Quantitative real-time RT-PCR Calcium imaging
Background
Satellite glial cells (SGCs) are a major cellular component of the peripheral nervous system, surrounding the cell bodies of sensory neurons in dorsal root (DRG) and trigeminal ganglia (Hanani, 2005). Long thought to be merely supportive of neurons, increasing evidence suggests that SGCs are dynamic regulators of neuronal processing, including pain on the peripheral nervous system (Ji et al., 2013). Pain affects more than 1.5 billion people worldwide, with hundreds of millions suffering from unrelieved chronic pain (Kennedy et al., 2014). Given the overuse of opioid prescriptions to treat chronic pain, new therapeutics are urgently needed (Grosser et al., 2017).
While there is a growing consensus that SGCs play a role in important physiological and pathological pain mechanisms (Hanani and Spray, 2020; Gazerani, 2021; Andreeva et al., 2022) and may provide new therapeutic targets for pain relief, our knowledge of SGC biology is still limited because of the lack of techniques to study these cells in vitro. Previous protocols for the dissociation and culture of SGCs have been generated from neonatal and immature tissues, contained neuronal and non-neuronal cells, or mostly used rats (Chen et al., 2008; Belzer et al., 2010; de Corato et al., 2011; Poulsen et al., 2014; Wang et al., 2019). Recently, there have been publications using cultured SGCs from adult mice (Su et al., 2022; Tonello et al., 2023). However, a detailed and well-characterized protocol for the dissociation and culture of mouse SGCs is still lacking.
Here, we describe a step-by-step protocol used in our recent publication (Tonello et al., 2023) that dissociates adult SGCs from mouse DRG tissues. The protocol results in cultures that are more than 90% pure and take approximately 10–14 days to be suitable for molecular and functional testing. As a proof of concept, we have also characterized these cultures for the transcriptional expression of SGC-enriched genes, such as the endothelin receptor type B (EDNRB), and functional responses to a specific EDNRB agonist by calcium imaging.
Materials and reagents
Biological materials
CD1 mouse (6–12 weeks) (Charles River, catalog number: 022)
Reagents
Hanks’ balanced salt solution (HBSS) without Ca2+/Mg2+ (Thermo Fisher Scientific, catalog number: 14175095)
1 M HEPES (Thermo Fisher Scientific, catalog number: 15630080)
Penicillin-Streptomycin (P/S), 10,000 U/mL (Thermo Fisher Scientific, catalog number: 15140122)
Papain (buffered aqueous suspension, ≥16 units/mg protein) (Millipore Sigma, catalog number: P3125)
Collagenase from Clostridium histolyticum (Millipore Sigma, catalog number: C6885)
Dulbecco’s modified Eagle medium (DMEM), low glucose (Thermo Fisher Scientific, catalog number: 11885084)
Fetal bovine serum (FBS), heat inactivated (Fisher Scientific, catalog number: MT35011CV)
Amphotericin B (Thermo Fisher Scientific, catalog number: 15290026)
Solutions
Note: Prepare all solutions in a laminar flow cell culture hood.
HBSS dissociation solution (see Recipes)
Papain solution (see Recipes)
Collagenase stock (see Recipes)
Collagenase solution (see Recipes)
DMEM culture medium (see Recipes)
Recipes
HBSS dissociation solution (store at 4 °C for up to six months)
Reagent Final concentration Quantity
HBSS n/a 490 mL
HEPES 1% (v/v) 5 mL
P/S 1% (v/v) 5 mL
Total n/a 500 mL
Papain solution (freshly prepared and used)
Reagent Final concentration Quantity
Papain 40 units total 108 μL
HBSS dissociation solution n/a 3 mL
Total n/a 3.108 mL
Collagenase stock (aliquot at 200 μL, and store at -20 °C for up to six months)
Reagent Final concentration Quantity
Collagenase 22.5 mg/mL 100 mg
HEPES 1% (v/v) 44 μL
HBSS n/a 4.4 mL
Total n/a 4.444 mL
Collagenase solution (freshly prepared and used)
Reagent Final concentration Quantity
Collagenase stock 1.5 mg/mL 200 μL
HBSS dissociation solution n/a 2.8 mL
Total n/a 3.0 mL
DMEM culture medium (store at 4 °C for up to six months)
Reagent Final concentration Quantity
FBS 10% (v/v) 50 mL
P/S 1% (v/v) 5 mL
Amphotericin B 1% (v/v) 5 mL
DMEM, low glucose n/a 440 mL
Total n/a 500 mL
Laboratory supplies
Isoflurane (Henry Schein, catalog number: 1182097)
Spray bottle with 70% ethanol (Fisher Scientific, catalog number: BP82031GAL)
Tissue culture dish (35 × 10 mm) (Fisher Scientific, catalog number: 08-772A)
Microtubes, 1.7 mL (Fisher Scientific, catalog number: 14-222-168)
Centrifugation tubes, 50 mL (Fisher Scientific, catalog number: 06-443-18)
Cell strainer 40 μm (Fisher Scientific, catalog number: 22-363-547)
Cell strainer 10 μm (PluriSelect, catalog number: 43-50010-03)
Cell culture 12-well microplates (Fisher Scientific, catalog number: 07-000-202)
Cover glass (Fisher Scientific, catalog number: 22-050-232)
Equipment
Stereomicroscope system (Olympus, catalog number: SZ51)
Biosafety cabinet (Fisher Scientific, catalog number: NC0986267)
Isotemp water bath (Fisher Scientific, catalog number: FSGPD2S)
Water-jacketed CO2 incubator (Fisher Scientific, catalog number: 13-998-078)
Sorvall Legend Micro 21R microcentrifuge (Thermo Fisher Scientific, catalog number: 75002445)
Centrifuge 5702 (Eppendorf, catalog number: 022628102)
Standard scissors (Fine Science Tools, catalog number: 14002-12)
Forceps (World Precision Instruments, catalog number: 501987)
Student spring scissors (Fine Science Tools, catalog number: 91500-09)
Dumont #5 forceps (Fine Science Tools, catalog number: 11251-20)
Gilson (or equivalent) pipettes and tips (P20/P200/P1000) (Fisher Scientific, catalog number: various)
Procedure
Initial preparations
Prior to beginning any procedures, disinfect all surgical tools and working areas with 70% ethanol.
Next, prepare a culture dish by filling it with 4 mL of HBSS dissociation solution (see Recipes) for the initial collection and cleaning of DRG tissues. Prepare two microtubes with 1 mL of HBSS dissociation solution each, where the cleaned tissues will be transferred. Place the culture dish and microtubes on ice at 4 °C.
The papain and collagenase solutions (see Recipes) can also be prepared at this time and kept at 4 °C. Warm these solutions and DMEM culture medium (see Recipes) to 37 °C just before use.
Terminally anesthetize a mouse by placing the animal in a jar/chamber for anesthesia. Add a gauze with 1 mL of isoflurane to the jar, and the mouse will be anesthetized within a minute. To ensure complete anesthesia, check for a lack of response to a pinch of the rear footpad.
Next, use scissors to decapitate the mice in a sink for the purpose of euthanasia and blood removal. This will facilitate the tissue dissection process.
Tissue dissection
Move the mouse to a designed tissue dissection area and liberally spray the back fur with 70% ethanol.
To expose the back muscles, make a large transverse cut in the middle of the back skin using standard scissors. Then, pull the skin in opposite directions to remove all of the back skin.
To remove the back muscles, make two long cuts close to the left and right sides of the spinal column. Then, use forceps to pull away the muscles starting from the rostral end.
To expose the spinal cord and DRGs, perform a laminectomy by removing the top of the vertebral canal. Use spring scissors to cut the bones on both sides of the vertebral canal in a 45° angle to avoid damaging or losing the DRGs. Continue cutting from side to side while lifting the top of the vertebral canal, starting from the rostral end and working your way down to the caudal end of the spinal column.
Next, use forceps to slowly remove the spinal cord in a rostral to caudal direction from the column and expose the DRGs. Discard the spinal cord. The DRGs are located in the dorso-lateral position of the vertebral canal and can be recognized by their round shape and hyaline appearance, which is different from the white color of the attached nerve fibers (Supplementary Figure 1).
Using the stereomicroscope, grasp the dorsal root with #5 forceps, carefully pull it a few millimeters, and cut the spinal nerve (immediately distal to the ganglion) with spring scissors. Next, cut the dorsal root and place the collected DRGs in the culture dish on ice. Typically, an experienced experimenter can collect approximately 40 ganglia per mouse.
If needed, move the culture dish under the stereomicroscope and use a pair of spring scissors and #5 forceps to remove any extra nerves, roots, and connective tissues from the DRGs. Then, transfer the cleaned DRGs equally into two microtubes.
Cell dissociation
Centrifuge the microtubes containing the cleaned DRGs at 200× g for 1 min and remove the HBSS dissociation solution.
Perform the first enzymatic dissociation by adding 1.5 mL of papain solution into the microtubes. Gently flick the microtubes to resuspend the DRGs and incubate them for 20 min in a 37 °C water bath.
Note: Do not resuspend with a pipette or vigorously shake to maintain the integrity of the cells.
Centrifuge the microtubes at 200× g for 1 min and remove the papain solution.
Wash the DRGs with 1 mL of HBSS dissociation solution, centrifuge (200× g for 1 min), and remove the solution.
Perform the second enzymatic dissociation by adding 1.5 mL of collagenase solution into the microtubes. Gently flick the microtubes to resuspend the DRGs and incubate them for 20 min in a 37 °C still water bath.
Note: Do not resuspend with the pipette or vigorously shake to maintain the integrity of the cells.
Centrifuge the microtubes at 200× g for 1 min and remove all the collagenase solution.
Note: From this point forward, the opening of tubes/plates that contain any tissue, cells, media, or reagents should be done in a laminar flow cell culture hood.
Resuspend the DRGs in 1 mL of DMEM culture medium. Centrifuge at 200× g for 1 min and carefully remove all culture solution without disturbing the tissue.
Resuspend the DRGs in 0.5 mL of DMEM culture medium. Perform mechanical dissociation by trituration using a P1000 pipette until the solution becomes cloudy (approximately 15–20 times up and down). During trituration, the tissues should pass through the tip with a little friction initially and progress to passing easily. Be gentle and avoid introducing air bubbles at this step; this is critical.
Add 1 mL of DMEM culture medium to each microtube. Next, transfer the contents of all microtubes into one 50 mL conical tube. Adjust the volume to approximately 5 mL with DMEM culture medium and gently mix the contents by flicking the tube.
Filter the mixture using a 40 μm cell strainer followed by a 10 μm cell strainer. Collect the filtrate each time in a 50 mL tube. If necessary, adjust the volume of the filtrate to 5 mL with DMEM culture medium and resuspend the cells by gently flicking the tube.
Cell plating and growth
Transfer the resuspended cells into a 12-well microplate by pipetting 400 μL of the filtrate into each well containing sterilized cover glasses. Then, incubate the plate in a CO2 cell culture incubator (37 °C, 5% CO2).
Note: During this incubation (24 h), due to the lack of coating of the cover glasses and use of DMEM low-glucose media, SGC cells will attach to the bottom of the dish while the DRG neurons and debris will remain in suspension.
One day after plating the cells, replace the DMEM culture medium with 400 μL of fresh DMEM culture medium to remove cell debris. Always pre-warm the culture medium to 37 °C before use.
SGCs proliferate and develop a spindle-shaped morphology (Figure 1) during the next 10–14 days. To maintain SGCs cultures, replace the DMEM culture medium every 2–3 days.
Validation of protocol
In our research article, we used this protocol to study the role of matrix metalloproteinases in SGCs and pain (Tonello et al., 2023). Here, we present additional evidence that this protocol is both robust and reproducible by assessing these cultures using molecular and functional approaches.
SGC cultures with an average cell purity of over 90%
We assessed the purity of cell culture using immunofluorescence, following the method described in our publication (Tonello et al., 2023). In brief, after 10–14 days, we replaced the DMEM culture medium with a 4% paraformaldehyde solution and incubated it for 15 min at room temperature (RT). Next, we rinsed the cultures with PBS and blocked them with 1% BSA and 0.2% Triton X-100 in PBS (BSA solution). We then incubated them overnight at 4 °C in the BSA solution with the anti-glutamine synthetase primary antibody (GS, rabbit, 1:5,000). Afterward, we incubated the cultures with the anti-rabbit secondary antibodies conjugated to Alexa Fluor 546 (1:1,000) for 1 h at RT in the BSA solution. Finally, we treated them with the universal nuclear staining DAPI in PBS for 1 min at RT. We then removed the cover glasses from the microplate and covered the side with the cultured SGCs with Prolong Gold Antifade Mountant, mounting them on glass microscope slides. Images were acquired using a Keyence BZ-X800 microscope, and cells were manually counted using NIH Image J open-source software (Schindelin et al., 2012). Cell purity was assessed as a percentage, calculated by dividing the number of GS+ cells (i.e., SGCs) by the number of DAPI+ cells (i.e., all cells in the culture) (Figure 1A). Our protocol resulted in SGC cultures with an average cell purity of 91.4% ± 1.1% (Figure 1B). Approximately 9% of the cells positive only for DAPI were considered potential contaminants. Fibroblasts, characterized as flat cells with a large nucleus, were identified as a potential major contaminant (Figure 1A). It should be noted that a reduced expression of GS has been reported in SGC culture (Belzer et al., 2010). Other markers of SGC, such as fatty acid binding protein 7 (Avraham et al., 2020), may be used instead.
Figure 1. Cell purity of satellite glial cell (SGC) cultures. A. Representative image of immunofluorescence of GS protein and DAPI in SGC culture after 10–14 days. Arrows indicate large-nuclei cells (e.g., fibroblasts). B. Percentage of GS+ cells (i.e., SGCs) over DAPI + cells (i.e., all cells). Graph was generated using GraphPad Prism (v. 10.0.2). Mean ± SEM, n = 4 independent SGC cultures.
Cultures show high expression of SGC transcriptional markers
We assessed the transcriptional expression levels of various cell markers using qPCR, following the method described in our publication (Tonello et al., 2023). In brief, after 10–14 days, we replaced the DMEM culture medium with TRIzol reagent to lyse SGCs and isolate the RNA. We extracted the total RNA from the TRIzol reagent using the Direct-zol RNA MiniPrep kit and then assessed the amount and quality using a UV-Vis spectrophotometer. We then converted the RNA into cDNA using a high-capacity cDNA reverse transcription kit. Specific primers for various cell markers and Gapdh (used for normalization) were obtained from PrimerBank (Wang et al., 2012). The primer sequences can be found in Supplementary Table 1. qPCR was conducted using PowerUp SYBR Green Master Mix on a QuantStudio 3 Real-Time PCR System. Relative transcriptional expression ratios were calculated using the Pfaffl method (Pfaffl, 2001). Figure 2 shows that our protocol resulted in cultures with high expression of SGC gene markers such as Glul, Gja1, and Ednrb (Feldman-Goriachnik and Hanani, 2017; Hanani and Spray, 2020), while showing low expression of gene markers for non-myelinated (Scn7a) and myelinated Schwann cells (Mpz and Ncmap), macrophages (Aif1, Itgam, and Cx3xr1), neurons (Nefh and Prph), and fibroblasts (Fgf13 and Fgf9) (Avraham et al., 2020; Jager et al., 2020; Tonello et al., 2020; Chu et al., 2023; Tonello et al., 2023).
Figure 2. Transcriptional analysis of satellite glial cell (SCG) cultures. Quantification of various cell markers using qPCR shows the higher expression of markers for SGCs over markers for Schwann cells (Schw.), macrophages (Macro.), neurons, and fibroblasts (Fibro.). The graph was generated using GraphPad Prism (v. 10.0.2). Mean ± SEM, n = 6 independent SGC cultures.
Cultures exhibit functional response to EDNRB agonist
We assessed the functional integrity of our SGC cultures using calcium imaging, following the method outlined in our publications (Lee et al., 2020; Ford et al., 2021). In brief, after 10–14 days, we replaced the DMEM culture medium and incubated the SGC culture for 45 min at room temperature with a solution of Fura2-AM (3 μg/mL) in DMEM containing 10% FBS and 1% P/S. We then removed the cover glasses from the cell culture microplates and transferred them to a recording chamber containing an external recording solution with the following concentrations: 130 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 30 mM glucose, and 10 mM HEPES. Illuminance signal was acquired using 365/385-nm switching LED controlled by the MetaFluor software (Molecular Devices) on an Olympus BX51 microscope. Previous research shows that endothelin-1 effectively increases intracellular calcium levels in SGCs by activating the endothelin receptor EDNRB (Feldman-Goriachnik and Hanani, 2017). Here, we show that IRL 1620 (10 nM), a potent and selective EDNRB agonist (Takai et al., 1992), induced an intracellular calcium increase in most, if not all, SGCs (Figure 3A and B). This experiment serves as a proof of concept for the functional integrity of our SGC cultures. These cultures may be used in future studies to evaluate and measure calcium responses to various agonists, including neurotransmitters or cytokines (Souza et al., 2013; Afroz et al., 2019).
Figure 3. Calcium responses in satellite glial cell (SGC) cultures. (A) Images of Fura-2 signaling in SGC cultures and (B) representative calcium imaging traces of SGCs (arrows) in response to the EDNRB agonist IRL 1620 (10 nM). Colors correspond to encircled regions of interest in A.
General notes and troubleshooting
SGCs have been identified as a potential therapeutic target for the treatment of chronic pain. However, our understanding of SGC biology is limited due to the lack of techniques to study these cells in vitro. Here, we provide a step-by-step protocol for the dissociation and culture of adult mouse SGCs:
This protocol is easy to implement and takes approximately 2 h for one person to complete the entire dissection and plating process with some practice. Typically, CD1 mice aged 8–12 weeks are used for this protocol. However, this protocol has been successful with mice of different ages and genetic backgrounds.
Dissection of DRG tissues is carried out similarly to a previously published protocol for the production of sensory neurons (Malin et al., 2007). However, we take extra care to remove any attached spinal nerve and dorsal root to minimize potential contamination from Schwann cells (Supplementary Figure 1).
To maximize cell survival and number in culture, we recommend a dissection time of no more than 30–45 min. Tissues should never be left on ice for longer than 1 h.
The growth of SGC cultures on cover glasses is not necessary. Dissociated cells can be plated directly into a 12-well microplate, for example for qPCR analysis.
Cultured SGCs require 10–14 days of incubation to reach a confluency of 60%–80% and acquire a spindle-shaped, bipolar morphology. Longer incubations risk contamination by fibroblasts.
In conclusion, this protocol results in cultures more than 90% pure and suitable for molecular and functional testing. These cultures may be used to better understand the biology of SGCs and to develop new therapeutics for the relief of chronic pain.
Acknowledgments
This work is supported by the National Institutes of Health (NIH) through the grants NS113243 to T.B. and NS130138 to S.D. Partial support was also provided by the University of Cincinnati Gardner Neuroscience Institute Pilot Award. The protocol was first described and validated in our original publication (Tonello et al., 2023), which investigated metalloproteinase signaling in satellite glial cells and its relation to pain.
Competing interests
The authors declare no competing interests.
Ethical considerations
Animals were used in accordance with NIH guidelines and approved by the Institutional Animal Care and Use Committee at the University of Cincinnati.
References
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Belzer, V., Shraer, N. and Hanani, M. (2010). Phenotypic changes in satellite glial cells in cultured trigeminal ganglia. Neuron Glia Biology 6(4): 237–243.
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Chu, Y., Jia, S., Xu, K., Liu, Q., Mai, L., Liu, J., Fan, W. and Huang, F. (2023). Single-cell transcriptomic profile of satellite glial cells in trigeminal ganglion. Front. Mol. Neurosci. 16: e1117065.
de Corato, A., Capuano, A., Currò, D., Tringali, G., Navarra, P. and Dello Russo, C. (2011). Trigeminal satellite cells modulate neuronal responses to triptans: relevance for migraine therapy. Neuron Glia Biology 7: 109–116.
Feldman-Goriachnik, R. and Hanani, M. (2017). The effects of endothelin-1 on satellite glial cells in peripheral ganglia. Neuropeptides 63: 37–42.
Ford, Z. K., Reker, A. N., Chen, S., Kadakia, F., Bunk, A. and Davidson, S. (2021). Cannabinoid Receptor 1 Expression in Human Dorsal Root Ganglia and CB13-Induced Bidirectional Modulation of Sensory Neuron Activity. Front. Pain Res. 2: e721332.
Gazerani, P. (2021). Satellite Glial Cells in Pain Research: A Targeted Viewpoint of Potential and Future Directions. Front. Pain Res. 2: e646068.
Grosser, T., Woolf, C. J. and FitzGerald, G. A. (2017). Time for nonaddictive relief of pain. Science 355(6329): 1026–1027.
Hanani, M. (2005). Satellite glial cells in sensory ganglia: from form to function. Brain Res. Rev. 48(3): 457–476.
Hanani, M. and Spray, D. C. (2020). Emerging importance of satellite glia in nervous system function and dysfunction. Nat. Rev. Neurosci. 21(9): 485–498.
Jager, S. E., Pallesen, L. T., Richner, M., Harley, P., Hore, Z., McMahon, S., Denk, F. and Vægter, C. B. (2020). Changes in the transcriptional fingerprint of satellite glial cells following peripheral nerve injury. Glia 68(7): 1375–1395.
Ji, R. R., Berta, T. and Nedergaard, M. (2013). Glia and pain: Is chronic pain a gliopathy?. Pain 154: S10–S28.
Kennedy, J., Roll, J. M., Schraudner, T., Murphy, S. and McPherson, S. (2014). Prevalence of Persistent Pain in the U.S. Adult Population: New Data From the 2010 National Health Interview Survey. J. Pain 15(10): 979–984.
Lee, S. H., Tonello, R., Choi, Y., Jung, S. J. and Berta, T. (2020). Sensory Neuron–Expressed TRPC4 Is a Target for the Relief of Psoriasiform Itch and Skin Inflammation in Mice. J. Invest. Dermatol. 140(11): 2221–2229.e6.
Malin, S. A., Davis, B. M. and Molliver, D. C. (2007). Production of dissociated sensory neuron cultures and considerations for their use in studying neuronal function and plasticity. Nat. Protoc. 2(1): 152–160.
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Poulsen, J. N., Larsen, F., Duroux, M. and Gazerani, P. (2014). Primary culture of trigeminal satellite glial cells: a cell-based platform to study morphology and function of peripheral glia. Int. J. Physiol. Pathophysiol. Pharmacol 6(1): 1–12.
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.
Souza, G. R., Talbot, J., Lotufo, C. M., Cunha, F. Q., Cunha, T. M. and Ferreira, S. H. (2013). Fractalkine mediates inflammatory pain through activation of satellite glial cells. Proc. Natl. Acad. Sci. U.S.A. 110(27): 11193–11198.
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Tonello, R., Silveira Prudente, A., Hoon Lee, S., Faith Cohen, C., Xie, W., Paranjpe, A., Roh, J., Park, C. K., Chung, G., Strong, J. A., et al. (2023). Single-cell analysis of dorsal root ganglia reveals metalloproteinase signaling in satellite glial cells and pain. Brain Behav. Immun. 113: 401–414.
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Wang, X., Spandidos, A., Wang, H. and Seed, B. (2012). PrimerBank: a PCR primer database for quantitative gene expression analysis, 2012 update. Nucleic Acids Res. 40: D1144–D1149.
Wang, X. B., Ma, W., Luo, T., Yang, J. W., Wang, X. P., Dai, Y. F., Guo, J. H. and Li, L. Y. (2019). A novel primary culture method for high-purity satellite glial cells derived from rat dorsal root ganglion. Neural Regen Res 14(2): 339–345.
Supplementary information
Supporting information can be downloaded here.
Supplementary Figure 1
Supplementary Table 1
Article Information
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© 2023 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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4,907 | https://bio-protocol.org/en/bpdetail?id=4907&type=0 | # Bio-Protocol Content
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This is a correction notice. See the corrected protocol.
Peer-reviewed
Correction Notice: Microscopy and Plate Reader–based Methods for Monitoring the Interaction of Platelets and Tumor Cells in vitro
VT Veeresh Toragall *
EH Elizabeth J. Hale *
KH Kenneth R. Hulugalla
TW Thomas A. Werfel
(*contributed equally to this work)
Published: Nov 20, 2023
DOI: 10.21769/BioProtoc.4907 Views: 219
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After the official publication in Bio-protocol, we noticed a typo in the Background section of our article (https://bio-protocol.org/e4856). Namely, the citation “Wojtukiewiczet al., 2017” in the sentence “Since platelet activation pathways are likely contributors to cancer growth and metastasis, antiplatelet drugs have immense potential for the treatment of cancer metastasis by inhibiting a myriad of events that drive cancer growth and metastasis (Gay and Felding-Habermann, 2011; Wojtukiewiczet al., 2017; Lucotti et al., 2019; Tao et al., 2021)” should be “Wojtukiewicz et al., 2017” instead.
We made that correction and added the following reference to the Reference section:
Wojtukiewicz, M. Z., Sierko, E., Hempel, D., Tucker, S. C. and Honn, K. V. (2017). Platelets and cancer angiogenesis nexus. Cancer and Metastasis Reviews 36(2): 249–262.
Reference
Toragall, V., Hale, E. J., Hulugalla, K. R. and Werfel, T. A. (2023). Microscopy and Plate Reader–based Methods for Monitoring the Interaction of Platelets and Tumor Cells in vitro. Bio-protocol 13(20): e4856. DOI: 10.21769/BioProtoc.4856.
Article Information
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Quantifying Cell Proliferation Through Immunofluorescence on Whole-Mount and Cryosectioned Regenerating Caudal Fins in African Killifish
AG Augusto Ortega Granillo
RS Robert R. Schnittker
WW Wei Wang
AA Alejandro Sánchez Alvarado
Published: Vol 13, Iss 24, Dec 20, 2023
DOI: 10.21769/BioProtoc.4908 Views: 1350
Reviewed by: Anna Sloutskin Ivonne SehringJohn W Peterson
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Original Research Article:
The authors used this protocol in Science Sep 2020
Abstract
The African killifish Nothobranchius furzeri is an attractive research organism for regeneration- and aging-related studies due to its remarkably short generation time and rapid aging. Dynamic changes in cell proliferation are an essential biological process involved in development, regeneration, and aging. Quantifying the dynamics of cell proliferation in these contexts facilitates the elucidation of the attendant underlying mechanisms. Whole-mount and cryosectioning sample preparation are the preferred approaches to investigate the distribution of cellular structures, cell–cell communication, and spatial gene expression within tissues. Using African killifish caudal fin regeneration as an example, we describe an efficient and detailed protocol to investigate cell proliferation dynamics in both space and time during caudal fin regeneration. The quantification of cell proliferation was achieved through high-resolution immunofluorescence of the proliferation marker Phospho-Histone H3 (H3P). We focused on the characterization of epithelial and mesenchymal proliferation in three-dimensional space at two regeneration time points. Our protocol provides a reliable tool for comparing cell proliferation under different biological contexts.
Key features
• Elaborates in detail the method used by Wang et al. (2020) to quantify whole-organ mitotic events during tail fin regeneration in vertebrates.
• Enables proliferation analysis of millimeter-sized homeostatic and regenerating tissues.
• Three-day alternative method to whole mount using cryosections.
• Allows automatic quantification using ImageJ macros and R scripts.
Graphical overview
Keywords: Proliferation African killifish Fin regeneration Whole mount Cryosections Immunofluorescence Fluorescence quantification
Background
Understanding the genetic and molecular basis of post-embryonic development in vertebrates, such as regeneration and aging, are long-standing quests in biology. One of the limiting steps in addressing such questions is the relative long generation time of most existing research organisms. As a result, identifying and characterizing phenotypes associated with adult stages is much more time-consuming when compared to embryonic stages. The emerging research organism African killifish, Nothobranchius furzeri, has received increasing attention in the fields of regeneration and aging due to its remarkable biology, including fast sexual maturation time and rapid aging (Genade et al., 2005; Harel et al., 2015; Hu et al., 2020; Wang et al., 2020). Quantifying the dynamic changes of cell proliferation is a critical procedure that has been frequently conducted in many different animal models (Kang and Sánchez Alvarado, 2009; Poleo et al., 2001; Wang et al., 2011).
Currently, cryosection and whole-mount sample preparations are efficient and generally accepted approaches to examine the dynamics of cell proliferation associated with a biological process of interest. The former allows for a quick analysis of a single plane in Z along the tissues, and high-resolution imaging of deep tissue structures is possible using this technique. In contrast, the whole-mount approach provides spatial information in 3D that is missing in cryosections at the expense of reduced ability for high-resolution imaging (usually requires short working distance lenses) due to the thickness of tested samples. Further, whole-mount samples retain all information on cell–cell interactions that are only apparent when looking at the whole structure. Integration of the two approaches makes it possible to characterize local features at high-resolution and perform three-dimensional analysis of biological markers of interest.
Phospho-histone 3 (H3P) has been widely used as a proliferation marker because this post-translational modification is deeply conserved across many species with a large phylogenetic distance (Newmark and Sánchez Alvarado, 2000; Nielsen et al., 2013; Wang et al., 2011). The epitope recognized by commercially available antibodies tends to be cross-reactive among wide ranges of species, making H3P the go-to mitosis marker in many research laboratories. In contrast to other proliferation markers such as Ki67 or EdU, H3P displays higher signal to noise ratios and its sparsity within the tissue allows for robust nuclei segmentation. In this protocol, we use the quantification of cell proliferation in killifish fin regeneration as an example to illustrate the challenges and discuss tips for successful H3P immunofluorescence on both cryosections and whole mounts in terms of tissue penetration, pigment bleaching, clearing, imaging, and data processing.
Materials and reagents
Common between whole mount and cryosections
16% Paraformaldehyde solution (Electron Microscopy Sciences, catalog number: 15710)
Methanol (MetOH) (Sigma-Aldrich, catalog number: 34860)
Monoclonal mouse antibody anti-E-Cadherin (Monoclonal Ecad Ab) (BD Biosciences, catalog number: 610182)
Monoclonal rabbit antibody anti-Phospho-Histone H3 (Ser10) (D2C8) (Monoclonal H3P Ab) (Cell Signaling Technology, catalog number: 3377S)
Polyclonal goat antibody anti-Rabbit IgG H&L Alexa Fluor 647 preadsorbed (Abcam, catalog number: ab150083) or F(ab')2-Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor Plus 647 (Thermo Fisher, catalog number: A48285)
Polyclonal goat antibody anti-Mouse IgG H&L Alexa Fluor 555 preadsorbed (Abcam, catalog number: ab150118) or F(ab')2-Goat anti-Mouse IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa FluorTM Plus 555 (Thermo Fisher, catalog number: A48287)
TWEEN® 20 (Sigma-Aldrich, catalog number: P1379)
100 mm Petri dish (Fisher Scientific, catalog number: FB0875712)
Horse serum, heat inactivated (Thermo Fisher, catalog number: 26050070)
Goat serum (Thermo Fisher, catalog number: 16210072)
Fetal bovine serum (FBS) (Thermo Fisher, catalog number: A5256801)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: 472301)
Sodium azide (NaN3) (Sigma-Aldrich, catalog number: S2002)
Roche western blocking reagent solution (Sigma-Aldrich, catalog number: 11921673001)
Glycerol (Sigma-Aldrich, catalog number: G7893)
YOYO-1 iodide (491/509), 1 mM solution in DMSO (Thermo Fisher, catalog number: Y3601)
30 mL polypropylene jar (Fisher Scientific, catalog number: 02-891A)
MS-222 (Sigma-Aldrich, catalog number: E10521)
Formic acid (Sigma-Aldrich, catalog number: 695076)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S5886)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P3911)
Sodium phosphate dibasic (Na2HPO4) (Sigma-Aldrich, catalog number: S9763)
Potassium phosphate monobasic (KH2PO4) (Sigma-Aldrich, catalog number: P5655)
500 mL bottle top vacuum filter, 0.22 µm pore (Corning, catalog number: 431118)
Whole mount
60 mm Petri dish (Fisher Scientific, catalog number: 08-757-100B)
Dumont forceps (Fisher Scientific, catalog number: NC9518792)
Sterile 50 mL conical tubes (Fisher Scientific, catalog number: 14-432-22)
10 mL pipette (Fisher Scientific, catalog number: 13-678-12E)
Razor blade (Fisher Scientific, catalog number: 12-640)
Transfer pipette (Fisher Scientific, catalog number: 13-711-5B)
Ibidi USA µ-dish coverslip bottom 35 mm high Petri dishes (Fisher Scientific, catalog number: 50-305-805)
White lamp with goose neck (Ikea JANSJÖ 3W lamp)
DER 332 (Sigma-Aldrich, catalog number: 31185)
DER 736 (Sigma-Aldrich, catalog number: 31191)
Isophorone diamine (IPDA) (Sigma-Aldrich, catalog number: 118184)
Hydrogen peroxide (H2O2) solution 30% (w/w) in H2O (Sigma-Aldrich, catalog number: H1009)
Dichloromethane (DCM) (Sigma-Aldrich, catalog number: 270997)
Dibenzyl ether (DBE) (Sigma-Aldrich, catalog number: 33630)
Corning 10 mL polystyrene serological pipette (Fisher Scientific, catalog number: 07-200-12)
Cryosections
30% Sucrose solution (Sigma-Aldrich, catalog number: S0389). Store the solution at 4 °C
FalconTM 15 mL conical centrifuge tubes (Fisher Scientific, catalog number: 14-959-49B)
FisherbrandTM disposable base molds (Thermo Fisher, catalog number: 22-363-552)
SuperFrost Plus microscope slides (Avantor, VWRI631-0108)
Super PAP pen (Thermo Fisher, catalog number: 008899)
TritonTM X-100 solution (Sigma-Aldrich, catalog number: 93443)
Reagents for bleaching:
H2O2 (Sigma-Aldrich, catalog number: 88597)
Formamide (Sigma-Aldrich, catalog number: 11814320001)
Optimal cutting temperature (OCT) embedding compound (SAKURA, catalog number: 4583)
TrueBlack lipofuscin (Biotium, catalog number: 23007)
ProLong Gold antifade mountant (Thermo Fisher Scientific, catalog number: P36930)
Tsugar memory miniature paint brushes (Amazon, catalog number: ASIN B0BJQ2CC8D)
DAPI solution 1 mg/mL (Thermo Fisher Scientific, catalog number: 62248)
200 proof ethanol (Sigma-Aldrich, catalog number: E7023)
Solutions
PBS-Tween (see Recipes)
Fixative solution (see Recipes)
10% Tween 20 (see Recipes)
PBS-Tween-NaN3 (see Recipes)
123 mM NaN3 (see Recipes)
Bleaching solution (see Recipes)
25% MetOH (see Recipes)
50% MetOH (see Recipes)
75% MetOH (see Recipes)
Blocking solution (see Recipes)
Primary antibody solution (see Recipes)
Secondary antibody solution (see Recipes)
Secondary antibody stock solution (see Recipes)
Nuclear dye solution (see Recipes)
DCM-MetOH (see Recipes)
DAPI solution (see Recipes)
Phosphate-buffered saline (PBS 1× pH 7.4) (see Recipes)
TrueBlack solution (see Recipes)
Recipes
PBS-Tween
Reagents Final concentration Quantity Unit
10% Tween 20 (Recipe 3) 0.1% (0.89 mM) 10 mL
PBS 1× pH 7.4 1× 990 mL
Final volume 1,000 mL
Note: No need to adjust pH.
Fixative solution
Reagents Final concentration Quantity Unit
16 % paraformaldehyde 4% (1.17 M) 10 mL
Formic acid 0.5% (132 mM) 200 μL
PBS-Tween (1×) (Recipe 1) 1× 29.8 mL
Final volume 40 mL
Note: No need to adjust pH.
10% Tween 20
Reagents Final concentration Quantity Unit
Tween 20 10% (89 mM) 10 mL
dH2O 90 mL
Final volume 100 mL
Note: Filter solution (0.22 μm pore filter). Store at 4 °C. No need to adjust pH.
PBS-Tween-NaN3
Reagents Final concentration Quantity Unit
123 mM NaN3 (Recipe 5) 6.15 mM 500 μL
PBS-Tween (Recipe 1) 9.5 mL
Final volume 10 mL
Note: No need to adjust pH.
123 mM NaN3
Reagents Final concentration Quantity Unit
NaN3 123 mM 400 mg
PBS 1× pH 7.4 50 mL
Final volume 50 mL
Note: No need to adjust pH.
Bleaching solution
Reagents Final concentration Quantity Unit
30% H2O2 6% (1.95 M) 2 mL
MetOH 80% (19.75 M) 8 mL
Final volume 10 mL
Note: Always make a fresh bleaching solution. No need to adjust pH.
25% MetOH
Reagents Final concentration Quantity Unit
MetOH 25% (6.17 M) 12.5 mL
PBS-Tween (Recipe 1) 75% 37.5 mL
Final volume 50 mL
Note: No need to adjust pH.
50% MetOH
Reagents Final concentration Quantity Unit
MetOH 50% (12.35 M) 25 mL
PBS-Tween (Recipe 1) 50% 25 mL
Final volume 50 mL
Note: No need to adjust pH.
75% MetOH
Reagents Final concentration Quantity Unit
MetOH 75% (18.52 M) 37.5 mL
PBS-Tween (Recipe 1) 25% 12.5 mL
Final volume 50 mL
Note: No need to adjust pH.
Blocking solution
Reagents Final concentration Quantity Unit
Horse serum 2.5% 250 μL
DMSO 5% (0.7 M) 500 μL
123 mM NaN3 (Recipe 5) 6.15 mM 500 μL
Roche blocking solution 5% 500 μL
Goat serum 10% 1 mL
PBS-Tween (Recipe 1) 7.25
Final volume 10 mL
Note: No need to adjust pH.
Primary antibody solution
Reagents Final concentration Quantity Unit
Monoclonal H3P Ab 1 in 400 (92.5 ng/mL) 10 μL
Monoclonal Ecad Ab 1 in 200 (1.25 μg/mL) 20 μL
DMSO 5% (0.7 M) 200 μL
123 mM NaN3 (Recipe 5) 6.15 mM 200 μL
FBS 10% 400 μL
PBS-Tween (Recipe 1) 3.17 mL
Final volume 4 mL
Note: No need to adjust pH.
Secondary antibody solution
Reagents Final concentration Quantity Unit
Polyclonal Anti-Mouse AF555 Ab stock (Recipe 13) 1 in 1,000 (2 μg/mL) 10 μL
Polyclonal Anti-Rabbit AF647 Ab stock (Recipe 13) 1 in 800 (2.5 μg/mL) 8 μL
DMSO 5% (0.7 M) 200 μL
123 mM NaN3 (Recipe 5) 6.15 mM 200 μL
FBS 10% 400 μL
PBS-Tween (Recipe 1) 3.182 mL
Final volume 4 mL
Note: No need to adjust pH.
Secondary antibody stock solution
Reagents Final concentration Quantity Unit
Polyclonal secondary Ab 1 in 2 (1 mg/mL) 100 μL
Glycerol 50% (6.78 M) 100 μL
Final volume 200 μL
Note: Mix well. Store at -20 °C. No need to adjust pH.
Nuclear dye solution
Reagents Final concentration Quantity Unit
YOYO-1 Iodide 1 in 10,000 (0.1 nM) 1 μL
PBS-Tween (Recipe 1) 10 mL
Final volume 10 mL
Note: The use of cyanine dyes with the iDISCO clearing method is preferred over DAPI (Renier et al., 2014). No need to adjust pH.
DCM-MetOH
Reagents Final concentration Quantity Unit
MetOH 33.3% (13 M) 3 mL
DCM 66.6% (10.4 M) 6 mL
Final volume 9 mL
Note: Always handle DCM in chemical hood. Make fresh every time. No need to adjust pH.
DAPI solution
Reagents Final concentration Quantity Unit
DAPI 1 mg/mL 2 ng/μL (5.7 μM) 1 μL
PBS-Tween (Recipe 1) 499 μL
Final volume 500 μL
Note: No need to adjust pH.
Phosphate-buffered saline (PBS 1× pH 7.4)
Reagents Final concentration Quantity Unit
NaCl 136.89 mM 8 g
KCl 2.68 mM 0.2 g
Na2HPO4, dibasic, anhydrous 10.14 mM 1.44 g
KH2PO4 1.76 mM 0.24 g
dH2O 1,000 mL
Final volume 1,000 mL
Note: Combine ingredients. Adjust pH to 7.4 with 7 N HCl. Bring to final volume with H2O. Autoclave for 20 min.
TrueBlack solution
Reagents Final concentration Quantity Unit
TrueBlack lipofuscin 1× 50 µL
200 proof EtOH 665 µL
dH2O 285 µL
Note: Combine ingredients by vortexing. No need to adjust pH.
Equipment
Shared by both whole mount and cryosections
Open air platform shaker (Eppendorf, New Brunswick, Innova 2000, catalog number: M1190-0000)
Spinning disk confocal microscope (Nikon CSU-W1) or laser scanning confocal microscope (Zeiss, model: LSM 780)
Imaging workstation (65 GB RAM)
Whole mount
Rocking shaker (Fisherbrand, Fisher Scientific, catalog number: 88-861-025)
Pipet controller (Fisherbrand, Fisher Scientific, catalog number: FB14955202)
Cryosections
Cryostat (Leica Microsystems Inc., Leica Biosystems, Leica CM1950)
SlideTray Slide Staining System (Sigma-Aldrich, Z670146-1EA)
Software and datasets
Fiji (https://imagej.net/software/fiji/)
FCS Express 7 Research Edition (De Novo Software, https://denovosoftware.com/)
Procedure
Sample collection and fixation (both whole mount and cryosections)
Prepare 200 mL of 100–150 mg/L MS-222 in fish system water into a glass beaker. Use an appropriate net size to catch the fish from the tank and place it into the beaker. Anesthetize fish for at least 5 min and transfer them to a Petri dish using a plastic spoon to perform fin amputation.
Amputate the caudal fin using a sterile razor blade at the desired position and allow for regeneration to proceed to the chosen time point (Figure 1A and 1B). For simplicity, we recommend amputations perpendicular to the anterior–posterior axis, removing 50% of the area of the fin. Most fins will show robust regeneration in a range from 30% to 70% removed area. Be aware that amputation position will change regeneration speed as reported before (Uemoto et al., 2020).
Euthanize the fish in accordance with animal care protocols approved by your research institution. We recommend the use of hypothermic shock to perform euthanasia. Immerse the fish in a sufficient volume of chilled system water (2–4 °C) to allow for locomotion. Wait until orientation and operculum movements are lost; this may vary depending on fish size and age. Leave the fish for at least 10 min following loss of orientation and cessation of operculum movement before disposing in biohazard waste bag and deposit into a biological waste freezer, following your home institution guidelines.
Pour 20 mL of fixative solution (see Recipe 2) into a 100 mm Petri dish.
Note: Although 0.5% formic acid may be left out of the fixative solution, its inclusion has been shown to improve penetration of whole-mount samples (Guerrero-Hernández et al., 2021).
Collect the whole tail with a fragment of the spine as shown in Figure 1C; this step is important to avoid folding or wrinkling of the fin tissue during fixation.
Figure 1. Caudal fin sample collection prevents folding of the tissue during fixation. A. Anesthetized full size adult male prior to amputation. B. Caudal fin amputation and plane of future sample collection. C. Fixation step in orbital shaker placed in a cold room; a close-up picture of the sample is shown in the top right corner, where ~4 mm of the spine was included during sample collection to prevent the fin from folding during fixation.
Transfer the sample to a Petri dish with fixative solution and incubate at 4 °C overnight or at room temperature for 2 h in an orbital shaker at 50 rpm.
Wash with 20 mL of PBS-Tween (see Recipe 1) three times for 15 min each in an orbital shaker at 50 rpm.
All steps described above are common for both whole-mount and cryosections. For cryosections, skip to section F; otherwise, continue for whole-mount preparation.
Permeabilization and bleaching (whole mount)
Transfer samples to a polypropylene jar.
Permeabilize samples by incubating with 10 mL of PBS-Tween-NaN3 (see Recipe 4) for 12 h in an orbital shaker at 50 rpm at room temperature.
Incubate in 10 mL of 25% MetOH (see Recipe 7) for 4 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of 50% MetOH (see Recipe 8) for 4 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of 75% MetOH (see Recipe 9) for 4 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of 100% MetOH overnight (>12 h) at room temperature in an orbital shaker at 50 rpm.
Transfer samples to a 60 mm Petri dish.
Incubate in 10 mL of bleaching solution (see Recipe 6) for 12 h at room temperature under direct exposure to white light from a white lamp with goose neck, as shown in Figure 2.
Note: Make sure the lid of the Petri dish is properly placed to avoid evaporation and that the lamp is not touching the Petri dish to avoid both heating up the sample and media evaporation.
Figure 2. Hydrogen peroxide methanol bleaching. A. Dehydrated fin before bleaching in the top panel and after bleaching in the bottom panel. B. Setup with lamp and Petri dish with bleaching solution. C. Fin placement for optimal bleaching with the lamp.
Repeat previous step every 12 h with freshly made bleaching solution until desired bleaching; the more pigmented the sample, the more bleaching iterations this will take to dissolve all pigment. For most specimens, two steps of bleaching will suffice (1 day), and three bleaching steps should be sufficient to bleach very pigmented specimens (1.5 days).
Transfer samples to a polypropylene jar.
Incubate in 10 mL of 50% MetOH for 4 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of 25% MetOH for 4 h at room temperature in an orbital shaker at 50 rpm.
Wash with 10 mL of PBS-Tween-NaN3 overnight (>12 h) at room temperature in an orbital shaker at 50 rpm.
Staining (whole mount)
Incubate in 10 mL of blocking solution (see Recipe 10) for 24 h at room temperature in an orbital shaker at 50 rpm.
Wash with 20 mL of PBS-Tween twice for 15 min each at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of primary antibody solution (see Recipe 11) for 24 h at room temperature in an orbital shaker at 50 rpm.
Wash with 20 mL of PBS-Tween four times for 15 min each at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of secondary antibody solution (see Recipe 12) for 24 h at room temperature in an orbital shaker at 50 rpm.
Wash with 20 mL of PBS-Tween four times for 15 min each at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of nuclear dye solution (see Recipe 14) for 24 h at room temperature in an orbital shaker at 50 rpm.
Repeat the previous step until desired nuclear staining has been reached. This may be necessary when working with larger tissues or when staining multiple fins in the same container. In our experience, two rounds of nuclear staining are sufficient to fully stain large tissues with high signal to noise ratio.
Clearing (whole mount)
Incubate in 10 mL of 25% MetOH for 4 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of 50% MetOH for 4 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of 75% MetOH for 4 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of 100% MetOH for 4 h at room temperature in an orbital shaker at 50 rpm.
Fully stained and dehydrated samples can be stored in the freezer at -20 °C in 100% MetOH for extended periods of time (~1 month). It is key that containers are tightly sealed to avoid evaporation and drying of samples in storage.
Incubate in 10 mL of DCM-MetOH solution (see Recipe 15) for 3 h at room temperature in an orbital shaker at 50 rpm.
Incubate in 10 mL of DCM for 15 min at room temperature in an orbital shaker at 50 rpm.
Repeat the previous step at least one more time and until the sample sinks to the bottom of the jar.
Fill a 5 mL Eppendorf tube with 5 mL of DBE.
Transfer the sample to a Kimwipe tissue to remove excess of DCM, transfer to the Eppendorf tube filled with DBE, and incubate in a rocking shaker for 15 min.
Discard DBE and fill with fresh DBE. Leave the sample overnight protected from light.
Mounting (whole mount)
Heat up DER 332 in a 60 °C oven or water bath for at least 30 min; it should look homogeneous and much more liquid than at room temperature.
In a 50 mL conical tube, pour 23 mL of DER 332, 7 mL of DER 736, and 6 mL of IPDA. Be careful to pour all the volumes from the plastic pipette as resin may adhere to the walls of the pipette.
Homogenize the solution by circular hand movements until the solution is clear. You will introduce bubbles to the resin, which is expected (see Figure 3A and 3B).
Centrifuge the conical tube at ≥ 3,000× g for 10 min at room temperature. If bubbles are still present, you may increase the centrifugation speed (see Figure 3C).
With extreme care to avoid making bubbles, pour 10 mL of resin into a polypropylene jar using 10 mL pipettes with very slow pipetting motion only once. Reusing pipettes may result in bubbles being introduced to the resin.
Transfer all contents from the Eppendorf tube into a 100 mm Petri dish. The sample should be transparent after clearing and may be hard to find. If the sample did not come out when pouring the contents of the Eppendorf tube into the Petri dish, use forceps to carefully extract the sample from the tube (see Figure 3D).
With extreme care to avoid making bubbles, use forceps to pick up the sample and gently insert the sample into the resin located in the polypropylene jar (see Figure 3E).
Stir the sample very gently while holding it with forceps to wash away the DBE for a couple of turns, being extremely careful not to make bubbles.
Transfer the sample to the coverslip bottom Petri dish and, with extreme care without making bubbles, place it on top of the coverslip.
Orient the sample according to your imaging needs, pushing the sample to the bottom against the coverslip using forceps so it locates adjacent to the coverslip. This is particularly important to avoid placing the sample too far from the objective working distance.
Pour enough resin to fill the coverslip but maintain part of the sample protruding from the layer of resin. This is important because if the sample is completely submerged in resin it will float over time and get away from the coverslip (see Figure 3F).
Incubate the coverslip bottom Petri dish with the sample in a dark environment at room temperature for 12 h, covering it with the lid to prevent any dust particles from attaching to the resin.
Prepare new resin according to the procedure described above and pour enough volume to completely submerge the sample in resin (see Figure 3G).
Optional: place a coverslip on top of the resin without introducing bubbles; this may be of particular interest for transmitted light imaging.
Cure the coverslip bottom Petri dish with the sample in a dark environment at room temperature for at least three days before imaging (see Figure 3H and 3I).
Figure 3. Mounting sample into coverslip bottom Petri dish provides optimal imaging conditions. A. Conical tube after pouring the three components of the resin. B. Resin after mixing, with the expected bubbles. C. Ready-to-use resin without bubbles after centrifugation. D. Way to locate the sample (blue arrow) by pouring the contents of the Eppendorf tube into a 100 mm Petri dish. E. Sample (blue arrow) immersed in resin previously poured into a 30 mL polypropylene jar; this step washes away any excess DBE from the sample. F. Sample placed in the coverslip bottom dish. G. Full immersion of the sample in the resin poured 12 h after the sample has been sitting at the bottom of the coverslip bottom dish. H and I. Sample ready for imaging with or without reflective light for visualization. All images were taken with a smartphone except for H and I, thus the scale is not comparable across them.
Cryosectioning (cryosections)
Remove excess tissue to facilitate embedding and sectioning orientation.
Soak specimen overnight in 30% sucrose dissolved in 1× PBS at 4 °C in a 15 mL Falcon tube.
Pause point: You may leave this in 30% sucrose at 4 °C for up to a week.
Transfer samples from 30% sucrose to a suitable tissue mold with OCT and equilibrate samples in OCT for 30 min (Figure 4A).
Figure 4. Caudal fin sample collection and cryosectioning preparation. A. Sample placed and oriented into the OCT embedding molds and the mold after OCT is frozen. B. SlideTray slide staining system where subsequent steps are performed.
Transfer samples to a new tissue mold with fresh OCT and position samples according to the sectioning plane under dissecting microscope with a fine brush or probe.
Place the OCT containing tissue mold on dry ice or other freezing medium such as liquid nitrogen (-40 °C to -70 °C) to freeze the samples. OCT is viscous at room temperature but freezes into a solid support below -10 °C.
Note: Liquid nitrogen is not recommended because it may cause tissues or blocks to crack.
Cut sections 10–12 μm thick in the cryostat at approximately -18 °C. If necessary, adjust the temperature of the cryostat chamber ±3 °C according to the tissues under study.
Note: The optimal temperature for obtaining high-quality cryosections may vary from tissue to tissue. A camel hairbrush is recommended to guide the emerging section over the knife blade or flatten the curved section.
Within 30 s of cutting a tissue section, transfer the section with acceptable morphology to a room temperature microscope slide by touching the slide to the tissue. The tissue section will immediately melt onto the slide due to temperature differences.
Note: This must be completed within 30 s of cutting to avoid freeze drying and unwanted morphological changes of the tissue. Poly-L-lysine-coated or silanized slides improve the adherence of the section (for reference, watch 9:40–9:45 clip at https://youtu.be/tA7S75MFdNk?si=7oq-uMaaxVEi1i_e&t=580).
Quickly evaluate tissue preservation/orientation and check your slide under a dissecting microscope. Collect enough sections per slide according to the need or the size of your samples. On most cases, 10–12 sections can be accommodated on a single slide.
Allow the section to air dry onto the slide at room temperature for 2 h to maximize the adherence of collected sections to the slide.
Air-dried slides can be processed immediately, or they can be placed in a box and stored for up to a month in -20 °C. When finished sectioning, you may store the remaining unused tissue block by covering it with a layer of OCT to prevent freeze drying.
Staining and mounting (cryosections)
Use the SlideTray Slide Staining System to perform the following steps on the slides, as shown in Figure 4B.
Circle the tissue with Super PAP pen to avoid liquid leakage outside of the region of interest.
Wash twice with PBS-Tween (see Recipe 1) for 10 min at room temperature. Slides may come from the freezer or right after air drying.
Block with 500 μL per slide of PBS-Tween with 10% FBS and 5% DMSO for 1 h at room temperature.
Wash once with PBS-Tween for 10 min at room temperature.
Incubate with 200 µL of TrueBlack solution for 1 min at room temperature.
Wash twice with PBS (without Tween) for 10 min at room temperature.
Immunolabel with 500 μL per slide of primary antibodies (H3P, 1:400; Ecad, 1:200) PBS-Tween with 10% FBS and 5% DMSO solution for 1 h at room temperature.
Wash twice with PBS-Tween for 10 min at room temperature.
Develop fluorescent signal with 500 μL per slide of secondary antibodies (Rabbit-AF647, 1:400; Mouse-AF555, 1:500) in PBS-Tween with 10% FBS and 5% DMSO for 1 h at room temperature.
Wash three times with PBS-Tween for 10 min at room temperature.
Stain nuclei with 500 μL per slide of nuclear dye solution (see Recipe 14) for 1 h at room temperature. Alternatively, you may use DAPI solution (see Recipe 16) for 15 min at room temperature.
Wash three times with PBS-Tween for 10 min at room temperature.
Pour one drop of ProLong Gold antifade mountant.
Mount coverslip and leave the slide to cure overnight protected from the light.
Imaging (both whole mount and cryosections)
Place the sample on the stage and image according to the fluorophores conjugated to the secondary antibodies.
For volumetric imaging of whole-mount samples, use a 10× air objective on a Nikon spinning disk confocal microscope and collect a slice every 5 μm for the entirety of the sample (between 80 and 120 slices), making sure that the fluorescent signal does not saturate the detection range of the camera (see Figure 5).
Figure 5. Whole-mount sagital view max projection. Top images show mitotic cells stained by H3P-AF647; middle images show epidermis stained by Ecad-AF555; bottom images show YOYO-1 nuclear staining. Left column: representative images from homeostasis uncut caudal fins; middle column: regenerating 1-day-post-amputation samples: right column: regenerating 2-day-post-amputation caudal fins. 400 μm thick confocal stacks were acquired with a 10× air objective on a spinning disk confocal microscope.
For cryosections, use a 40× immersion objective on a point laser scanning confocal microscope and collect a slice every 1 μm for the entirety of the section (10 slices, see Figure 6).
Figure 6. Max projection of coronal-plane cryosections. Top images show mitotic cells stained by Phospho-Histone H3 (H3P)-AF647; middle images show epidermis stained by Ecad-AF555; bottom images show YOYO-1 nuclear staining. Left column corresponds to regenerating 1-day-post-amputation samples; right column shows regenerating 2-days-post-amputation caudal fins. 25 μm thick confocal stacks were acquired with a 40× water objective on a laser scanning confocal microscope.
Data analysis
Data preprocessing
Stitch the stacks together using the Grid/Collection stitching tool from Fiji (https://imagej.net/software/fiji/) located under the tab Plugins/Stitching/Grid/Collection stitching.
Data analysis
Data analysis of H3P staining is commonly done by manual annotation due to its sparse labeling. However, in whole-mount samples and large volumetric imaging, this task becomes extremely time-consuming. To address this problem, thresholding by particle area and fluorescence intensity can be done using Fiji macros to quantify large volumes of tissue. Below, we describe our quantification pipeline, which yields adequate discrimination between H3P stained nuclei and large fluorescent debris coming from secondary antibody aggregates. See Supplementary information (Code folder) for a ready-to-use pipeline for data analysis.
For every Z plane, perform the following steps (macro 01-Make_pH3_csv.ijm, see Figure 7A):
Duplicate the image.
Blur the image with Median filter, 5 pixel radius.
Subtract background with 50 pixel rolling.
Blur the image with Gaussian Blur filter, 4 pixel sigma.
Find Maxima with a prominence of 10 and output Point Selection.
Measure to record X and Y positions of each maxima.
Save coordinates in a csv file.
Close blurred image and return to the original image.
For every maxima, perform the following steps (macro 02-H3P_Intensity_csv.ijm, see Figure 7B):
i. Draw a square around the point coordinate of 40 pixels by 40 pixels.
ii. Duplicate this image.
iii. Optional: save image for future reference.
iv. Blur the image with Median filter, 3 pixel radius.
v. Blur the image with Gaussian Blur filter, 3 pixel radius.
vi. Duplicate the blurred image.
vii. Set Auto Threshold with the "Otsu" algorithm.
viii. Convert to mask.
ix. Run Watershed.
x. Analyze particles with size 10–400 pixels excluding the edges and adding to the ROI manager.
xi. If there are not any particles, discard the coordinate.
xii. Otherwise, select the raw fluorescent image and measure using the particles from the ROI manager. In most cases, it will be one particle but there may be more than one in a cropped image.
Figure 7. Data analysis workflow. A. Maximum projection on the left; a single Z slice in the middle; and maxima overlay on the right. B. Three examples of H3P positive nuclei showing the raw image on the left, the masked signal in the middle, and the merge of both images on the right.
Save the area and intensity measurements for every analyzed particle with the corresponding X, Y, and Z coordinates.
You should have one table per sample with all the particles corresponding to X, Y, Z, Area, and Intensity values.
Split the table by Z steps and iterate through the whole stack, discarding any particles that are within 5 pixels of each other in X and Y and have lower intensity values than the particle in the adjacent Z plane. This helps filter your table to keep only the brightest measurement for each particle (script 03-ReduceZ.R).
Import the filtered table into FCS express (De Novo Software, https://denovosoftware.com/) to draw a gate for positive events according to Area and intensity. It is important that you quantify the noise and have clear separation between noise particles and true signal.
Figure 8 shows the results for automatic H3P quantification in a whole-mount regenerating fin. We observed 12.7% positive events within the particle area and fluorescence intensity gate. This value will change depending on the prominence value when detecting maxima on step B1e. To validate image quantification, we examine representative images from different areas of the plot. Images that show nuclear morphology are included within the gate and the images that display small debris are excluded from analysis. The high event number is key to reveal clear separation between nuclear morphology particles (top right gated events) and debris.
Figure 8. Phospho-Histone H3 (H3P) gating strategy. After particle quantification, a gate is drawn to include particles of nuclear size with high fluorescence intensity. In this example, 12.7% of all particles are within this gate and the rest of the particles are considered background noise. Representative images of the corresponding distribution are shown for validation.
General notes and troubleshooting
Staining intensity may vary significantly even with the exact same reagent concentration and volume due to cell mass differences between tissues. If you stain in parallel large and small tissues, the local concentration of the reagents as they are penetrating through the tissue will introduce technical variability that will result in noticeable differences in fluorescence intensity among true positive signal. An example is evident in Figure 5 in YOYO-1 staining, where the uncut sample is significantly understained compared to the rest of the samples.
Blur pixel sizes may need to be adjusted depending on the image resolution in the steps B1b, B1c, B1d, and B3. For data analysis, we used images with 1.5577 pix/μm.
Acknowledgments
We thank the Howard Hughes Medical Institute and the Stowers Institute for Medical Research for funding to support this work. We also wish to thank C. Guerrero for sharing fixation protocols with us prior to their publication; J. Unruh, B. Slaughter, C. Wood, R. Alexander, S. McKinney, and L. Maddera for advice on image acquisition and image quantification; J. Jenkin, C. Guerrero, and D. Zamora for help with animal care; and Y. Wang, L. Holmes, J. Blanck, J. Haug and A. Box for extensive discussion about staining optimization, reagent titration, and cytometry analysis; all members of the Sánchez Alvarado lab for insightful scientific discussion. This protocol is an updated and detailed version of the protocol used in: Science (2020), DOI: 10.1126/science.aaz3090.
Competing interests
There are no conflicts of interest or competing interests.
Ethical considerations
All African killifish work was performed according to protocols approved by The SIMR Institutional Animal Care and Use Committee (Protocol ID 2022-137).
References
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Supplementary information
The following supporting information can be downloaded here:
Code.zip.
Article Information
Copyright
© 2023 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Developmental Biology > Cell growth and fate > Regeneration
Cell Biology > Cell imaging > Glue impression
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4,909 | https://bio-protocol.org/en/bpdetail?id=4909&type=0 | # Bio-Protocol Content
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A Protocol to Depict the Proteolytic Processes Using a Combination of Metal–Organic Materials (MOMs), Electron Paramagnetic Resonance (EPR), and Mass Spectrometry (MS)
QL Qiaobin Li
ML Mary Lenertz
ZA Zoe Armstrong
AM Austin MacRae
LF Li Feng
AU Angel Ugrinov
ZY Zhongyu Yang
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4909 Views: 454
Reviewed by: Thomas SchmidtNeha Nandwani Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in ACS Applied Materials & Interfaces 2023
Abstract
Proteolysis is a critical biochemical process yet a challenging field to study experimentally due to the self-degradation of a protease and the complex, dynamic degradation steps of a substrate. Mass spectrometry (MS) is the traditional way for proteolytic studies, yet it is challenging when time-resolved, step-by-step details of the degradation process are needed. We recently found a way to resolve the cleavage site, preference/selectivity of cleavage regions, and proteolytic kinetics by combining site-directed spin labeling (SDSL) of protein substrate, time-resolved two-dimensional (2D) electron paramagnetic resonance (EPR) spectroscopy, protease immobilization via metal–organic materials (MOMs), and MS. The method has been demonstrated on a model substrate and protease, yet there is a lack of details on the practical operations to carry out our strategy. Thus, this protocol summarizes the key steps and considerations when carrying out the EPR/MS study on proteolytic processes, which can be generalized to study other protein/polypeptide substrates in proteolysis. Details for the experimental operation and cautions of each step are reported with figures illustrating the concepts. This protocol provides an effective approach to understanding the proteolytic process with the advantages of offering time-resolved, residue-level resolution of structural basis underlying the process. Such information is important for revealing the cleavage site and proteolytic mechanisms of unknown proteases. The advantage of EPR, probing the target substrate regardless of the complexities caused by the proteases and their self-degradation, offers a practically effective, rapid, and easy-to-operate approach to studying proteolysis.
Key features
• Combining protease immobilization, EPR, spin labeling, and MS experimental methods allows for the analysis of proteolysis process in real time.
• Reveals cleavage site, kinetics of product generation, and preference of cleavage regions via time-resolved SDSL-EPR.
• MS confirms EPR findings and helps depict the sequences and populations of the cleaved segments in real time.
• The demonstrated method can be generalized to other proteins or polypeptide substrates upon proteolysis by other proteases.
Graphical overview
Keywords: Protease immobilization Metal–organic materials (MOMs) Electron paramagnetic resonance (EPR) Site-directed spin labeling Mass spectrometry (MS) Tandem MS
Background
Proteolysis is a critical cellular/biochemical process receiving extensive research attention that has found wide applications in industry and biomedicine [1–6]. Most current progress has been focused on the structures [7–9], cleavage sites [10–13], and proteolytic mechanisms of commonly seen proteases [14–17]. Meanwhile, a knowledge gap still remains regarding the molecular level details of the step-by-step, proteolytic actions on the substrate polypeptides/proteins. Bridging this gap is challenging because it requires revealing, in real time, the details of the reaction mixture, a complex ensemble containing peptide pieces with constantly changing lengths and populations. Perhaps the most feasible practice to probe the lengths and populations of the broken peptide sequences of the entire ensemble would be tandem mass spectrometry (MS/MS or MS2) [18–21], which also resolves amino acid sequence [22–24]. However, MS does not conveniently provide time-resolved information due to the change in product size and population as well as the self-degradation of the protease, which may create additional peptide sequences in the reaction mixture in real time, complicating the mass analysis and even the proteolytic kinetics due to the loss in protease over time. Thus, MS is often applied to analyze the final products after a certain stage (or upon completion) of a proteolytic reaction.
Recently, we found it possible to track the proteolytic process using site-directed spin labeling (SDSL) in combination with electron paramagnetic resonance (EPR) spectroscopy, which is sensitive only to the spin-labeled peptide truncations regardless of the presence of the protease and other broken peptide pieces [25]. This is because EPR line width is sensitive to the molecular size of the labeled polypeptides, which can be tracked via time-resolved, two-dimensional (2D) EPR [26–27]. We also found that immobilizing proteases on the surface of metal–organic materials (MOMs) crystals via co-crystallization allowed for the separation of protease from the soluble reaction mixture during a proteolytic reaction, which contains intact and broken peptides, by centrifugation [28–30]. The soluble portion can then be subjected to MS/MS study to determine the peptide sequences of the entire ensemble. Thus, we proposed a combination of MOMs, EPR, and MS to reveal the global (via MS) and local (via EPR) structural information of the peptide substrates in real time upon proteolysis, which ended up with a dynamic movie depicting the cleavage process [25]. As that was the first time that such a combination was applied to study proteases, we believe it is necessary to summarize the practical protocols to carry out such a complicated study. This protocol is based on our recent publication and utilized the data therein to demonstrate the protocol [25].
Materials and reagents
Reagents
Biphenyl-4,4′-dicarboxylic acid (BPDC-COOH) (Sigma-Aldrich, catalog number: 225266)
NaOH (Sigma-Aldrich, catalog number: 221465)
2-Isopropanol (Fisher Scientific, catalog number: A416-4)
CaCl2 (Sigma-Aldrich, catalog number: C4901)
Trypsin (Sigma-Aldrich, catalog number: T1426)
1-Oxyl-2,2,5,5-tetramethyl-∆3-(methanesulfonyloxymethyl)pyrroline (MTSL) (Toronto Research Chemicals, catalog number: O872400)
Acetonitrile (Fisher Scientific, catalog number: A21-200)
3-(N-Morpholino)propanesulfonic acid, 4-Morpholinepropanesulfonic acid (MOPS) (Sigma-Aldrich, catalog number: M1254)
NaCl (Sigma-Aldrich, catalog number: S9888)
4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid, N-(2-Hydroxyethyl)piperazine-N′-(2-ethanesulfonic acid) (HEPES) (Sigma-Aldrich, catalog number: H3375)
Solutions
Spin buffer (pH 6.8) (see Recipes)
HEPES buffer (pH 7.4) (see Recipes)
Recipes
Spin buffer (pH 6.8)
Reagent Final concentration Quantity
MOPS 50 mM 10.46 g
NaCl 25 mM 1.46 g
H2O n/a 1 L
Total n/a 1 L
HEPES buffer (pH 7.4)
Reagent Final concentration Quantity
HEPES 50 mM 11.92 g
NaCl 50 mM 2.92 g
H2O n/a 1 L
Total n/a 1 L
Laboratory supplies
Beaker (100 mL) (Fisher Scientific, catalog number: 02-540N)
Magnetic spin bar (VWR, catalog number: 58948-080)
Buchner funnel (Sigma-Aldrich, catalog number: Z247340)
Side arm flask (Sigma-Aldrich, catalog number: Z740685)
Microcentrifuge tube (Fisher Scientific, catalog number: 02-682-004)
Amicon spin concentrator (Millipore Sigma, catalog number: ACS 501024)
Borosilicate capillary tube (DWK Life Sciences, catalog number: 13-707-47)
Filter paper (Millipore Sigma, catalog number: WHA1093125)
Equipment
Oven, sample drying (Global Industrial, catalog number: T9FB918772)
Stirring hot plate, mixing (Thermo Scientific, catalog number: SP88854100)
Vortex, mixing (VWR, catalog number: 10153-838)
Nutation mixer, incubation (VWR, catalog number: 82009-202)
Centrifuge, separation (Thermo Scientific, catalog number: 75887203)
EPR, dynamic monitoring (Bruker, model: ESC-106)
Mass spectrometer, peptide mass (Water, model: SYNAPT MS)
Ultra performance liquid chromatography (UPLC), peptide separation (ACQUITY UPLC I-class System)
Software and datasets
Software and algorithms
Multi-component for CW EPR spectral simulation (https://sites.google.com/site/altenbach/labview-programs/epr-programs/multicomponent?authuser=0)
Confirm peptide sequence ProteinLynx Global SERVER 3.0.3
Procedure
The key to achieving the full picture of substrate cleavage by a protease is to obtain both the change in the length/population of different substrate segments in real time via EPR and the peptide sequences of the whole ensemble via MS. This requires protease immobilization, followed by EPR and MS measurements to be carried out separately.
Protease immobilization on MOMs
This major step is to immobilize the target protease onto MOMs during the MOM formation process via co-precipitation. We chose Ca-BPDC as the MOM for protease immobilization due to its high reliability and reproducibility when co-crystalizing with enzymes as well as the proper size of co-crystals formed this way [28, 29]. This material is constructed via incubation of CaCl2 and disodium biphenyl-4,4′-dicarboxylate (BPDC-Na2) in aqua condition, an enzyme friendly environment. Here, biphenyl-4,4′-dicarboxylic acid (BPDC-COOH) has to be converted to BPDC-Na2 due to its low solubility in water.
Preparation of BPDC-Na2
Mix 9.69 g (~40 mmol) of BPDC-COOH and 3.20 g (~80 mmol) of NaOH in a molar ratio of 1:2 in 100 mL of double-deionized (dd-) water. Upon immediate mixing, the reaction system should be white and cloudy because of the low solubility of BPDC-COOH in water.
Vigorously stir the mixture at room temperature for 2 h. Over time, a transparent solution will be obtained due to the enhanced solubility of BPDC-Na2.
Slowly add 1 L of cold 2-isopropanol to the mixture to precipitate BPDC-Na2 and collect the obtained BPDC-Na2 by filtration through a standard filter paper.
Wash the collected filtration with cold isopropanol until a final pH of 7.0 (for the flowthrough). Usually, 0.5 L of cold isopropanol is needed.
Dry the obtained BPDC-Na2 at 75 in an oven overnight.
Immobilization of protease on Ca-BPDC
Prepare stock solutions of CaCl2 and BPDC-Na2 in water at 0.5 M and 0.25 M, respectively. Usually, the stock volume is 5–10 mL, which can be stored for a few months.
Disperse 50 μL of 0.25 M BPDC-Na2 into 1 mL of dd-water, followed by the addition of 20 μL of 20 mg/mL protease and 50 μL of 0.5 M CaCl2. The amount of added protease is adjustable, depending on its solubility and activity. Here, we are using trypsin, a widely seen serine protease, as an example.
Immediately mix the reaction system via vigorous vortex for 30 s and incubate at ambient temperature overnight with gentle nutation (~25 RPM nutation rate is suggested).
Wash the obtained protease@Ca-BPDC with dd-water three times. Each wash is done by centrifugation at 19,000× g for 5 min and resuspend it in 1 mL of water. The obtained co-crystal can be stored for weeks at 4 as judged by testing the crystal properties of the formed Ca-BPDC for many weeks (for characterization see step A3).
Caution: One has to confirm the complete removal of unreacted species in the reaction mixture by determining that no protease (by checking the protein UV absorption at 280 nm) or other involved chemicals (by checking MS) are present in the supernatant after the final wash. If needed, more washes can be applied. Depending on protease, the nutation can be carried out in a fridge if self-degradation is serious.
Characterization
Confirm the structure of Ca-BPDC and inclusion of protease by standard MOM characterization approaches, such as confocal fluorescence microscopy, Powder X-ray diffractometer, thermal gravimetric analysis, and scanning electron microscopy.
Caution: Usually, we do not run nitrogen isotherm [also known as the BET (Brunauer, Emmett, and Teller), experiment] on Ca-based MOMs. Because these techniques are well-established, a detailed procedure to run each equipment is not provided here [25].
Spin labeling of the substrate
This major step is to create an EPR sensitive substrate for monitoring the proteolysis process. Spin labeling is required for protein dynamics, and thus protein size studies by EPR for most proteins. T4 phage lysozyme (T4L) is selected in this protocol as an example to facilitate the discussion, due to its well-studied structure and high population of lysine (K) and arginine (R), which are known as the typical cleavage sites of trypsin [31]. We are using the most widely used spin labeling approach, SDSL of proteins via a methanethiosulfonate-based nitroxide labeling reagent developed by Hubbell and coworkers [32, 33]. The basis of this approach is to create a single cysteine mutation at the site of interest, followed by reaction with the labeling reagent. In principle, any substrate protein can be studied this way.
Caution: For cysteine-rich substrates, an alternative labeling strategy is needed (see below) [34].
Expression of substrate cysteine mutants
Depending on the structure of the substrate, multiple single cysteine mutants need to be created to cover most regions of the substrate and monitor their cleavage over time. The general rule is to place at least one cysteine for each secondary structure (for example α-helix, β-strand, and loop), in order to cover the degradation of each segment by a protease. If there are specific regions of the target protein that need special attention in a proteolytic study, more cys mutants can be placed in these regions. The position of the cys mutants should be solvent accessible. For example, in T4L, 44C, 65C, 72C, 89C, 109C, 118C, 131C, and 151C (each “C” indicates the native residue at the corresponding position was mutated to a cysteine) were generated, one at a time, which is sufficient to cover most regions of the protein. The procedures of site-directed mutagenesis are well documented in the literature and thus not repeated here [32]. Verification of each mutation can be carried out via standard sequencing service available commercially. Nowadays, the cost of sequence is easily affordable.
The generated substrate mutants should be over-expressed and purified in appropriate cell lines. In our case, T4L mutants were expressed in E. coli and purified using published procedures [27, 35], which are not repeated here for the conciseness of this protocol.
Verification of the purity of obtained substrate mutants should be confirmed with circular dichroism (CD) spectroscopy, protein gel–electrophoresis, and protein activity if possible. In our case, each T4L mutant was confirmed to have the expected secondary structure, molecular weight, and catalytic activity [27, 32].
Spin labeling of the substrate mutants
The most widely used spin labeling compound for cysteine mutants is the 1-Oxyl-2,2,5,5-tetramethyl-∆3-(methanesulfonyloxymethyl)pyrroline (MTSL), available at Toronto Research Chemicals, Inc. Powder MTSL should be dissolved in acetonitrile with the concentration of 200 mM.
Add 10-fold molar excess of MTSL into purified T4L mutants and incubate at 4 °C overnight. This will yield a spin labeled sidechain typically designated as R1. Usually, the protein concentration depends on its stability. For most proteins, we suggest maintaining a relatively low protein concentration, ca. < tens of micromolar, to avoid aggregation during labeling.
Filtrate the mixture after incubation to remove excess MTSL by using the Amicon spin concentrator (10,000 MWCO cutoff, 50 mL).
Wash the spin-labeled T4L mutants with a spin buffer three times to further remove the excess MTSL. Removal of all unreacted labeling compound should be confirmed by EPR. Typical EPR settings include ~1 mW power, 3300–3400 G scan range, 30 s scan time for each scan, 1 G modulation depth, and 250 video gain. Incomplete removal is often indicated by an additional sharp component on top of the regular EPR spectrum of a labeled protein. Note that because proteolytic reactions also generate short peptides and thus a sharp spectral component, it is critical to completely remove all unreacted labeling reagent (Figure 1).
Figure 1. Representative electron paramagnetic resonance (EPR) spectrum of an intact spin-labeled protein (A) and that of the cleaved pieces with spin labels attached (B). The smaller the cleaved protein piece, the sharper the continuous wave (CW) EPR spectrum.
Caution: For cysteine-rich proteins, spin labeling based on unnatural amino acids and the corresponding labeling reagents need to be employed as detailed in the literature [34, 36]. The selection of the labeling reagent needs additional caution so that relatively more rigid spin labels should be preferred such as the R1p and RX discussed in a review [26]. This way, the narrowing of the continuous wave (CW) EPR spectrum is mainly caused by the cleavage of the substrate protein, maximizing the sensitivity of the CW EPR spectrum to the protein size. It is also critical to completely remove all unreacted labeling reagent.
EPR study of the proteolytic process
Solution state EPR study as a control
An advantage of time-resolved EPR is to monitor the change in the rotational tumbling rate and thus the molecular weight of the labeled substrate protein pieces in the presence of protease in both the solution state and upon the presence of MOMs. Usually, the proteolytic process could take from minutes to hours, depending on the concentrations of the substrates and protease. Thus, a proper selection of concentration is necessary so that the whole process can be revealed without lengthy experimentation time. In addition, a control of free protease activity against the substrate in solution is often needed in order to confirm the operation. Finally, the EPR spectrum of the sole substrate is needed in order to compare with that of the degraded substrate and confirm cleavage. In our case, we monitored trypsin activity upon degradation of spin-labeled T4L in real time by a Bruker ECS-106 Electron Paramagnetic Resonance (EPR) spectrometer equipped with a cavity resonator (W1700750). The spectrum of each spin-labeled T4 mutant with a 1 mg/mL concentration was acquired prior to later steps discussed below.
Dissolve an appropriate volume of protease into HEPES buffer to make a 1.75 mg/mL (10 times of the final concentration) stock solution for the protease. Note that this stock solution should be freshly prepared due to its self-digestion tendency. Also, HEPES buffer is needed to be consistent with the MOM study because MOMs are stable in HEPES buffer.
Prepare spin-labeled T4L substrate to 10 mg/mL with HEPES buffer. Depending on substrate solubility, this stock concentration can be adjusted.
Bring 2 μL of stock protease and 2 μL of 10 mg/mL T4 substrate to 16 μL of HEPES buffer to make a proteolytic system with 0.175 mg/mL and 1 mg/mL final concentrations of protease and T4L, respectively.
Load the mixture into a borosilicate capillary tube and subject immediately for time-resolved EPR study for 2 h with a 5 min interval. We found 2 h to be sufficient to observe the completion of the proteolytic process using the concentrations detailed above.
An observe power of 200 µW, modulation frequency of 100 kHz, and a modulation amplitude of 0.5 G were used for the acquisition of all CW EPR spectra. Note that a narrow modulation amplitude is needed to detect the narrow linewidth of spectral components originated from cleaved, short peptide chains, bearing in mind that the shorter the peptides, the faster their rotational tumbling is, and the narrower their linewidths.
Caution: The concentrations of protease and substrate could vary depending on the system of study and instrument sensitivity. The EPR data at 0 min could not be acquired yet is often assumed that zero degradation occurs upon immediate mixing. A 5 min interval is often more than sufficient to observe the whole process, yet depending on protease and substrate, instrument sensitivity, and rate of reaction, smaller intervals can be applied.
EPR on protease@MOMs to degrade peptide substrates
Centrifuge 800 µL of protease@CaBPDC suspension to collect all composite. Then, resuspend the composite into 20 µL of buffer, which contains 1 mg/mL of spin-labeled T4 substrate.
Load the mixture into a borosilicate capillary tube and subject immediately for time-resolved EPR study for 2 h with a 5 min interval. Loading the mixture of co-crystals and substrate solution is often completed via capillary effects with tubes open in both ends. Upon completion of loading, the end containing the sample should be sealed to prevent leaking during the measurement.
An observe power of 200 µW, modulation frequency of 100 kHz, and a modulation amplitude of 0.5 G was used for the acquisition of all CW EPR spectra.
MS/MS data acquisition
Protease immobilization on MOMs allows for ease of separation from the soluble products and thus MS analysis of the product ensemble without the interference of protease. A major difference from EPR measurements is that the ion concentration in the buffer needs to be minimized in MS studies, which requires a buffer switch via wash–resuspension. The reaction rate will also be reduced as compared with that in buffer. However, although the reaction kinetics can be altered, being able to carry out time-resolved MS study will still offer valuable information such as the sequences of various broken protein pieces on the dynamic picture of the substrate cleavage process, supporting and confirming the findings from EPR. Thus, MS is necessary in this protocol.
Sample preparation for MS in principle does not need to offer the same time resolution as EPR studies due to the slower reaction rate and the difficulty in practical operation. For example, in our case, upon mixing 40 folds of trypsin@Ca-BPDC with T4L (which does not need to be spin labeled) in 120 μL with 1 mg/mL concentration of the composite and T4L, 20 µL of the mixture was removed from the reaction at 30 min, 1 h, 1.5 h, 2 h, 2.5 h, and 3 h, followed by a quick centrifugation (19,000× g for 1 min) and collection of the supernatant.
Each supernatant should be diluted with dd-water to 100 ng/μL to match the most sensitive concentration range for mass spectrometry.
The data acquisition and analysis can be done in a regular modern MS spectrometer. In our case, a Waters SYNAPT MS system controlled by MassLynx 4.2 Software was employed. An ACQUITY I-class UPLC System equipped with 2.1 × 100 mm BEH300 1.7 μm Peptide Separation Technology C18 column was used to separate fragmented polypeptides.
Run each sample with a 20 min gradient (3%–37% B), after 1 min non-gradient flow at initial conditions (3% B). Mobile phase A was 0.1% formic acid (FA) in water and B was 0.1% FA in acetonitrile. The flow rate was 0.2 mL/min, and the column temperature was 65 °C. An auxiliary pump delivered a lockmass solution [100 fmol/μL (GLu1)-fibrinopeptide B (GFP) in 50:50 ACN/water containing 0.1% FA] for mass accuracy reference.
The instrument is operated in the positive ion V-mode. Use an alternating low collision energy (5 V) and elevated collision energy (ramping from 17 to 40 V) acquisition to acquire peptide precursor (MS) and fragmentation (MSE) data. Scan time was 0.5 s (1 s total duty cycle). The capillary voltage was 3.0 kV, source temperature 110 °C, cone voltage 30 V, and cone gas flow 10 L/h. Sampling of the lock spray channel was performed every 1 min.
Process the acquired data with IdentityE Software of ProteinLynx Global SERVER 3.0.3. Search the processed data against the manual database consisting of single protein (T4L in our case) sequence. Primary Digest Reagent will be selected as non-specific, while Cysteine (C) carbamidomethylation and methionine (M) oxidation will be allowed as optional modifications in this search. The average mass of the intact protein can be determined by MassLynx software.
Data analysis
This major step is to depict the processes of proteolysis via spectral analysis of data acquired from EPR and MS. To obtain the population changes of cleaved shorter peptides from EPR data, spectral simulation is needed. Since the simulation procedure has been published, only cautions and specific details are highlighted below [27]. An additional bonus is that by plotting the populations of the cleaved segments of T4L, it is possible to obtain the reaction kinetics and compare the effects of MOMs on protease performance (see below).
Simulate all collected EPR spectra using the Multi-component program available from Prof. Hubbell’s website. Note that the runtime engine of Labview available at National Instrument is needed for software installation. Before simulation, a careful baseline correction and normalization is needed for all spectra. Details of the baseline correction, spectra normalization, and simulation can be found in our protocol on the resolving enzyme orientation and dynamics via SDSL-EPR [32].
A key difference from our previous simulation works is that the two components involved in protease studies via EPR do not have the traditional immobile peak corresponding to very slow and/or highly restriction motion of the spin label. Instead, one component from our simulation originates from the uncleaved substrate; the linewidth and position fall within the classic mobile component, as described in our recent protocol. The other component, originated from the shorter peptides due to cleavage, results in a sharper peak located close to the classic mobile. The software we are using is able to distinguish the two components as long as the sharper component is populated by 5%–10% during cleavage.
Caution: Protease will continue cleaving other peptide bonds when no more K and R residues in the sequence are available for proteolysis. This will make the truncated peptides become shorter and shorter over time. Since the CW EPR spectra are sensitive to sub-ns motions, when the peptides are too short, EPR loses the sensitivity. Thus, our data analysis was only focused on the first 30–40 min. A representative data is shown in Figure 2.
Figure 2. Two-dimensional electron paramagnetic resonance (2D-EPR) data under the optimal trypsin concentration within a 2 h timeframe to degrade 151R1 of T4L using trypsin (a) and trypsin@Ca-BPDC (b)
Plot the mobile population vs. time to reveal the changes in the rotational tumbling rate and population of the spin-labeled protein segment and thus the proteolytic process. At all spin-labeled sites, the sharper mobile population increases over time and then tends to flatten. At the beginning of 40 min, the greater mobile population slope indicates the generation of a shorter fragmented polypeptide (Figure 3).
Figure 3. Plotting the mobile population of each labeled segment of T4L substrate upon reaction with free (left) and Ca-BPDC-encapsulated trypsin (right) for 40 min
The proteolytic kinetics can also be obtained from the plot via fitting with the Michalis-Menten equation. For example, in our case (Figure 4), we found that reduced km indicates an enhanced binding affinity of trypsin@CaBPDC toward 151R1 as compared with free trypsin. The Vmax of trypsin@CaBPDC is lower than that of free trypsin in buffer. However, this does not mean trypsin@CaBPDC has a lower catalytic efficiency. To compare the catalytic efficiencies of enzymes, one often utilizes kcat/km, wherein kcat is the turnover number that is related to Vmax by Vmax = kcat[Etotal]. Based on the enhanced kcat/km, we believe that roughly 5–10 times catalytic efficiency enhancement is offered by entrapping trypsin in Ca-MOM.
Figure 4. Fitting the velocity (V) as a function of substrate concentration to determine the Vmax and Km of free and MOM-encapsulated trypsin under varied substrate (151R1) concentrations
For MS, taking trypsin as the example protease, it is already well known that it hydrolyzes peptide bonds of arginine and lysine at c-terminal side. Also, the amino acids sequence of the substrate is known. The IdentityE Software of ProteinLynx Global SERVER 3.0.3 could result in all fragmented peptide sequences and their intensities. A typical MS data set is shown in Figure 5. The products of proteolysis separated from the reaction mixture contain substrate and trypsin@Ca-BPDC at various time points. The ease of separation between the trypsin@Ca-BPDC composite and the broken peptide pieces in the solution allows the characterization of the ensemble of substrate protein and hydrolyzed peptides. The drop in the intensity of the peak highlighted in the red block confirms the reduction in molecular mass of the protein (Figure 5).
Figure 5. Representative mass spectrometry (MS) data from the products of proteolysis on our trypsin@Ca-BPDC study
Plot the intensities of different polypeptides from MS vs. timeline. A rapid growth rate of a certain polypeptide fragment indicates that the cleavage sites near the fragment are more likely to be captured and broken down by trypsin. Thus, the intensities and slopes monitor the proteolytic process roughly. A representative data set is shown in Figure 6. Most segments show an increased population over time upon cleavage.
Figure 6. Representative mass spectrometry (MS) data at the beginning (1.5 h) of the reaction when trypsin@CaBPDC catalyzed the T4L substrate
Lastly, the plots of population vs. time from EPR and MS need to be compared to confirm the findings. In our case, we found the 66–76 segment is favored to be cleaved by trypsin according to both EPR and MS. A quick way to prove this finding is mutagenesis, wherein 65K was mutated out and we did observe the disappearance of the 66–76. The site at 76 was still cleaved, resulting in the 61–76 truncation, which was generated at a much lower rate, confirming our speculation that 65K sticking out of the protein favors T4L contact with trypsin@Ca-BPDC. The other three dominant truncations were observed with a similar trend for each, as the mutation does not affect these regions.
Validation of protocol
The whole procedure is validated in our recent work and the supplemental information [25].
General notes and troubleshooting
General notes
For proteolytic processes in a faster timescale (<30 min), the protocol should be modified to offer a higher time resolution. It has to be noted that the immobilization can only reduce the chance for two proteases to collide and cleave each other; it cannot completely prevent self-degradation. However, as shown in our recent work, the reusability of the protease@Ca-BPDC composites is very high, indicating that the self-degradation of the immobilized protease is marginal [25].
Additionally, the protease we are using is a commercial trypsin, which is relatively stable under our immobilization process. However, for a general protease that is sensitive to certain metal ions (ca. Ca2+) or ligands, caution should be given when immobilizing it. We recently published a paper on more possible combinations of metal ions and ligands to immobilize enzymes in water, which may help the custom immobilization of a general protease [37].
Troubleshooting (Table 1)
Table 1. Troubleshooting
Trouble Possible cause Suggested solution
Too slow or fast kinetics Improper protease@CaMOMs concentration Adjust concentration of protease@CaMOMs
Low protease activity upon immobilization Self-degradation during co-crystallization Adjust initial concentration of protease; reduce co-crystallization temperature
Acknowledgments
This work is supported by the National Science Foundation (NSF: MCB 1942596 and DMR 2306137). We appreciate Dr. Peter G. Fajer for generously donating the Bruker ECS-106 to our institution (North Dakota State University) and Dr. Wayne Hubbell for generously providing the EPR data analysis software. This protocol is adapted from our recent work (Li et al., 2023).
Competing interests
The authors declare no competing interests.
Ethical considerations
No human subjects are involved in this work.
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4,910 | https://bio-protocol.org/en/bpdetail?id=4910&type=0 | # Bio-Protocol Content
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In vitro Assay to Examine the Function of Tears on Corneal Epithelial Cells
MM Moumita Mondal
MV Mehak Vohra
JS Jyoti Sangwan
SV Sudhir Verma
Vivien J. Coulson-Thomas
AC Arun Chandru
TB Tuhin Bhowmick
VS Virender S. Sangwan
MA Manisha Acharya
AT Anil Tiwari
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4910 Views: 784
Reviewed by: Alberto RissoneAbhishek vats Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Pathogens Feb 2023
Abstract
Tears contain numerous secreted factors, enzymes, and proteins that help in maintaining the homeostatic condition of the eye and also protect it from the external environment. However, alterations to these enzymes and/or proteins during pathologies such as mechanical injury and viral or fungal infections can disrupt the normal ocular homeostasis, further contributing to disease development. Several tear film components have a significant role in curbing disease progression and promoting corneal regeneration. Additionally, several factors related to disease progression are secreted into the tear film, thereby serving as a valuable reservoir of biomarkers. Tears are readily available and can be collected via non-invasive techniques or simply from contact lenses. Tears can thus serve as a valuable and easy source for studying disease-specific biomarkers. Significant advancements have been made in recent years in the field of tear film proteomics, lipidomics, and transcriptomics to allow a better understanding of how tears can be utilized to gain insight into the etiology of diseases. These advancements have enabled us to study the pathophysiology of various disease states using tear samples. However, the mechanisms by which tears help to maintain corneal homeostasis and how they are able to form the first line of defense against pathogens remain poorly understood and warrant detailed in vitro studies. Herein, we have developed an in vitro assay to characterize the functional importance of patient isolated tears and their components on corneal epithelial cells. This novel approach closely mimics real physiological conditions and could help the researchers gain insight into the underlying mechanisms of ocular pathologies and develop new treatments.
Key features
• This method provides a new technique for analyzing the effect of tear components on human corneal epithelial cells.
• The components of the tears that are altered in response to diseases can be used as a biomarker for detecting ocular complications.
• This procedure can be further employed as an in vitro model for assessing the efficacy of drugs and discover potential therapeutic interventions.
Keywords: Ocular diseases In vitro assay Corneal epithelial cells Tear biomarkers Drug testing
Background
Tears are a vital component of the ocular surface, serving as a first line of defense against pathogens and extreme changes in the environment [1, 2] One of the main functions of tears is to act as lubricant, keeping the eye moist while also protecting it from physical damage [2]. Tears also help to maintain conjunctival and corneal health by providing nutrition, regulating optical properties, and boosting tissue cooling. The unique composition of tears also protects the ocular surface against various bacterial, fungal, and viral infections. The tear film is comprised of three layers: the innermost mucous layer, the middle aqueous part, and the outermost lipid layer. The biological composition of metabolites, electrolytes, mucins, lipids, water, proteins, salts, and organic molecules that are present in tears can be used to distinguish healthy and diseased states [1]. Several studies have shown how tears can be used as an indicator of ocular health and pathology, for example in diabetic retinopathy [3], dry eye disease [4], glaucoma, Graves’ ophthalmopathy, and in some ocular tumors [5, 6].
Given that tears are readily available and can be collected via non-invasive procedures, they have recently been recognized as more valuable tools in the identification of ocular health and in the diagnosis of diseased states when compared to other biological fluids. Therefore, studies are being dedicated towards characterizing the composition of tears and understanding how changes in this composition relate to pathology. In several ocular disorders, tears have been proposed to promote inflammation, and the precise secretome of the tears directly relates to the ocular microenvironment [6]. However, characterizing how tears affect ocular health and more specifically how tears affect corneal epithelial cells remains poorly understood. Herein, we describe procedures that can be easily implemented to study how healthy vs. diseased tears affect the corneal epithelial cells in vitro. This protocol is based on our recently published paper, which focuses on how the tear proteins of Persistent Epithelial Defect (PED) patients have the potential to alter the inflammatory, fibrotic, and corneal epithelial barrier profile of other healthy cornea cells [7]. Taken together, we can infer that patient-derived tears can be a valuable source of factors to create a disease-specific in vitro model to study the key features of the pathology. This system can also be used to study the patient-specific clinicopathological features.
There are standard protocols describing the methods for collecting tears, extracting the protein, and using it for biochemical characterization, proteomic analysis, and inflammatory cytokines analysis. Recently, tears have been explored as a non-invasive method to identify disease-specific markers. Recent reports highlight the presence of exosomes in keratoconus tears, hypothesized as a communication vehicle [8]. The in vitro model used for studying disease pathology, such as in dry eye, allergy, or inflammatory models, is based on using a single growth factor/cytokine, which does not represent the actual pathology, as the disease state is governed by multiple factors such as cytokines, growth factors, and metabolites. The novel aspect of the described protocol is the use of patient-derived factors in the form of tears, which is the true representation of the diseased state (including growth factors, cytokines, and metabolites). Co-culturing patient-specific and control tears with relevant cells (corneal epithelial cells in this case) will provide an in vitro model that truly depicts the diseased state, whereas the healthy individual will be taken as a control. To the best of our knowledge, the ease of collecting tears and using it as a supplement in cell culture has never been explored previously, and this is the first study describing the detailed methodology for establishing a tear-based in vitro model of ocular diseases.
This protocol highlights the effect of patient-derived tears on corneal epithelial cells’ morphology and gene expression pattern. The procedure to collect tears can be done using either the Schirmer strip or the plastic capillary tube method. Sampling with Schirmer strip is relatively easy, rapid, reliable, and free of risk, whereas obtaining tears from capillary tube requires a degree of practice and experience; this procedure is also interrupted by blinking and the investigator needs to hold the tube for the duration of the sampling process. Hence, the protocol developed using the Schirmer’s test strip is easy to carry out, reproducible, and can be used to further understand ocular disease progression. Proteins are isolated post tear collection using the Schirmer’s test strip from both healthy and diseased individuals. Isolated proteins are then used to treat human corneal limbal epithelial (HCLE) cells at a specified concentration for a defined period (48 h in our study) until visual changes are observed. Next, morphological changes in inflammatory or fibrotic gene expression are recorded and studied. Matrix metalloproteinases (MMPs) are iron- and calcium-dependent enzymes that degrade the corneal collagen and the extracellular matrix. The level of MMP9 activity in the tear fluids has been regarded as an indicator of ocular inflammation and damage in dry eye disease, vernal keratoconjunctivitis, and glaucoma. Thus, MMP9 activity of the tears of PED patients has been studied via gelatin zymography and is represented in Figure 1. This novel assay uses tear fluid as a medium for testing and analyzing biomarkers, allowing for a more cost-effective and non-invasive diagnosis. The technique is also beneficial for tracking disease progression, as it can be used to detect changes in biomarkers over time. Furthermore, this method presents a convenient and time-saving method for healthcare professionals to monitor the health of their patients. Tears can be used for in vitro drug testing as a more effective and efficient means to test for the efficacy and safety of drugs. This model also mimics the in vivo conditions and is thus useful in drug testing.
Figure 1. Graphical representation of human corneal limbal epithelial (HCLE) cells treated with tear protein extracts and their downstream processing
Materials and reagents
50 mL conical centrifuge tubes (Thermo Fisher Scientific, catalog number: 339652)
15 mL conical centrifuge tubes (Thermo Fisher Scientific, catalog number: 339650)
100 mm culture dish (tissue culture treated) (Corning, catalog number: 430167)
12-well plate (tissue culture treated) (Thermo Fisher Scientific, catalog number: 173095)
10 mL serological pipettes (Thermo Fisher Scientific, catalog number: 170356N)
1,000 μL micro tip (Thermo Fisher Scientific, catalog number: 90030210-P)
200 μL micro tip (Thermo Fisher Scientific, catalog number: 90030130-P)
10 μL micro tip (Thermo Fisher Scientific, catalog number: 90030000-P)
1.5 mL microcentrifuge tube (Tarsons, catalog number: 500010)
Human corneal limbal epithelial (HCLE) cells (a kind gift from Dr. Ilene Gipson, Harvard Medical School, Schepen Eye Research Institute of Mass Eye and Ear)
Keratinocyte-serum free media (KSFM), supplemented with bovine pituitary extract and epidermal growth factor (EGF) (Gibco, Thermo Fisher Scientific, catalog number: 10724-011-500ML)
Penicillin/streptomycin solution (100×) (Diagnovum, catalog number: D910-100ML)
TrypLETM Express w/o phenol red (Gibco, Thermo Fisher Scientific, catalog number: 12604-013-100ML)
Minimal essential medium (MEM) (Gibco, Thermo Fisher Scientific, catalog number: 32571-036-500ML)
Fetal bovine serum (FBS) (Gibco, Thermo Fisher Scientific, catalog number: 10270-106-500ML)
Dulbecco’s PBS w/o Ca2+, Mg2+ and phenol red (1×) (Diagnovum, catalog number: D402-500ML)
Trypan blue (Thermo Fisher Scientific, catalog number: T10282)
Ammonium persulphate (APS) (Invitrogen, Thermo Fisher Scientific, catalog number: HC2005)
SureCastTM TEMED (Invitrogen, Thermo Fisher Scientific, catalog number: HC2006)
SureCastTM 40 % (w/v) acrylamide (29:1 acrylamide:bis-acrylamide) (Invitrogen, Thermo Fisher Scientific, catalog number: HC2040)
2.5× Tris-SDS buffer (pH 8.8) (HIMEDIA, catalog number: MB039-500ML)
5× Tris-SDS buffer (pH 6.8) (HIMEDIA, catalog number: MB040-500ML)
Gelatin, Hi-LRTM (HIMEDIA, catalog number: GRM019-500G)
0.2 μm Polyethersulfone (PES) syringe filter (mdi Membrane Technologies, catalog number: SYKG0601MNXX204)
Mini-RNA Isolation kit (Qiagen, catalog number: 74004)
Verso c-DNA Synthesis kit (Thermo Fisher Scientific, catalog number: AB1453)
SYBR Green (Applied Biosystems, Thermo Fisher Scientific, catalog number: A25742)
NuPage LDS sample buffer (4×) (Invitrogen, Thermo Fisher Scientific, catalog number: 2463558)
Calcium chloride (CaCl2) (HIMEDIA, catalog number: GRM3906-500G)
Triton X-100 (HIMEDIA, catalog number: MB031-100M)
Coomassie brilliant blue R-250 (HIMEDIA, catalog number: MB153-25G)
Methanol (Rankem, catalog number: M0140)
Acetic acid (Rankem, catalog number: A0060)
Sodium chloride (HIMEDIA, catalog number: MB023-500G)
1 mL syringes (DISPOVAN, model: U-40 Insulin syringe)
Schirmer tear test strips/tear touch (Madhu Instruments Pvt. Ltd.)
Gloves (Kimberly-Clark, catalog number: KM50601)
Kimwipes (Kimtech Science, catalog number: 34120)
Complete KSFM+GF (500 mL) (see Recipes)
Basal media: incomplete KSFM-GF (see Recipes)
1% Triton X-100 solution (see Recipes)
Developing buffer (see Recipes)
Coomassie staining solution (see Recipes)
Destaining solution (see Recipes)
Recipes
Complete KSFM+GF (500 mL)
Bovine pituitary extract, 25 μg/mL (half vial provided)
EGF (0.2 ng/mL)
CaCl2 (0.4 mM)
Penicillin/streptomycin 100× antibiotic (5 mL to achieve a final concentration of 1×)
Basal media: incomplete KSFM-GF
Penicillin/streptomycin 100× antibiotic (5 mL to achieve a final concentration of 1×)
1% Triton X-100 solution
1% Triton X-100 (v/v)
Autoclaved Milli-q water
Developing buffer
10 mM Tris buffer
200 mM NaCl
6 mM CaCl2
pH 7.4
Coomassie staining solution
Coomassie brilliant blue R-250 (0.05%)
Methanol (50% v/v)
Acetic acid (10% v/v)
Make up the remaining volume with Milli-Q water
Destaining solution
Acetic acid (10% v/v)
Methanol (50% v/v)
Milli-Q water (40% v/v)
Equipment
Swinging bucket centrifuge (NEUATION, model: ifuge UC02; catalog number: UC- 2002- EU/UK/AUS)
Tabletop centrifuge (Thermo Fisher Scientific, model: Sorvall Legend Micro 21R; catalog number: 75002447)
Class II biological safety cabinet (ESCO, model: Class II BSC)
Cell culture CO2 incubator (ESCO, model: CCL-170B-8-UV)
EVOSTM XL Core imaging system (Thermo Fisher Scientific, model: AMEX1200)
Automated cell counter (Thermo Fisher Scientific, model: CountlessTM 3 AMQAX2000)
Hemacytometer (Thermo Fisher Scientific, model: CountlessTM Reusable Slide, catalog number: A25750)
NanoDropTM Lite spectrophotometer (Thermo Fisher Scientific, catalog number: ND-LITE)
Gel Doc (Azure Biosystems, model: c600)
Real-time PCR (Azure Biosystems, model: Ceilo 6)
Thermocycler (Bio-Rad, model: T1000)
Rocker shaker (Scientek Hub, model: SR-1)
SureCastTM glass plates for PAGE ((Invitrogen, Thermo Fisher Scientific, catalog number: HC1001)
Mini gel tank (Invitrogen, Thermo Fisher Scientific, AMEX 1200; catalog number: A25977)
-80°C laboratory freezer (Thermo Fisher Scientific, model: Forma 900)
Milli-Q (Elga, model: PQ00018907)
Vortex (NEUATION, model: iSWIX)
Pipettes (1,000, 200, 10, 2.5 μL) (Eppendorf, model: Research Plus)
Software and datasets
Prism 9 (GraphPad Prism version 9.4.1)
BioRender for graphical images (https://www.biorender.com/)
Azure Cielo Manager software (version 1.0.4.0)
Procedure
Culturing and seeding of HCLE cells
Remove a vial of cryopreserved HCLE cells from -80 °C.
Place the vial at 37 °C for 3–4 min to thaw the cells.
Add 1 mL of the cryopreserved cells to 5 mL of MEM+10% FBS media in a 15 mL centrifuge tube in a BSL2 safety cabinet.
Centrifuge at 160× g for 5 min at room temperature.
Discard the supernatant.
Resuspend the pellet in 1 mL of KSFM+GF media.
Add the resuspended cells to a 100 mm tissue culture plate and add 7 mL of fresh KSFM+GF media.
Incubate the cells at 37 °C with 5% CO2 in the incubator to grow.
Once the cells attain 70% confluency, passage the cells twice before seeding for experiment.
For passaging the cells, remove the spent media from the 100 mm tissue culture plate.
Wash the plate with 4 mL of 1× Dulbecco’s PBS.
Add 1 mL of TrypLE solution to the plate and incubate at 37 °C with 5% CO2 for 3–5 min. Collect the trypsinized cells in a 15 mL centrifuge tube.
Neutralize the effects of TrypLE by adding 5 mL of MEM+10% FBS to the 15 mL centrifuge tube containing the trypsinized cells.
Centrifuge at 160× g for 5 min at room temperature. Discard the supernatant.
Resuspend the pellet in 1 mL of KSFM+GF media and add 500 μL to two 100 mm tissue culture plates. Make up the volume to 8 mL by adding 7.5 mL of complete KSFM media to each plate with the help of 10 mL serological pipettes.
Collection of tears from PED patients
Note: Prior to sample collection, patient informed consent must be taken.
A Schirmer strip test, as routinely used to evaluate the degree and severity of dryness in various disease conditions, is used to collect tears. Kindly note that a minimum tear migration up to 5 mm is necessary for optimal protein isolation.
Wear gloves and fold the end of the Schirmer strip within the packet at the ungraduated end.
Remove the Schirmer strip from the opposite end, place the folded end inside the lower lid of both eyes, and ask the patient to close their eyes.
After 5 min, the Schirmer strips are removed from both eyes.
The tear migration distance is noted from the graduated markings.
Collect the strips and place the entire strip containing the adsorbed tears in a 1.5 mL microcentrifuge tube (MCT) labeled with complete patient information.
Note: The important parameters to note during tear collection include the affected eye, patient medication, and the distance of tear migration.
Post tear collection, immediately place the tubes on ice (Figure 2).
Next, collect a second round of tears by repeating steps B2–B7. For control samples, tears are collected from patients not suffering from any ocular diseases.
Finally, store the vials in the -20°C freezer; those can be used within a period of six months.
Figure 2. Schematic depicting the method of tear collection from the control and patients’ eyes
Extraction of proteins from tears of patients
Thaw tear vials on ice.
Add 200 μL of 1× PBS to each vial using a 200 μL pipette. It is very crucial that no protease inhibitor cocktail is added while extracting proteins from tear samples, as it will interfere with protein activity.
With the help of a 1,000 μL pipette tip, grind the Schirmer strips in the MCTs.
Vortex for 30 s using a vortexer and then incubate on ice for 2 min.
Repeat step C4 3–4 times.
Centrifuge the MCTs at 16,000× g for 5 min at 4 °C.
Collect the extracted proteins present in the supernatant in a fresh 1.5 mL MCT using a 200 μL pipette.
Quantify proteins by measuring absorbance at 280 nm using NanoDrop (Figure 3).
For protein concentration measurement by NanoDrop, first set blank with 1 μL of 1× PBS using a 2.5 μL pipette. Clean the pedestal with Kimwipes.
Next, add 1 μL of the sample to be quantified as in step C9 and measure absorbance at 280 nm. Note the reading.
Use a 2 mL syringe to attach the 0.22 μm PES syringe filter to filter the extracted proteins.
Equilibrate the filter by adding 500 μL of KSFM-GF media to the 2 mL syringe using a 1,000 μL pipette. Filter the media and then discard the filtered media.
Next, filter the tear proteins before treating cells using a 0.22 μm PES syringe filter.
Add 200 μL of tear proteins to the same syringe filter inside a class II biological safety cabinet and collect the filtered tear proteins into a sterile 15 mL conical centrifuge tube.
Note: Please note that equilibrating the syringe filter with media is necessary to prevent any loss of tear proteins during the filtering process.
Figure 3. Schematic representation of protein extraction from Schirmer’s test strips
Seeding of HCLE cells for the experiment
Use the previously prepared 100 mm tissue culture dishes of HCLE cells at 70% confluency for seeding the experiment.
Discard the spent KSFM media from the culture dish using a 10 mL serological pipette in a BSL2 safety cabinet.
Wash the plate once with 3–4 mL of 1× Dulbecco’s PBS using a 1,000 μL pipette.
Add 1.5 mL of TrypLE solution to the plate after removing 1× PBS from the plate using a 1,000 μL pipette and incubate the plate at 37 °C for 5 min in a CO2 incubator.
Take out the plate from the CO2 incubator post incubation and tap the plate on both sides to dislodge the trypsinized cells.
In a Class II biological safety cabinet, collect the trypsinized cells in a 15 mL centrifuge tube containing 5 mL of MEM+10% FBS solution to neutralize the effects of TrypLE. Centrifuge the cells at 160× g in a swinging bucket centrifuge for 5 min at room temperature.
Discard the supernatant. Resuspend the pellet in 1 mL of KSFM+GF media using a 1,000 μL pipette.
Mix 10 μL of cells with 10 μL of Trypan Blue solution using a 10 μL pipette. Add 10 μL of this mix to one chamber of a hemacytometer.
Switch on the automated cell counter to count the cells.
Insert the hemacytometer into the automated cell counter. Adjust the brightness to focus the cells. Then, measure the cells’ density as indicated by the percentage of live cells calculated by the automated cell counter. Seed 50,000 cells/well in a 12-well tissue culture plate in 1 mL of KSFM+GF media for 24 h in a CO2 incubator at 37 °C.
Treating HCLE cells with proteins isolated from tears
Once the density reaches 60%–70% confluency, approximately 20 h after seeding, treat cells with tear proteins.
Remove the complete media from each well of the 12-well seeded plate using a 1,000 μL pipette.
Wash the cells once with 500 μL of 1× PBS to remove the remaining growth factors.
Add 1 mL of incomplete KSFM-GF media to each well using a 1,000 μL pipette.
Add an equal concentration of control and test tear proteins to the respective wells in a drop-wise manner. The protein concentration used for treating cells in vitro lies in the range of 60–100 μg/mL to each well.
Incubate the tear protein–treated cells in a CO2 incubator at 37 °C for 48 h.
Note the morphological changes taking place by treating HCLE cells with tear proteins of diseased patients. (Figure 4)
Figure 4. Morphological changes observed after tear treatment to human corneal limbal epithelial (HCLE) cells using EVOS Core imaging system. A. Control HCLE cells appear to be cuboidal maintaining epithelial stratification. B. HCLE cells treated with protein extracts of diseased Persistent Epithelial Defect (PED) patients were found to be more rounded, with decreased cell number and increased number of cellular vacuoles, thereby disrupting the normal stratification process. Please look out for changes in the cellular appearance, cellular vesicles, and the total number of cells present at the end of the experiment.
Analysis of MMP activity using gelatin zymography
After 48 h, collect the culture supernatant of both the control and tear protein–treated cells in 1.5 mL Eppendorf tubes to check the activity of MMPs by gelatin zymography.
Estimate the protein concentration of the supernatant using NanoDrop, following steps C9–C10.
Note: Make a note that for estimating protein concentration of culture supernatants, culture media devoid of growth factors is used to set the blank (KSFM w/o GF, in this case)
Next, mix an equal protein concentration of healthy control tears and patients’ tear-treated culture supernatant with 4× gel loading dye. Mix using a 200 μL pipette.
Note: Please make sure to avoid heating the samples or adding any reducing agent like β-mercaptoethanol during sample preparation.
Prepare an 8% acrylamide gel using 40% acrylamide, TEMED, 2.5× Tris-SDS buffer (pH 8.8), 5× Tris-SDS buffer (pH 6.8), and APS incorporated with 1 mg/mL of gelatin prepared in water.
Load the protein sample made with NuPage sample buffer (1×) and the pre-stained protein ladder onto the gel.
Run the gel at 30 mA for 1.5 h using the mini gel tank apparatus.
When the dye has completely run off, stop running the gel.
Transfer the gel to a staining box and incubate in 30 mL of 1% Triton X -100 for 30 min in a rocker shaker.
Note: Please note that the gelatin zymogram should be incubated in 1% Triton X-100 in order to get rid of SDS and renature proteinases that might affect MMP activity.
Discard the 1% TritonX-100 solution. Rinse the gel with Milli-Q water three times to remove the remaining Triton X-100.
Add 30 mL of developing buffer to the gel. Incubate overnight at 37 °C.
On the next day, remove the developing buffer. Stain the gel with 15 mL of Coomassie blue stain for 10 min in a rocking shaker.
Destain the gel in destaining solution for 5–7 min.
Note: Care must be taken not to extensively destain the gel.
Remove the destaining solution and store the gel in autoclaved water. Visualize the gel using a gel doc system to check for clear bands that indicate MMP activity (Figure 5 shows gelatin zymography depicting the levels of MMP9).
You can quantify the differences in the control and test samples using ImageJ. Refer to [7] for detailed information regarding the densitometric analysis of the MMP9 levels from gelatin zymography.
Figure 5. Gelatin zymography depicting the levels of MMP9 activity in the culture supernatants of human corneal limbal epithelial (HCLE) cells treated with tear proteins of control and Persistent Epithelial Defect (PED) patients
Gene expression analysis of tear protein–treated HCLE cells
Lyse cells in RNA lysis buffer and isolate RNA from the cells using Qiagen Mini-RNA Isolation kit, following the manufacturer’s instructions.
Quantify the RNA concentration by measuring absorbance at 260 nm using a NanoDrop, following steps C9–C10. Use the elution buffer provided in the RNA Isolation kit to set a blank for quantifying RNA.
The RNA concentration required to prepare 800 ng of cDNA in 20 μL falls within the range of 90–105 ng/μL. To ensure consistency, both the control and test samples should be converted to equal concentration of RNA to c-DNA using the Verso cDNA Synthesis kit, following the instructions provided by the manufacturer. Perform real time PCR analysis of the samples in duplicates using SYBR Green on an Azure Biosystems qPCR system. Run the housekeeping genes β-actin and RPL10 to study the effect on pro-inflammatory cytokines (as TNF-α, IL-1β, IL-6, TGFβ1, and TGF-β2), cell–cell communication, corneal epithelial homeostasis genes (such as E-cadherin, beta-catenin, and desmoglein), transcription factors like KLF4 (cell cycle inhibitor, EMT regulator, maintenance of corneal epithelial homeostasis), and other genes like GPX4 and NCO4.
For real time PCR, dilute the c-DNA to a final ratio of 1:10. Prepare a final reaction of 15 μL using SYBR green using 5 μL of diluted c-DNA.
Run the samples in duplicate in a 96-well Azure qPCR plate.
The cycling settings are as follows:
Step 1: 95 °C, 3 min
Step 2: 95 °C, 15 s
Step 3: 60 °C, 20 s
Step 4: Go to Step 2, 40 times
Step 5: start 60 °C; end 95 °C
Conduct the analysis of qPCR data using Azure Cielo Manager software and Microsoft Excel.
Employ the geometric mean of the housekeeping genes RPL10 and beta-actin for data normalization.
Utilize the relative expression (∆∆Ct) of genes to present the results, comparing the disease group to the control group.
Generate graphs using Prism 9. Incorporate the normalized fold change and standard errors of the mean for the qPCR data in the graphical representations.
Analyze the graphs to interpret the relative expression levels between the disease and the control groups.
Present the results by comparing the normalized fold changes and standard errors of the means for each gene. (Refer to Figure 6 for the gene expression data performed by real-time PCR.)
Figure 6. Gene expression analysis of HCLE cells treated with tear proteins (N = 3). A. Transcript level changes of the proinflammatory cytokines were studied in the control and PED tear–treated groups. B. The gene expression changes of the corneal homeostasis markers and transcription factors were analyzed in the control and PED tear–treated groups. The error bars in the figures represent the standard error of the means. An unpaired t-test was conducted, and the asterisk (*) denotes statistical significance (p < 0.05) [7].
Validation of protocol
Dutta et al. (2023). Prolonged Inflammation and Infectious Changes in the Corneal Epithelium Are Associated with Persistent Epithelial Defect (PED). Pathogens (Figure 4, panel A and B).
Acknowledgments
A.T. thanks SERB-DST Start-up research grant and Eicher Group Foundation (EGF) for supporting the study.
Competing interests
There are no conflicts of interest or competing interests.
References
Azkargorta, M., Soria, J., Acera, A., Iloro, I. and Elortza, F. (2017). Human tear proteomics and peptidomics in ophthalmology: Toward the translation of proteomic biomarkers into clinical practice. J. Proteomics 150: 359–367. doi: 10.1016/j.jprot.2016.05.006
Raposo, A. C., Portela, R. D., Aldrovani, M., Barral, T. D., Cury, D. and Oriá, A. P. (2020). Comparative Analysis of Tear Composition in Humans, Domestic Mammals, Reptiles, and Birds. Front. Vet. Sci. 7: e00283. doi: 10.3389/fvets.2020.00283
Torok, Z., Peto, T., Csosz, E., Tukacs, E., Molnar, A., Maros-Szabo, Z., Berta, A., Tozser, J., Hajdu, A., Nagy, V., et al. (2013). Tear fluid proteomics multimarkers for diabetic retinopathy screening. BMC Ophthalmology 13(1): e1186/1471–2415–13–40. doi: 10.1186/1471-2415-13-40
Aluru, S. V., Agarwal, S., Srinivasan, B., Iyer, G. K., Rajappa, S. M., Tatu, U., Padmanabhan, P., Subramanian, N. and Narayanasamy, A. (2012). Lacrimal Proline Rich 4 (LPRR4) Protein in the Tear Fluid Is a Potential Biomarker of Dry Eye Syndrome. PLoS One 7(12): e51979. doi: 10.1371/journal.pone.0051979
Ambroziak, A.M., Szaflik, J., Szaflik, J.P., Ambroziak, M., Witkiewicz, J., Skopiñski, P. (2016). Immunomodulation on the ocular surface: A review. Cent. Eur. J. Immunol. 41 (2): 195–208. doi: 10.5114/ceji.2016.60995
Hagan, S., Martin, E. and Enríquez-de-Salamanca, A. (2016). Tear fluid biomarkers in ocular and systemic disease: potential use for predictive, preventive and personalised medicine. EPMA Journal 7(1): e1186/s13167–016–0065–3. doi: 10.1186/s13167-016-0065-3
Dutta, T., Sangwan, J., Mondal, M., Vohra, M., Nidhi, V., Gour, A., Kapur, N., Gupta, N., Bhowmick, T., Chandru, A., et al. (2023). Prolonged Inflammation and Infectious Changes in the Corneal Epithelium Are Associated with Persistent Epithelial Defect (PED). Pathogens 12(2): 261. doi: 10.3390/pathogens12020261
Hefley, B. S., Deighan, C., Vasini, B., Khan, A., Hjortdal, J., Riaz, K. M., Liu, Y. and Karamichos, D. (2022). Revealing the presence of tear extracellular vesicles in Keratoconus. Exp. Eye Res. 224: 109242. doi: 10.1016/j.exer.2022.109242
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Multiple Labeling of Compartmentalized Cortical Neurons in Microfluidic Chambers
GM Guillermo Moya-Alvarado
AA Alejandro Aguirre-Soto
FB Francisca C. Bronfman
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4911 Views: 1052
Reviewed by: Miao HeCarlos Wilson Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Feb 2023
Abstract
Neurons are complex cells with two distinct compartments: the somatodendritic and the axonal domains. Because of their polarized morphology, it is challenging to study the differential cellular and molecular mechanisms that occur in axons and impact the soma and dendrites using conventional in vitro culture systems. Compartmentalized cultures offer a solution by physically and chemically separating the axonal from the somatodendritic domain of neurons. The microfluidic chamber model presented in this work is valuable for studying these mechanisms in primary cortical cultures derived from rat and mouse. In addition, this chamber model is compatible with various microscopy methods, such as phase contrast, and fluorescence imaging of living and fixed cells.
Key features
• Preparation and attachment of PDMS microfluidic chambers to glass coverslips.
• Primary culture of cortical neurons and plating cortical neurons in microfluidic chamber.
• Confirmation of compartmentalization using the retrograde transport of the fluorescently labeled form of cholera toxin subunit B (f-Ctb).
• Immunofluorescence and multilabeling of compartmentalized cortical neurons.
• Retrograde transport of fluorescently labeled BDNF.
Keywords: Compartmentalized cultures Microfluidic chambers Cortical neurons Endocytosis BDNF Axons Long-distance signaling Cholera toxin subunit B
Background
The nervous system (NS) connectivity depends on the polarized morphology of neurons, as they receive multiple inputs across dendrites and integrate and then propagate responses through a single axon. This specialized morphology depends on proper neuronal intracellular transport achieved by microtubule-based molecular motors, such as dynein and kinesins. The classic view is that the flow of information in neuronal circuits is from dendrites to axons; however, there are local events in axons that are communicated to the somatodendritic domain. Local axonal processes include, among others, the local translation of proteins after injury or retrograde transport of signaling endosomes initiated by extracellular cues (Hirokawa et al., 2010; Panayotis et al., 2015; Yamashita and Kuruvilla, 2016; Guedes-Dias and Holzbaur, 2019; Pathak et al., 2021). Axonal translation has been reported for various physiological and pathophysiological processes in the peripheral NS (PNS) and in the central NS (CNS). In the PNS, axonal local translation of proteins is required for regenerative responses in cell bodies upon axonal injury (Koley et al., 2019). In the CNS, the translatome of visual circuit axons has been established (Shigeoka et al., 2016), and local protein synthesis in the presynaptic compartment has been described as a ubiquitous feature of the adult brain (Hafner et al., 2019).
The endocytic system of axons actively participates in neuronal homeostasis. In the PNS, signaling endosomes are essential for long-range signaling from the axon to the nucleus, impacting cell survival (Yamashita and Kuruvilla, 2016; Villarin et al., 2016). In the CNS, our recent findings suggest a role for signaling endosomes in the wiring of neuronal circuits (Moya-Alvarado et al., 2023). Indeed, we have demonstrated that signaling endosomes transmit retrograde signals from the axon to the nucleus in a dynein-dependent manner, increasing dendritic branching in cortical neurons. Additionally, in the hippocampus, signaling endosomes locally regulate neurotransmission in the presynaptic compartment (Andres-Alonso et al., 2019; Moya-Alvarado et al., 2022 and 2023; Lazo and Schiavo, 2023).
The cellular and molecular aspects that govern axonal homeostasis, such as those discussed above, and how they impact the somatodendritic domain are difficult to study in normal in vitro cultures because we are unable to control the microenvironment of the axonal compartment. There are different types of compartmentalized systems for the in vitro culturing of neurons, including microfabricated platforms (Campenot, 1977), scaffold-based systems (Zhang et al., 2022), and microfluidic chambers (Park et al., 2006). Microfabricated platforms, such as Campenot chambers, were first developed to study the role of trophic factors applied to long-projecting neurons of the PNS (MacInnis and Campenot, 2002). However, the utility of Campenot chambers has remained limited given the arduous manufacturing process, incompatibility with high-resolution optical imaging, and limited throughput (Jadhav et al., 2016). In this protocol, we describe the use of a microfluidic chamber fabricated using lithography in polydimethylsiloxane (PDMS) as a platform for culturing primary cortical neurons from rat or mouse embryos. The advantages of microfluidic systems include compatibility with different types of neurons, improved fluidic compartmentalization, and enhanced imaging capabilities (Southam et al., 2013; Stuardo et al., 2020; De Vitis et al., 2021). Here, we describe a protocol to prepare and assemble microfluidic chambers from epoxy molds and to use them as a platform for primary culture of cortical neurons for fluorescence microscopy analysis of retrogradely transported proteins.
Materials and reagents
Mice (Mus musculus) or rats (Rattus norvegicus) embryos (16–18 days old)
Hank’s Balanced Salt Solution plus calcium and magnesium (HBSS) (Gibco, catalog number: 14025134)
Modified Eagle’s medium (MEM), high glucose, pyruvate (Gibco, catalog number: 11995081)
Horse serum (Gibco, catalog number: 16050122)
Coverslips (Marienfeld, catalog number: 0111650)
Neurobasal medium (Gibco, catalog number: 21103049)
B27 (Gibco, catalog number: 17504044)
GlutaMax supplement (Gibco, catalog number: 35-050-061)
Penicillin/streptomycin (10,000 U/mL) (Gibco, catalog number: 15140-122)
Cytosine B-D-arabinofuranoside (AraC) (Sigma-Aldrich, catalog number: C1768)
Microfluidic chamber (Xona microfluidics, catalog number: RD450 or SND450) or epoxy microfluidics molds. The molds used to prepare the compartmentalized chambers were fabricated at the microfluidic core facility of Tel Aviv University and donated by Professor Eran Perlson from the Department of Physiology and Pharmacology, Sackler Faculty of Medicine, Tel Aviv University (Gluska et al., 2016)
Cholera toxin subunit B (recombinant), Alexa Fluor 555 (f-Ctb555) (Invitrogen, catalog number: C34776)
Cholera toxin subunit B (recombinant), Alexa Fluor 647 (f-Ctb647) (Invitrogen, catalog number: C34778)
Mouse anti-βIII tubulin (Sigma, catalog number: T8578)
Rabbit anti-Rab11a (Invitrogen, catalog number: 715300)
Donkey anti-mouse IgG, Alexa 647 (Invitrogen, catalog number: A31571)
Donkey anti-rabbit IgG, Alexa 488 (Invitrogen, catalog number: A21206)
Poly-D-lysine (PDL) (Corning, catalog number: 354210)
Natural mouse laminin (Gibco, catalog number: 23017015)
Antibiotic/antimycotic 100× (Gibco, catalog number: 15240062)
6-well plate (Corning, catalog number: 15240062)
Cell culture dish 35 mm (Corning, catalog number: 353001)
SylgardTM 184 silicone elastomer kit (Dow Corning, catalog number: 4019862)
Trypsin 2.5% 10× (Gibco, catalog number: 15090046)
Mowiol 4-88 (Millipore, catalog number 475904)
Detergent Micro90 cleaning solution (Cole-Parmer, catalog number: 18100-05)
Ethanol (TCL Group, catalog number: IN-0150)
Distilled water (Sanderson, catalog number: SAN0003)
Phosphate buffered saline (PBS) 10× (Winkler, catalog number: BM-1340)
Glucose (Merck, catalog number: 108337)
Boric acid (Sigma-Aldrich, catalog number: B6768)
Borax anhydrous (Sigma-Aldrich, catalog number: B0127)
Syringe driven filters 0.22 µm (Jet Biofil, catalog number: JET-022)
Triton X-100 (Merck, catalog number: K42092903 220)
Hoechst (Invitrogen, catalog number: R37605)
Paraformaldehyde (Sigma, catalog number: 158127)
Sucrose (Merck, catalog number: 1.07687.1000)
Biopsy punch (Miltex, catalog number: 33-55)
Single edge blade (Stanley, catalog number: 28-510)
Cell culture dish 150 mm × 25 mm (SPL Life Sciences, catalog number: 201505SPL)
Parafilm (Amcor, catalog number: PM-996)
15 mL conical tubes (SPL Life Sciences, catalog number: 50015)
50 mL conical tubes (SPL Life Sciences, catalog number: 50150)
Pasteur pipettes (SPL Life Sciences, catalog number: 91010)
HEPES (Sigma, catalog number: 1001889449)
Sodium bicarbonate (Merck, catalog number, K50321729 832)
Bovine serum albumin (BSA) (Jackson ImmunoResearch Laboratories, catalog number: 001-000-162)
TrkB-fc (B&D Systems, catalog number: 688TK)
Fish gelatin (Sigma, catalog number: G7765)
Saponin (Sigma, catalog number: S4521)
Human BDNF-biotin (Alomone labs, catalog number: B-250-B)
Transferrin from human serum, Alexa 555 conjugate (Invitrogen, catalog number: T35352)
Streptavidin DyLight 488 (Invitrogen, catalog number: 21832)
Solutions
Polydimethylsiloxane (PDMS) mixture (see Recipes)
Poly-D-lysine (PDL) with laminin (see Recipes)
Borate buffer pH 8.3 (see Recipes)
HBSS 1× (see Recipes)
HBSS-trypsin (see Recipes)
MEM-HS (100 mL) (see Recipes)
Neurobasal-B27 (see Recipes)
Neurobasal-B27 with AraC (see Recipes)
Neurobasal-Cholera toxin subunit B (see Recipes)
Paraformaldehyde with sucrose (PFA solution) (see Recipes)
Blocking solution (see Recipes)
Antibody solution (see Recipes)
Recipes
Polydimethylsiloxane (PDMS) mixture
Reagent
Final concentration
Quantity
PDMS base (part A)
n/a
40.5 g
PDMS curing (part B)
n/a
4.5 g
Total
45 g
Note: SylgardTM 184 silicone elastomer kit contains the two different PDMS viscous solutions.
Poly-D-lysine (PDL) with laminin
Reagent
Final concentration
Quantity
Poly-D-lysine (1 mg/mL) prepared in borate buffer pH 8.3
0.1 mg/mL
1 mL
Laminin
20 µg/mL
0.1 mL
H2O
n/a
8,900 mL
Total
10 mL
Borate buffer pH 8.3
Reagent
Final concentration
Quantity
Boric acid
3.1 mg/mL
155 mg
Borax anhydrous
4.75 mg/mL
237.5 mg
H2O
n/a
50 mL
Total
50 mL
Note: Mix boric acid and borax anhydrous in distilled water using a magnetic bar and mix the solution overnight with a magnetic stirrer. After, adjust the pH and filter through a syringe driven filter (0.2 µm).
HBSS 1×
Reagent
Final concentration
Quantity
HBSS 10×
1×
10 mL
HEPES pH 7.4 1 M
10 mM
1 mL
Sodium bicarbonate
7.5%
0.5 mL
Antibiotic/antimycotic 100×
1×
1 mL
Autoclaved H2O
n/a
87.5 mL
Total
100 mL
HBSS-trypsin
Reagent
Final concentration
Quantity
10× Trypsin
1×
1 mL
HBSS 1×
1×
9 mL
Total
10 mL
MEM-HS (100 mL)
Reagent
Final concentration
Quantity
Horse serum
10%
10 mL
Glucose
0.6%
3 mL
GlutaMax 100×
1×
1 mL
Antibiotic/antimycotic 100×
1×
1 mL
MEM 1×
1×
86 mL
Total
100 mL
Neurobasal-B27
Reagent
Final concentration
Quantity
B27
2%
2 mL
GlutaMax 100×
1×
1 mL
Penicillin/streptomycin 100×
1×
1 mL
Neurobasal
1×
96 mL
Total
100 mL
Neurobasal-B27 with AraC
Reagent
Final concentration
Quantity
AraC 1 mM
1 µm
10 µL
Neurobasal-B27 (Recipe 7)
n/a
990 µL
Total
1 mL
Neurobasal-Cholera toxin subunit B
Reagent
Final concentration
Quantity
Cholera toxin subunit B Alexa 1 mg/mL
1 μg/mL
1 µL
Neurobasal
n/a
999 µL
Total
1 mL
Paraformaldehyde with sucrose (PFA solution)
Warm up PBS to 65 °C and add paraformaldehyde and sucrose. Mix every 5 min to completely dissolve the paraformaldehyde and complete volume to 5 mL with PBS 1×.
Reagent
Final concentration
Quantity
PBS 1×
n/a
Approximately 5 mL
Paraformaldehyde
4%
0.2 g
Sucrose
4%
0.2 g
Total
5 mL
Blocking solution
Reagent
Final concentration
Quantity
BSA
3%
0.3 g
Fish gelatin
5%
50 µL
Saponin 10% in PBS
0.2%
20 µL
PBS 1×
n/a
930 µL
Total
1 mL
Antibody solution
Reagent
Final concentration
Quantity
BSA
3%
0.3 g
Fish gelatin
5%
50 µL
Saponin 10% in PBS
0.02%
2 µL
PBS 1×
n/a
948 µL
Total
1 mL
Laboratory supplies
Pipette p2 (Thermo Scientific Finnpipette® F2, catalog number: 4642010)
Pipette p10 (Thermo Scientific Finnpipette® F2, catalog number: 4642040)
Pipette p20 (Thermo Scientific Finnpipette® F2, catalog number: 4642060)
Pipette p200 (Thermo Scientific Finnpipette® F2, catalog number: 4642080)
Pipette p1000 (Thermo Scientific Finnpipette® F2, catalog number: 4642090)
Alcohol burner (Usbeck), butane/propane mix (Providus s.r.l. 75% butane, 25% propane)
Neubauer counting chamber (HGB Germany)
Glass vacuum desiccator 210 mm (ISOLAB GmbHha, see Figure 1A)
HYBAID Incubator Shake 'n' Stack (Thermo Scientific, catalog number: HBMOVCST220)
Mini dialysis devices, Slide-A-Lyzer (Thermo Fisher, catalog number: 69590)
Equipment
Forceps short and extra fine length 110 mm (Dumont, catalog number: 11251-20)
Fine scissors 11.5 cm, 4 1/2" (Rudolf, catalog number: RU 1631-11 M)
Stereomicroscope for dissection (Leica, model: S6D L2)
Inverted phase contrast microscopy to visualize cells (Brand Motic, model: AE31E)
Horizontal laminar flow hood (Labtech, catalog number: LCB-0122H)
Vertical laminar flow hood (Labconco, catalog number: 3620924)
Cell vertical incubator (Forma Scientific, model: 3111)
Centrifuge (Hermle Labortechnik GmbH, catalog number: Z 233 MK-2)
Light microscope (Motic, model: AE31E Trinocular)
Confocal microscope (Leica, model: TCS SP8)
Software and datasets
LAS X version 3.5.5.19976 software
ImageJ (version 1.54J)
Procedure
Preparation and attachment of PDMS microfluidic chambers to glass coverslips
Preparation of microfluidic chambers
Take the epoxy molds, which are in a 10 cm plastic Petri dish, and use pressurized air/N2 to blow any remaining dust off the epoxy molds (Gluska et al., 2016) (see Figure 1).
Figure 1. Preparing coverslips and microfluidic chambers for seeding cortical neurons. (A) Vacuum desiccator containing microfluidics molds. The molds are placed into a 10 cm plastic Petri dish as shown in B. (B) Mold before being put into the oven. (C) No. 1 glass coverslips (25 mm) placed in a piece of parafilm in a plastic dish inside the hood. (D) Each coverslip was covered with 500 µL of poly-D-lysine (PDL) and laminin mixture and incubated overnight in the cell incubator at 37 °C. (E) Coverslips are washed three times with 600 µL of autoclaved distilled water using a p1000 pipette. (F) Photograph shows how to remove the coverslip with fine forceps to dry out the water if liquid is below the cover. (G) The silicone device contains four chambers (indicated by the black arrows) that will be cut after punching the four holes. (H) Silicone device with four punch holes in the first chamber. (I) Picture shows how to place the microfluidic chambers and the coverslips in the hood for drying. (J) The microfluidic chamber is attached to the coverslips. (K) The microfluidic chambers are attached to the coverslip in a 6-well plate containing plating media. The arrow indicates a small piece of paper below the cover that was added so the cover will not stick to the plastic well. (L) Close view of the microgrooves of the microfluidic chamber with plating media before plating cells. The arrow indicates a microgroove. Scale bar, 400 µm.
Keep the mold plates closed until casting using a piece of parafilm around the plate.
Prepare the PDMS mixture at a 1:10 ratio (see Recipe 1).
Pour the PDMS mixture into the molds. Place the molds in a vacuum desiccator for 2–3 h until the PDMS becomes clear without air bubbles (Figure 1B).
Move molds into a 70 °C oven for at least 3 h, up to overnight.
Using gloves, carefully position a scalpel at the edge of the mold and the PDMS cast. Precisely incise along the border of the cast with the scalpel to facilitate the detachment of the PDMS from the epoxy mold. Subsequently, with the aid of a spatula, extract the PDMS cast from the mold. Then, place the PDMS cast on a flat surface to separate the chambers and punch out the holes for plating the cells.
Use 5 mm biopsy punchers to cut holes in the PDMS. Punch four holes to allow the flux of media from side-to-side of the channel (see Figure 1H and Figure 2).
Note: The microgrooves must face up during this step. This will prevent disrupting microgrooves with the punch.
Figure 2. Plating cortical neurons in a microfluidic chamber. (A) Scheme of the microfluidic chamber used in this protocol. Five-millimeter punch holes are made on each side of the compartments in the chamber to allow the flux of media. Punch holes 1 and 2 define the cell body compartment; punch holes 3 and 4 define the axonal compartment. (B) When plating cells, the pipette tip is placed near the channel's entrance (red circle) and pipetted up and down in hole 2. Green dots represent cells. The black arrow shows the movement of the cells when plating. (C) Schematic representation of neuron distribution inside the cell body compartment. Red circles indicate where the pipette tip must be placed to pipette up and down and distribute the cells.
If the mold has more than one chamber, each chamber is cut into single chambers using a single-edge blade.
If you do not have the molds, you can use the commercial microfluidic chambers.
Place the chambers in a 10 cm plastic Petri dish and cover and seal them with parafilm until needed.
Glass coverslips and chambers preparation prior to neuronal seeding (Figure 1)
Inside the laminar flow hood, add an 8 cm2 parafilm layer into a sterile 150 mm ×25 mm plastic culture dish (Figure 1C).
Place the 25 mm No. 1 glass coverslips on parafilm (Figure 1C).
Coat each coverslip with 500 µL of PDL and laminin mixture (see Recipe 2) and incubate overnight in the incubator (Figure 1D).
The next day, hand wash (using plastic gloves) the microfluidic chambers for 5 min with 500 mL of 2% Micro90 detergent in distilled water. You can use your fingers to remove any dust or dirt from the chamber.
Place the chambers in a beaker and add 400 mL of distilled water, mix gently with a plastic spoon to remove the detergent thoroughly, and wait 10 min; repeat this process six times.
Put the chambers in 100 mL of 100% ethanol for 10 min at room temperature.
Mounting the chamber on top of the cover
Remove the plastic dishes containing coverslips from the incubator and place them inside a laminar flow hood.
Remove the mixture of PDL and laminin with the p1000 pipette.
Wash the coverslips three times with 600 µL of autoclaved distilled water (using the p1000 pipette, Figure 1E).
Dry out the coverslips using a vacuum and wait until the cover is completely dry. Place the coverslips inside a cell culture dish with the substrate side facing up.
Note: If there is liquid below the cover, take the cover with fine forceps and dry out the water (Figure 1F).
Put the microfluidic chambers inside the hood on delicate paper wipers with the microgrooves looking up (Figure 1I).
Turn on the UV light from the hood and irradiate and sterilize the chambers and the coverslips for 5 min.
Attach the microfluidic chamber to the coverslips. Adjust the chamber to the edge of the coverslip and drop the chamber. Gently touch the chamber to seal the chamber to the coverslip (Figure 1J).
Note: PDMS microfluidics exhibit a remarkable capacity to achieve a robust and reproducible seal with glass covers without requiring plasma-bonding procedures. This approach facilitates the removal and reusability of the microfluidic chamber for future experiments.
Place the microfluidic chamber attached to the glass coverslips inside a 35 mm dish or 6-well plate (Figure 1K).
Note: Below the glass coverslip, add a small piece of tissue paper. This will prevent the glass from adhering to the plastic (Figure 1K).
Add 400 µL of MEM-HS medium (see Recipe 6) to the chamber and put it inside the incubator until use (Figure 1K).
Notes:
i. At this point, you can check if the chambers are leaking by adding 200 µL of MEM-HS to the cell body compartment and wait for 2–3 min. Subsequently, observe the axonal compartment (Figure 1K) to detect any potential leaks.
ii. Since there is no hydrophobic bonding between the cover and the chamber, it is recommended to add MEM-HS (200 µL) directly to the channels of the cell body compartment (and not in the punch hole space) (see Figure 2A). If bubbles are left in the cell body compartment channel, you should pipette out the media with a p200 and refill until no bubbles are left.
Primary culture of cortical neurons and plating neurons in microfluidic chambers
Primary cultures of cortical neurons modified from Kaech and Banker (2006) for cortical neurons (Moya-Alvarado et al., 2023)
Add each embryo in 10 mL of HBSS (see Recipe 4) inside a 10 cm plate on a bucket with ice.
Decapitate the embryos and place the heads on a 10 cm plate with 10 mL of HBSS on ice.
Hold the heads through the eyes and open the skin and skull with scissors to expose the dorsal side of the brain.
Remove the brain from the skull and place on a 10 cm plate with 10 mL of HBSS on ice.
Under the dissection scope, separate the brain hemispheres and remove the meninges.
Dissect the cortex, cut it into small pieces (0.5–1 mm2), and add them to a 60 mm cell culture dish containing 3 mL of HBSS 1× on ice.
Transfer the small pieces of cortex with a Pasteur pipette to a 15 mL conical tube with 10 mL of HBSS-trypsin solution (see Recipe 5) to digest the tissue.
Incubate the small pieces of cortex for 10 min at 37 °C in the incubator or water bath.
Prepare three Pasteur pipettes with tips of three different diameters (1, 0.75, and 0.35 mm approximately). This is achieved by placing the fine part of the pipette close to the flame of a small gas burner.
Wash the small pieces of cortex placed in the 15 mL conical tube three times with 5 mL of cold HBSS 1× and wait until the tissue decants in the tube.
Remove the HBSS 1× and resuspend the cortex in 2 mL of MEM-HS with a p1000 pipette.
Then, dissociate the small pieces of cortex until the solution becomes cloudy by pipetting repeatedly with glass pipettes. First, use a pipette with a larger diameter tip, and then use a pipette with a smaller diameter tip.
Note: Pipette up and down the MEM-HS with each glass pipette before using them with the tissue. This mitigates the adhesion of tissue fragments to the glass surfaces, ensuring optimal handling of the sample.
Plating neurons in the microfluidic chambers
Count the cells in a Neubauer counting chamber.
Dilute or concentrate the cells in MEM-HS to the desired volume, depending on the number of chambers, by using centrifugation. Five microliters of MEM-HS containing 40 × 103–50 × 103 cells was used per chamber.
Note: If you plate 10 chambers, you should take 50 × 104 cells (50 × 103 per chamber) and resuspend them in 50 µL of MEM-HS.
Take the microfluidic chambers from the incubator and remove the MEM-HS from all four-punch holes (there will still be media in the channel compartment).
Add 5 µL (for a right-handed person use hole 2) of MEM-HS containing 50 × 103 cells into the chamber and pipette up and down from both sides of the cell body compartment (three times on each side) (Figure 2B and C).
Note: To obtain better results, use a p10 tip to plate the cells inside the chamber. This approach ensures precise and consistent cell distribution, contributing to subsequent reproducible experimental results. After adding the cells, pipette up and down in both sides of the cell body compartment to allow a homogeneous distribution of cells inside the chamber.
Allow the neurons to attach to the glass coverslips for 30 min inside the incubator.
Then, add 60 µL of MEM-HS in each punch hole (240 µL total in the four of them, as shown in Figure 2A) to fill the chamber.
On the next day, remove the MEM-HS from the chamber and add 80 µL of Neurobasal-B27 with AraC (see Recipe 8) in each punch hole of the cell body compartment and 40 µL of Neurobasal-B27 with AraC each punch hole of the axonal compartment (240 µL total) (see Figure 2).
Note: To facilitate the growth of axons toward the distal axon compartment, add a total of 160 µL of Neurobasal-B27 to the cell body compartment (80 µL in hole 1 and 80 µL in hole 2, as shown in Figure 2); 80 µL in total should be added to the distal axonal compartment (40 µL in hole 3 and 40 µL in hole 4, as shown in Figure 2). To minimize cell disruption, carefully add the media directly to the wall of the punched-out holes of the microfluidic chambers. Generally, consider adding 2/3 of the total volume to the cell body compartment and 1/3 to the distal axonal compartment. The differential volume between both compartments must be at least 80–100 µL to ensure microfluidic isolation of the two compartments. This differential volume must be conserved along all the cultures if fluidic isolation is required; therefore, media supplementation must be performed every two days due to media evaporation.
Change 20% of Neurobasal-B27 media every two days for a better growth of neurons at 5 days in vitro (DIV). Several axons are already grown in the axonal compartment (Figure 3).
Figure 3. Growth of cortical neurons in microfluidic chambers. (A) Representative images of cortical neurons grown in microfluidic chambers during the days in vitro (DIV). Cortical neurons are plated in the cell body compartment. During the days in vitro, neurons extend their axons along the microgrooves. Neurons reach the axonal compartment approximately at DIV 4–5. (B) Representative images of axons during different DIV (1–5). Scale bar, 50 µm.
Retrograde transport of fluorescently labeled f-Ctb in compartmentalized cortical neurons
Confirmation of microfluidic chamber compartmentalization and retrograde labeling of neuronal somas
Ctb is a protein that is endocytosed in neurons by binding the ganglioside GM1 in the plasma membrane. Axonal endocytosed Ctb is efficiently retrogradely transported in neurons and accumulates in the Golgi apparatus in the cell bodies of neuronal cells in vitro and in vivo (Wang et al., 2016).
To study the accumulation of Ctb in neuronal cell bodies, treat 5–7 DIV compartmentalized cultures with f-Ctb in the axonal compartment overnight. At the desired DIV, remove all the media from the chamber and add 80 µL of warm Neurobasal-B27 in hole 1 and 2 (Figure 2) and 40 µL of warm Neurobasal-B27 with f-Ctb674 (1 µg/mL) (see Recipe 9) in hole 3 and 4 in the axonal compartment (Figure 2). Subsequently, transfer the cells to the incubator overnight. This procedure allows us to recognize chambers that are well compartmentalized (not all cell bodies should be labeled with f-Ctb) and to label neurons with axons in the axonal compartment (Figure 4). Therefore, if a treatment is added to the axonal compartment, e.g., BDNF, only neurons accumulating f-Ctb are considered to quantify responses. For instance, neurons presenting activation of the transcription factor cAMP response element binding protein (CREB) are responsive to BDNF, as was done in the study by Moya-Alvarado et al. (2023).
On the next day, wash the f-Ctb647 with neurobasal medium and add 80 µL of f-Ctb555 (1 µg/mL) to the axonal compartment for 30, 90, 120, and 180 min. These treatments are useful for visualizing the co-internalization of any fluorescently labeled protein with f-Ctb (Figure 4 and Figure 5).
Figure 4. Retrograde transport of fluorescently labeled Alexa Fluor 647 and 555 cholera toxin subunit B (f-Ctb) in microfluidic chambers. Five days in vitro (DIV) cortical neurons were treated with f-Ctb647 (red) overnight. This treatment allows to verify chamber compartmentalization and label neurons that projected their axons to the axonal compartment. At 6 DIV, neurons were treated with f-Ctb555 for 30, 90, 120, and 180 min (green). Hoechst staining is shown in blue. This treatment allows to study the kinetics of f-Ctb transport and is useful for co-transport studies, as shown in Figure 5. (A) Upper panels show the cell body compartment. Scale bar, 50 µm. Lower panels show a magnified view of a group of cells shown in the square in the upper panel. Scale bar, 25 µm. White arrowheads show co-localization of f-Ctb555 and f-Ctb647. (B) Upper panels show axons in microgrooves labeled with f-Ctb555 (green). Lower panels show axons in microgrooves labeled with f-Ctb647 and f-Ctb555. Scale bar, 50 µm.
Figure 5. Axonal BDNF-positive endosomes co-localized with Ctb and partially with transferrin in the neuronal soma in compartmentalized cultures of cortical neurons. (A) Experimental design used to study transferrin co-localization with BDNF and Ctb retrogradely transported into neuronal cell bodies. In the cell body (CB) compartment, TrkB-Fc (100 ng/mL) was added for 60 min; then, in the axonal compartment, f-BDNF488 was added together with f-Ctb647 (1 µg/mL) for 3 h, while in the cell body compartment (CB) f-Transferrin 555 (100 µg/mL) was added to label recycling endosomes 30 min before withdrawing the treatment with f-BDNF488. (B) Images show co-localization of BDNF (green) and f-Ctb647 (cyan). The right panel shows a plot profile with the fluorescence intensity for BDNF (green) and f-Ctb647 (cyan) of the line shown in the figure. (C) Images show partial co-localization of BDNF (green) and transferrin (red). The right panel shows a plot profile with the fluorescence intensity for BDNF (green) and transferrin (red) of the line shown in the figure. (D) Images show partial co-localization of f-Ctb647 (cyan) and f-transferrin 555 (red). The right panel shows a plot profile with the fluorescence intensity for f-Ctb647 (cyan) and f-transferrin 555 (red) of the line shown in the figure. (E) Images show partial co-localization of BDNF (green), f-Ctb647 (cyan), and f-transferrin 555 (red). In the lower right panel, a plot profile with the fluorescence intensity for f-BDNF488 (green), f-Ctb647 (cyan), and f-transferrin 555 (red) from the line shown in the figure is shown. These are representative images of an experiment and were acquired using the UNAB Leica SP8 microscope at 63× magnification with a 5× digital zoom. Scale bar, 10 µm.
After the incubation, remove the media from the microfluidic chambers with a p200 pipette.
Wash the chambers with 200 µL of PBS one time on each side of the chamber (cell body and axonal compartment).
Carefully remove the chamber from the side of the coverslip to re-use the microfluidic chamber for another experiment before fixing the neurons.
Note: This procedure can be pursued either within the plate where the cells were initially cultured or by transferring the glass coverslips, bearing the neurons facing upward, over a piece of parafilm placed inside a container with a wet paper towel to maintain humidity (humid chamber). Additionally, it is possible to fix the neurons while the microfluidic chamber is still attached, but this is not recommended if the intention is to re-use the chamber afterward.
To fix the cells, add 100 µL of PFA solution (see Recipe 10) to the neurons and incubate them for 18 min at room temperature.
Note: To avoid detachment of neurons from the coverslips, add the solution slowly through the edge of the glass coverslip.
Remove the PFA solution and carefully wash three times with 100 µL of PBS for 5 min each wash.
Incubate the samples with 100 µL of Hoechst staining solution (1:5,000) for 10 min.
Wash the cells with 100 µL of PBS three times for 3 min each and then one time with 100 µL of distilled water to remove the salts from the PBS. Carefully drop the coverslip into 50 µL of mounting media (Mowiol 4-88) with the neurons facing the slide.
Dry the slides at room temperature for at least 12 h and then store the samples at 4 °C until imaging. Drying time may vary with different types of mounting media.
Note: To identify neurons that have projected their axons into the axonal compartment, the Ctb tracer must undergo an incubation period of a minimum of 3 h within the axonal compartment. This temporal requirement is crucial for optimal uptake and retrograde transport of Ctb, ensuring reliable neuronal labeling for subsequent analysis.
Immunofluorescence and multiple labeling of compartmentalized cortical neurons
Multiple labeling of f-Ctb, endocytic structures, and βIII-tubulin identifying microtubules in axons and cell bodies
At 5–7 DIV, remove the media from the chamber and add a total of 240 µL of warm Neurobasal-B27: 160 µL to the cell body compartment (80 µL in hole 1 and 2) and 80 µL with f-Ctb555 (1 µg/mL) in the axonal compartment (40 µL in hole 3 and 4). Incubate for at least 3 h in the incubator (Figure 2).
Remove the media from the four holes of the microfluidic chambers with a p200 pipette.
Perform three quick washes of the neurons with 50 µL of PBS (at room temperature) in each of the four punch holes of the chamber, as shown in Figure 2.
Carefully remove the chamber from the side of the coverslip to re-use the microfluidic chamber for another experiment.
Note: We re-used microfluidic chambers 2–3 times. Chambers are re-washed as indicated in Section A, step 2d and 2e.
Gently add 100 µL of PFA solution (at room temperature) to the neurons and incubate them for 18 min at room temperature.
Remove the PFA solution and carefully wash three times with 100 µL of PBS for 5 min each wash.
Remove PBS and add 150 µL of blocking solution (see Recipe 11) for 1 h at room temperature.
Note: For the staining described here, we used BSA and fish gelatin combined with saponin as a cell permeabilizer to block the samples. In our hands, this blocking buffer allows better visualization of endosomes in neuronal cells in vitro. If you wish to stain nuclear proteins, you will have to replace the saponin for Triton X-100 (see Moya-Alvarado et al., 2023). Additionally, BSA can be replaced by serum in the blocking solution that matches the species of the secondary antibody to decrease nonspecific binding.
Remove the blocking solution and add 100 µL of primary antibody solution (see Recipe 12). Then, incubate the cells with mouse anti-βIII tubulin (1:750) and rabbit anti-Rab11a (1:400).
Note: The concentration, condition, and time of incubation of the primary and secondary antibodies can vary depending on the antibody used during the experiment.
Incubate the neurons with primary antibody solution overnight at 4 °C.
Then, wash the primary antibody three times with 100 µL of PBS for 5 min each.
Remove the PBS and incubate the neurons with 100 µL of secondary antibody dissolved in antibody solution for 1 h at room temperature. For the staining illustrated in this protocol, we used donkey anti-mouse Alexa 647 (1:500) and donkey anti-rabbit Alexa 488 (1:500).
Wash the secondary antibody three times with 100 µL of PBS for 5 min each.
Incubate with 100 µL of Hoechst solution (1:5,000) for 10 min.
Wash the cells with 100 µL of PBS three times for 3 min each and then one time with 100 µL of distilled water to remove the salts from the PBS. Put the coverslip in a slide with mounting media (Mowiol 4-88).
Dry the slides at room temperature for at least 12 h and then store the samples at 4 °C until imaging. Representative images of this protocol are shown in Figure 6.
Figure 6. Multiple labeling of cortical neurons in microfluidic devices. (A) Scheme of the experimental design to label the neurons in microfluidic chambers. Five days in vitro (DIV) cortical neurons were treated with f-Ctb647 (red) overnight to stain all the neurons that projected their axons. At 6 DIV, neurons were fixed and immunostained. (B) Representative image of immunofluorescence of neurons in microfluidic chambers labeled with βIII-tubulin (cyan), Rab11a (green), and f-Ctb555 (red). Scale bar, 100 µm. (C) Representative image of neurons in the cell body compartment, microgrooves, and distal axons. Scale bar, 10 µm.
Labeling signaling endosomes with biotinylated BDNF
Conjugation of biotinylated BDNF (BDNF-biotin) with fluorescently labeled (DyLight488) streptavidin (f-streptavidin488)
We have previously published a protocol to monobiotinylate recombinant BDNF and to conjugate it to f-streptavidin (Stuardo et al., 2020). Here, we describe the protocol to conjugate commercially available BDNF-biotin as performed in Moya-Alvarado et al. (2023); both protocols were used in this publication.
Dissolve 5 µg of BDNF-biotin in 50 µL of neurobasal media in 0.1% BSA.
Note: Prepare 2 µL aliquots so you do not freeze and defrost the BDNF-biotin more than once (store aliquots at -80 °C). BDNF-biotin is not stable when it is stored more diluted. The concentration of BDNF will be 100 µg/mL. According to the manufacturer’s instructions, BDNF can have one or two biotin molecules.
Resuspend 10 µg of Streptavidin DyLight 488 in 100 µL of neurobasal medium with 0.1% BSA.
Note: If f-streptavidin contains sodium azide, it will have to be dialyzed with mini dialysis devices.
Add to an Eppendorf tube 2 µL of BDNF-biotin solution (100 μg/mL), 6 µL of f-strepavidin 488 (100 μg/mL), and 72 µL of neurobasal medium (supplemented with 0.1% BSA); mix up and down with a p200 pipette.
Incubate the solution for 20 min at 37 °C.
Then, dilute the mixture in 320 µL of neurobasal medium with 0.1% BSA at 37 °C to obtain a final solution of ~5 nM BDNF-biotin conjugated to f-streptavidin488 (or 150 ng/mL).
Note: As a control for conjugated BDNF treatment, nonconjugated f-streptavidin488 at the same final concentration is used.
Multilabeling neuronal endosomes with fluorescent BDNF (f-BDNF), f-Ctb647 from axons, and fluorescent labeled Alexa Fluor 555 transferrin (f-transferrin 555) from cell bodies
Compartmentalized cortical neurons are grown as indicated in Section B until DIV 8.
Remove the media from the chamber (80 µL from each hole of the cell body compartment and 40 µL from each hole of the axonal compartment).
Add 80 µL of TrkB-fc (100 µg/mL) in neurobasal medium to each hole of the cell body compartment and 40 µL of neurobasal medium to each hole of the axonal compartment for 60 min at 37 °C. This treatment neutralizes endogenous BDNF in the cell body compartment.
Then, remove the media from the axonal compartment and add f-Ctb647 (1 µg/mL) and f-BDNF488 (150 ng/mL) diluted in a total of 80 µL of neurobasal medium (1% BSA) for 180 min at 37 °C.
After 150 min of the abovementioned incubation, remove the media (80 µL from each hole) from the cell body compartment and then add 80 µL of neurobasal medium containing f-transferrin 555 (100 µg/mL) to each hole of the cell body compartment for the next 30 min to label early/recycling endosomes.
After 180 min, replace the medium in both compartments with warm PBS.
Then, proceed as indicated in Section D.
Confocal microscopy settings for Figures 5 and 6
We used a Leica SP8 confocal microscope with LAS X version 3.5.5.19976 software. Laser Diode 405, Diode 638, OPSL 488, and OPSL 552.
Identify the cells using the EPI fluorescence system by Hoechst or Ctb555 staining in the cell body compartment using the 63× objective.
Note: To visualize the cell body of neurons, we applied a 5× zoom. No zoom was applied for imaging axons in the axonal compartment or in the microgrooves.
Take 6–14 pictures using an optical slice 1 µm (or less) thick using the Z-scan option.
Note: In Figures 5 and 6, pictures of 1,024 × 1,024 pixel resolution were taken. In Figure 6, the following settings were applied: a gain of 850 and an offset O, with a 2%–5% intensity of 405 laser, 3%–10% 488 laser, 0.1%–0.5% 555 laser, and 0.1%–0.5% 647 laser. The setting conditions will vary depending on the experiment and must be set up for each sample.
Co-localization analysis using the fluorescence intensity plot profile as shown in Figure 5
Open ImageJ in the computer and load the image acquired by the confocal microscope.
Select from the Z-stack images the picture of the optical slice with higher fluorescence intensity from the channel labeling f-BDNF in Figure 5.
Select the option Straight line in ImageJ. Draw a line of 2–4 µm on top of the particle for analysis.
Add the line to the ROI (region of interest) manager to save the position of the line.
Open the option Analyze and select Plot profile for each channel.
Note: We also performed an analysis of the co-localization of f-BDNF endosomes with other labels, such as phospho TrkB antibodies, to detect signaling receptors using single organelle analysis as described in the methods of Moya-Alvarado et al. (2023).
Acknowledgments
The authors gratefully acknowledge financial support from ANID (Agencia Nacional de Investigacion y Desarrollo) FONDECYT grant N° 1221203 and DGI-UNAB (DI-01-21/NUC) from UNAB to FCB. This protocol is based on our previous work (Gluska et al., 2016).
Competing interests
The authors declare no conflicts of interest.
Ethical considerations
Pregnant animals were euthanized under deep anesthesia according to bioethical protocols approved by the Bioethics Committee of the Pontificia Universidad Catolica de Chile (protocol ID:180822013) and Universidad Andres Bello (act of approval 022/2019 and 009/2022).
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Correction Notice: HIV-CRISPR: A CRISPR/Cas9 Screening Method to Identify Genes Affecting HIV Replication
FR Ferdinand Roesch
MO Molly OhAinle
Published: Dec 5, 2023
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After official publication in Bio-protocol (https://bio-protocol.org/e3614), we noticed that the number of PCR cycles listed for the Second-Round PCR (PCR2) (Table 5) in H. Viral RNA HIV-CRISPR Library Amplification is incorrect. The table reads “x12” cycles but it should be “x20” cycles.
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4,913 | https://bio-protocol.org/en/bpdetail?id=4913&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Engineering a CRISPRoff Platform to Modulate Expression of Myeloid Cell Leukemia (MCL-1) in Committed Oligodendrocyte Neural Precursor Cells
MG Melanie Gil §
CH Catherine A. Hamann
JB Jonathan M. Brunger §
VG Vivian Gama §
(§ Technical contact)
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4913 Views: 842
Reviewed by: Salma MerchantVishal Nehru Anonymous reviewer(s)
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Abstract
In vitro differentiation of human pluripotent stem cell (hPSC) model systems has furthered our understanding of human development. Techniques used to elucidate gene function during early development have encountered technical challenges, especially when targeting embryonic lethal genes. The introduction of CRISPRoff by Nuñez and collaborators provides an opportunity to heritably silence genes during long-term differentiation. We modified CRISPRoff and sgRNA Sleeping Beauty transposon vectors that depend on tetracycline-controlled transcriptional activation to silence the expression of embryonic lethal genes at different stages of differentiation in a stable manner. We provide instructions on how to generate sgRNA transposon vectors that can be used in combination with our CRISPRoff transposon vector and a stable hPSC line. We validate the use of this tool by silencing MCL-1, an anti-apoptotic protein, which results in pre-implantation embryonic lethality in mice; this protein is necessary for oligodendrocyte and hematopoietic stem cell development and is required for the in vitro survival of hPSCs. In this protocol, we use an adapted version of the differentiation protocol published by Douvaras and Fossati (2015) to generate oligodendrocyte lineage cells from human embryonic stem cells (hESCs). After introduction of the CRISPRoff and sgRNAs transposon vectors in hESCs, we silence MCL-1 in committed oligodendrocyte neural precursor cells and describe methods to measure its expression. With the methods described here, users can design sgRNA transposon vectors targeting MCL-1 or other essential genes of interest to study human oligodendrocyte development or other differentiation protocols that use hPSC model systems.
Key features
• Generation of an inducible CRISPRoff Sleeping Beauty transposon system.
• Experiments performed in vitro for generation of inducible CRISPRoff pluripotent stem cell line amenable to oligodendrocyte differentiation.
• Strategy to downregulate an essential gene at different stages of oligodendrocyte development.
Graphical overview
Workflow for generating inducible CRISPRoff stem cell line and assessing knockdown phenotype in stem cell–derived committed oligodendrocyte neural precursor cells
Keywords: Oligodendrocytes Oligodendrocyte precursor cells CRISPR/Cas Sleeping Beauty MCL-1 Pluripotent stem cells
Background
In vitro systems of differentiation have become useful tools to dissect the molecular mechanisms involved in cell fate transitions. The use of human pluripotent stem cells (PSCs) has revolutionized our understanding of early stages of human development that were previously inaccessible. Different gene-targeting techniques, such as RNAi, TALEN, Cas9 nuclease, and CRISPRi/a, are available to investigate the function of specific genes during development [1–3]. Partial gene knockdowns in PSCs are commonly used to study the function of genes that would be lethal if completely knocked out. However, most of these tools have transient genetic modifications and their effectiveness is diminished during in vitro differentiation of PSCs. Nuñez et al. pioneered CRISPRoff, a programmable epigenetic memory writer protein that can heritably silence genes [4]. This tool has greatly benefited the field, allowing developmental researchers to further understand the function of genes during long-term in vitro differentiations, as epigenetic memory is stable. We built on the CRISPRoff system by adding a Sleeping Beauty transposon system containing a reverse tetracycline-controlled transactivator (Figure 1A). Additionally, we designed a single-guide RNA (sgRNA) transposon plasmid that allows for efficient genomic integration (Figure 1B). With the CRISPRoff and sgRNA transposon vectors described here, we were able to develop a novel experimental approach to inducibly silence expression of MCL-1, an anti-apoptotic protein, in oligodendrocyte lineage cells.
Deletion of Mcl-1 results in pre-implantation embryonic lethality [5]. Consequently, investigating the function of MCL-1 during development has barriers as mouse embryos are not able to implant and PSCs undergo apoptosis once MCL-1 is knocked down or inhibited via small molecules [6, 7]. This technical barrier has halted further research on the function of MCL-1 during early human neurodevelopment. With CRISPRoff, we silenced expression of MCL-1 at different stages of neurodevelopment. In this protocol, we outline how this set of tools can be used to investigate the potential function of MCL-1 during development of oligodendrocyte lineage cells. We adapted this technology and linked it to an established oligodendrocyte generation protocol [8] to validate the use of these novel tools during differentiation of human PSCs. Moreover, we detail how to generate sgRNA transposon vectors for other genes of interest that can be adapted into alternative differentiation systems.
There is a lack of literature that combines culturing of human oligodendrocyte lineage cells with epigenetic editing strategies. This protocol aims to bridge that gap by using a proof-of-principle experiment where MCL-1 is silenced at an early stage of differentiation. This protocol can be used to investigate an array of genes that cause lethality in PSCs or for studying gene function at different stages of oligodendrocyte development.
Materials and reagents
Biological materials
H9 human embryonic stem cells (WiCell Research Institute, WA09, NIH Registration Number: 0062)
Competent E. coli cells (New England Biolabs, catalog number: C2987H/C2987I)
pSB-TRE-CRISPRoff-EF1A-TetOn (Addgene, catalog number: 203355)
pSB-BbsI-sgRNA (Addgene, catalog number: 203359)
pCMV(CAT)T7-SB100 (Addgene, catalog number: 34879)
Reagents
BbsI-HF endonuclease (New England Biolabs, catalog number: R0539/R0539L)
rCutSmart buffer (New England Biolabs, catalog number: B6004S)
Quick calf intestinal alkaline phosphatase (CIP) (New England Biolabs, catalog number: M0525S/M0525L)
PCR and/or gel extraction purification kit (Invitrogen, catalog number: K220001)
T4 DNA ligase reaction buffer (New England Biolabs, catalog number: B0202S)
T4 DNA ligase (New England Biolabs, catalog number: M0202S/M0202L)
Polynucleotide kinase (PNK) (New England Biolabs, catalog number: M0201S/M0201L)
Recommended colony PCR reagents include Taq DNA Polymerase with ThermoPol® buffer (New England Biolabs, catalog number: M0267S/M0267L) and deoxynucleotide (dNTP) solution mix (New England Biolabs, catalog number: N0447S/N0447L)
DifcoTM LB broth, Miller (Luria-Bertani) (Becton Dickinson, catalog number: 244610)
LB agar, Miller (Fisher Scientific, catalog number: BP1425-500/BP1425-2)
Plasmid DNA Miniprep kit (Qiagen, catalog number: 27104)
Ampicillin sodium salt (Thermo Fisher Scientific, catalog number: 611770250/611770050)
TRIzolTM reagent (Thermo Fisher Scientific, catalog number: 15596026)
RNaseZAP (Thermo Fisher Scientific, catalog number: AM9780)
Chloroform (Millipore Sigma, catalog number: C2432)
2-Propanol (Millipore Sigma, catalog number: I9516)
Ethyl alcohol (Millipore Sigma, catalog number: 459836)
Diethyl pyrocarbonate (DEPC)-treated water (KD Medical, catalog number: RGE3050)
Ethylenediaminetetraacetic acid (EDTA), 0.5 M, pH 8.0, molecular biology grade, DEPC-treated (Millipore Sigma, catalog number: 324506)
DNase I (New England Biolabs, catalog number: M0303)
Thermo Fisher High-Capacity cDNA Reverse Transcription kit (Thermo Fisher Scientific, catalog number: 4368814)
SYBR Green PCR Master Mix (Thermo Fisher Scientific, catalog number: 4309155)
Nuclease-free water (Thermo Fisher Scientific, catalog number: AM9906)
Phosphate-buffered saline (PBS) tablets (Fisher BioReagents, catalog number: BP2944100)
Paraformaldehyde (PFA) 16% aqueous solution (Electron Microscope Sciences, catalog number: 15710-S)
Triton X-100 (Millipore Sigma, catalog number: T87876)
Bovine serum albumin (BSA) (Millipore Sigma, catalog number: A4503-100G)
Fluoromount-G (Thermo Fisher Scientific, catalog number: 00-4958-02)
Sodium dodecyl sulfate (SDS) (Millipore Sigma, catalog number: L3771)
Trizma® base (Tris Base) (Millipore Sigma, catalog number: T6066)
Glycine (Millipore Sigma, catalog number: G7126)
Sodium chloride (NaCl) (Millipore Sigma, catalog number: S7653)
Hydrochloric acid (HCl) (Millipore Sigma, catalog number: 258148)
Dry powder milk (RPI, catalog number: M17200)
TWEEN® 20 (Millipore Sigma, catalog number: P7949)
Phosphatase inhibitor PhosSTOP (Roche, Thermo Fisher Scientific, catalog number: 4906845001)
Protease inhibitor phenylmethylsulfonyl fluoride (PMSF) (Roche, Thermo Fisher Scientific, catalog number: 10837091001)
cOmpleteTM, EDTA-free protease inhibitor cocktail (PIC) (Roche, Thermo Fisher Scientific, catalog number: 4693132001)
Pierce bicinchoninic acid (BCA) Protein Assay kit (Thermo Fisher Scientific, catalog number: 23227)
4× BlotTM lithium dodecyl sulfate (LDS) sample buffer (Thermo Fisher Scientific, catalog number: B0007)
2-Mercaptoethanol (Bio-Rad, catalog number: 1610710)
Hoechst 33342 solution (Thermo Fisher Scientific, catalog number: 62249)
4%–20% Mini-PROTEAN® TGXTM Precast protein gels (Bio-Rad catalog number: 4561091)
mTeSR1 (STEMCELL Technologies, catalog number: 85850)
Matrigel Matrix hESC-qualified mouse (Corning, catalog number: 354277)
DMEM/F-12 (N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid) HEPES (Thermo Fisher Scientific, catalog number: 11330032)
Gentle cell dissociation reagent (STEMCELL Technologies, catalog number: 100-0485)
ROCK1 and ROCK2 inhibitor Y-27632 (STEMCELL Technologies, catalog number: 72307)
Opti-MEM I reduced serum medium (Thermo Fisher Scientific, catalog number: 31985070)
TransIT-LT1 transfection reagent (Mirus, catalog number: 2304)
Accutase (Innovative Cell Technologies, catalog number: AT104)
Puromycin dihydrochloride from Streptomyces alboniger (Millipore Sigma, catalog number: P8833-10MG)
Hygromycin B (Thermo Fisher Scientific, catalog number: 10687010)
Caspase inhibitor Quinoline-Val-Asp-Difluorophenoxymethylketone (Q-VD-Oph) (SM Biochemicals LLC, catalog number: SMP001-5MG)
Doxycycline hyclate (Tocris Bioscience, catalog number: 4090)
Insulin solution human (Thermo Fisher Scientific, catalog number: I9278)
DMEM/F-12 (Thermo Fisher Scientific, catalog number: 11320033)
Penicillin-Streptomycin (Thermo Fisher Scientific, catalog number: 15140122)
MEM non-essential amino acids solution 100× (Thermo Fisher Scientific, catalog number: 11140050)
GlutaMAX supplement (Thermo Fisher Scientific, catalog number: 35050061)
Retinoic acid (Millipore Sigma, catalog number: R2625)
TGF-beta/Smad inhibitor LDN-193189 (Reprocell, catalog number: 04-0074)
TGF-beta/Smad inhibitor Stemolecule SB431542 (Reprocell, catalog number: 04-0010-10)
N-2 supplement 100× (Thermo Fisher Scientific, catalog number: 17502048)
Smoothened agonist (SAG) (STEMCELL Technologies, catalog number: 73412)
Ultrapure DNase/RNase-free distilled water (Thermo Fisher Scientific, catalog number: 10977015)
Solutions
DNase I reaction (see Recipes)
Reverse transcription master mix (see Recipes)
1% Triton (see Recipes)
4% PFA (see Recipes)
10% BSA (see Recipes)
Lysis buffer (see Recipes)
10× Tris-Gly-SDS buffer (see Recipes)
20× TBS (see Recipes)
TBST (see Recipes)
5% Milk in TBST (see Recipes)
Basal medium (see Recipes)
Neural induction medium (NIM) (see Recipes)
N2 medium (see Recipes)
Recipes
DNase I reaction
Note: Keep on ice and use immediately after preparation.
Reagent Final concentration Quantity or Volume
DNase I reaction buffer (10×) 1× 1 µL
DNase I (RNase-free) n/a 0.2 µL
RNA sample n/a 2 µg
Nuclease-free H2O n/a Up to 10 µL
Total n/a 10 µL
Reverse transcription master mix
Note: Keep on ice and use immediately after preparation.
Reagent Final concentration Quantity or Volume
10× RT buffer 1× 2 µL
25× dNTP mix 1× 0.8 µL
10× RT random primers 1× 2 µL
MultiScribeTM reverse transcriptase n/a Up to 10 µL
Nuclease-free H2O n/a 4.2 µL
Total n/a 10 µL
1% Triton
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
Triton X-100 1% 50 µL
1× PBS n/a 49.5 mL
4% PFA
Reagent Final concentration Quantity or Volume
16% PFA 4% 12.5 mL
1× PBS n/a 37.5 mL
10% BSA
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
BSA 10% 5 g
1× PBS n/a 50 mL
Lysis buffer
Note: Keep on ice and use immediately after preparation.
Reagent Final concentration Quantity or Volume
1% Triton (Recipe 3) 0.79% 79 µL
10× PhosSTOP 1× 10 µL
10× PIC 1× 10 µL
100 mM PMSF 1 mM 1 µL
Total n/a 100 µL
10× Tris-Gly-SDS buffer
Reagent Final concentration Quantity or Volume
Tris Base n/a 121.1 g
Glycine n/a 576 g
SDS 1% 200 mL
ddH2O n/a Up to 4 L
Total n/a 4 L
20× TBS
Note: Adjust pH to 7.6 with HCl.
Reagent Final concentration Quantity or Volume
Tris Base n/a 193.6 g
NaCl n/a 640 g
ddH2O n/a Up to 4 L
Total n/a 4 L
TBST
Reagent Final concentration Quantity or Volume
20× TBS 1× 50 mL
ddH2O n/a 949 mL
TWEEN® 20 0.1% 1 mL
Total n/a 1 L
5% Milk in TBST
Reagent Final concentration Quantity or Volume
Dry powder milk 5% 5 g
TBST n/a 100 mL
Total n/a 100 mL
Basal medium
Note: Filter sterilize and store at 4 °C.
Reagent Final concentration Quantity or Volume
DMEM/F12 n/a 485 mL
MEM non-essential amino acids solution 1× 5 mL
GlutaMAX supplement 1× 5 mL
Penicillin-Streptomycin 1× 5 mL
2-Mercaptoethanol 55 µM 1.93 µL
Total n/a 500 mL
Neural induction medium (NIM)
Note: Store at 4 °C. *Add small molecules fresh daily.
Reagent Final concentration Quantity or Volume
Basal medium (Recipe 7) n/a 50 mL
Insulin 25 µg/mL 108 µL
SB431542* 10 µM 25 µL
LDN193189* 250 nM 6.25 µL
Retinoic acid* 100 nM 50 µL
Total n/a 50 mL
N2 medium
Note: Store at 4 °C. *Add small molecules fresh daily.
Reagent Final concentration Quantity or Volume
Basal medium (Recipe 7) n/a 49.5 mL
N-2 supplement 100× 1× 500 µL
SAG* 250 nM 5 µL
RA* 100 nM 50 µL
Total n/a 50 mL
Laboratory supplies
1.5 mL microcentrifuge tubes (Fisherbrand, catalog number: 05-408-1317)
PCR tubes (Thermo Fisher Scientific, catalog number: AB2000)
Costar 6-well clear TC-treated well plates (Corning, catalog number: 3516)
Cell lifter (Corning, catalog number: 3008)
Surface-treated sterile tissue culture plates (Fisherbrand, catalog number: FB012928)
Stericup-GP 500 mL Express Plus (Millipore Sigma, catalog number: S2GPU05RE)
35 mm glass-bottom dish with 14 mm micro-well #1.5 cover glass (Cellvis, catalog number: D35-14-1.5-N)
Equipment
Applied BiosystemsTM QuantStudioTM 3 Real-Time PCR System, 96-well, 0.2 mL, laptop (Thermo Fisher Scientific, catalog number: A28567)
Thermocycler (Thermo Fisher Scientific, catalog number: 4484073)
Incubated shaker (Thermo Fisher Scientific, catalog number: SHKE6000)
Synergy HT Microtiter Plate reader (BioTek, catalog number: 7091000)
Amersham Imager 600 (General Electric, catalog number: AI600)
Stirring Hotplates (Thermo Fisher Scientific, catalog number: SP88854100)
Mini-PROTEAN Tetra Vertical Electrophoresis Cell (Bio-Rad, catalog number: 1658004
PowerPacTM HC power supply (Bio-Rad, catalog number: 1645052)
Spinning disk confocal microscope (Nikon Eclipse Ti-E equipped with a Plan Apo Lambda 20× 0.75 NA WD 1.00 mm objective and an Andor DU-897 EMCCD camera)
Software and datasets
Prism v9 (GraphPad, 10/24/2020)
Image Studio Lite
SnapGene v6.2.1
Procedure
Cloning strategy
Sleeping Beauty transposon plasmids (Figure 1) were cloned using NEBuilder® HiFi DNA Assembly Mastermix and are available from Addgene as noted in section Biological materials above.
Figure 1. Sleeping Beauty transposon plasmids. (A) CRISPRoff transposon plasmid (Addgene #203355) encodes constitutive expression of the puromycin N-acetyl-transferase gene (PuroR) and a reverse tetracycline-controlled transactivator (Tet-on 3G) that enables doxycycline-dependent activation of the tetracycline response element (TRE)-driven transgene. The TRE regulates transcription of the CRISPRoff enzyme, composed of the DNA methyltransferases 3a (Dnmt3a) and 3-like (Dnmt3L, as well as dCas9-KRAB fused to blue fluorescent protein (BFP). SV40 PolyA: Simian virus 40 polyadenylation signal sequence. EF1α: Elongation factor-1 alpha. P2A: Porcine teschovirus-1 2A self-cleaving peptide. bGH PolyA: Bovine growth hormone polyadenylation signal sequence. (B) Single-guide RNA (sgRNA) transposon plasmid (Addgene #203359) encodes constitutive expression of the hygromycin phosphotransferase (HygroR) transgene. The U6 type III polymerase promoter constitutively drives expression of the sgRNA, which consists of a constant scaffold region and a user-defined spacer sequence. Inset: Annealed sense and anti-sense oligos should include the appropriate BbsI overhangs for successful ligation. Both transposon plasmids contain inverted terminal repeats (ITRs) spanning either end for successful genomic integration mediated by the Sleeping Beauty 100× transposase.
sgRNA design
Numerous resources are available to aid in the selection of target-specific sgRNA sequences. For example, Horlbeck et al. and Sanson et al. have established sgRNA design rules for suppressing transcripts in the human genome via CRISPRi [9, 10]. Horlbeck et al. generated a database of predicted, active sgRNAs against the human genome, and this database was used to design sgRNAs in the seminal CRISPRoff work, suggesting that CRISPRi libraries serve as a reasonable starting point for selecting sgRNAs for CRISPRoff. The CRISPRi library developed by Sanson et al. is available via the CRISPick web portal located at https://portals.broadinstitute.org/gppx/crispick/public. Users select the appropriate genome editing mechanism (i.e., CRISPRi), and Cas protein (i.e., SpyoCas9) to identify sgRNAs with predicted high and specific activity. Addgene also maintains a table of gRNA sequences available from the depository, which users can mine to determine whether genes of interest have been effectively targeted in CRISPRi or CRISPRoff experiments (https://www.addgene.org/crispr/grnas/). It is useful to note, however, that effective sgRNAs for CRISPRi are typically localized to a narrow window downstream of the transcription start site (TSS), while CRISPRoff-mediated gene repression displays a wide targeting window spanning a distance in excess of 2 kb from the TSS. Thus, candidate sgRNA sequences that are not predicted by tools referenced above may be empirically tested and validated. Critically, users will need to identify potential Cas9 target sites that contain a protospacer adjacent motif (PAM) for Spyo Cas9 (5′-NGG-3′). These sites can be identified using tools such as the UCSC Genome Browser (https://genome.ucsc.edu/). The sgRNA sequences used in this protocol can be found in Table S1.
sgRNA transposon plasmid design
Preparation of sgRNA backbone
Perform BbsI digest and backbone dephosphorylation by combining ~1 µg of sgRNA transposon plasmid (Addgene #203359), 1 µL of BbsI endonuclease, 2 µL of rCutSmart Buffer, 1 µL of Quick CIP, and water to bring to a total final volume of 20 µL in a sterile tube. Place at 37 °C for 1–3 h.
Perform PCR purification or gel extraction to purify the digested backbone. In our experience, a PCR purification is adequate as the excised fragment is 22 bp long.
Preparation of sgRNA insert
Design and obtain oligos containing sgRNA sequence(s) for integration at the BbsI sites in the sgRNA transposon plasmid (Addgene #203359). Sense and anti-sense oligos should contain the appropriate overhang specific to the BbsI cut site to allow for successful ligation (Figure 1). Reconstitute oligos to 100 µM using ultrapure water.
Place 4 µL of each of the 100 μM stock of sense and anti-sense oligos along with 4 µL of T4 DNA ligase buffer, 1 µL of PNK, and 27 µL of ultrapure water in a sterile tube and mix well.
Phosphorylate and anneal oligos using the following protocol, which can be programmed into a thermocycler:
37 °C for 30 min
95 °C for 10 min
85 °C for 1 min
75 °C for 1 min
65 °C for 1 min
55 °C for 1 min
45 °C for 1 min
35 °C for 1 min
25 °C for 1 min
After oligos have been phosphorylated and annealed, dilute 1:50 in ultrapure water and mix well.
Ligation and verification
Add 75 ng of purified backbone along with 1 µL of insert (diluted annealed/phosphorylated oligos), 2 µL of T4 DNA ligase buffer, 1 µL of T4 DNA ligase, and water to bring to a final volume of 20 µL, mixing well. Incubate at 16 °C for 1 h.
Transform the reaction mixture into chemically competent E. coli, such as DH5α strain, spread onto an agar plate supplemented with ampicillin or, alternatively, carbenicillin (100 mg/mL) and incubate overnight at 37 °C [11].
Perform a colony PCR using bacterial colonies. We use a primer that binds the U6 promoter (5′-ttcttgggtagtttgcagtttt-3′) as a forward primer and the anti-sense oligonucleotide as a reverse primer. Use the bacterial colony as template DNA, making sure to swirl the colony once in PCR mix before also swirling the same tip in a small volume (i.e., 100 µL) of LB broth for growth of positively identified colonies.
Positively identified clones should be grown in overnight cultures in 3–5 mL of LB supplemented with ampicillin (100 μg/mL) on a shaking incubator at 37 °C. After 16 h in culture, plasmid DNA should be extracted from E. coli using a Miniprep Plasmid Purification kit.
Sanger sequencing can be performed spanning BbsI cut sites to ensure successful assembly of the sgRNA transposon vector. Again, we typically use the U6 primer noted previously for Sanger sequencing. Alternatively, users may also elect to submit samples for full plasmid sequencing service.
Maintenance of human embryonic stem cells (hESCs)
H9 hESCs were maintained in mTeSR1 on Matrigel-coated plates. Change media daily and passage when needed.
Prepare Matrigel (1 mg/24 mL of DMEM/F12, HEPES).
Coat plates (1.5 mL for each well of a 6-well plate) overnight in 37 °C incubator.
(Optional) Wash cells with 1 mL of gentle cell dissociation buffer. Aspirate.
Incubate with 1 mL of gentle cell dissociation buffer for 3–5 min at room temperature (RT) (check on the microscope for efficient dissociation). Aspirate buffer.
Add fresh mTeSR1, scrape gently with cell lifter, and resuspend the cells with a serological pipette such that they are in small clumps.
Split cells 1:6 (if the well is > 80% confluent) onto 6-well plates for maintenance.
Incubate cells at 37 °C and 5% CO2.
CRISPRoff transfection
Figure 2 outlines the experimental procedures in sections E–I.
Figure 2. Generation of cell lines. Day 0: Transfect human embryonic stem cells (hESCs) with CRISPRoff transposon and Sleeping Beauty 100× transposase plasmids. Blue wells indicate no DNA negative control and pink wells indicate experimental wells with DNA. Day 4: Passage cells when they reach 70%–80% confluency. Day 5: Start antibiotic selection with puromycin. Blue wells with - indicate no DNA negative control without puromycin. Blue wells with + indicate no DNA negative control with puromycin. Pink wells with - indicate experimental wells with DNA and without puromycin. Pink wells with + indicate experimental wells with DNA and with puromycin. Day 20: Transfect hESCs with sgRNA pool and transpose plasmids. Day 24: Passage cells when they reach 70%–80% confluency. Day 25: Start antibiotic selection with hygromycin. Blue wells with - indicate no DNA negative control without hygromycin. Blue wells with + indicate no DNA negative control with hygromycin. Pink wells with - indicate experimental wells with DNA and without hygromycin. Pink wells with + indicate experimental wells with DNA and with hygromycin. Day 40: Validate stem cell properties. SB = Sleeping Beauty
Day 1
Place 250 µL of mTeSR1 containing 1 µL of 10 mM Y27632 into a well of a Matrigel-coated 12-well plate.
Prepare a mixture with 0.75 µg of the Sleeping Beauty transposase plasmid [pCMV(CAT)T7-SB100] and 1.25 µg of the CRISPRoff plasmid in 200 µL of Opti-MEM I reduced serum medium supplemented with 6 µL of TransIT-LT1. Include a mixture without DNA. This is the no-DNA negative control.
After a 15-min incubation at RT, add the transfection mix into a well of a 12-well plate.
Add 1 mL of Accutase per well (for cells in a 6-well plate) to detach hESCs. Incubate for 5 min at 37 °C.
Dilute the Accutase solution by adding 2 mL of DMEM/F12 medium.
Use a cell lifter to remove cells from the well and gently pipette the mixture 2–5 times with the p1000 pipette to fully dissociate the hESC cell colonies to single cells.
Transfer the cells to a conical tube.
Centrifuge the cells at 200× g for 4 min at RT.
Resuspend the cell pellet in 1 mL of mTeSR1 containing 10 µM Y27632 and count the cells with a hemocytometer.
Dilute cell suspension to 1.5 × 106 cells/mL.
Place 500 µL of this cell suspension to the well containing the transfection mix.
Incubate cells overnight at 37 °C with 5% CO2.
Day 2
Aspirate media and replace with fresh mTeSR1 without Y27632.
Day 4
Cells should be 70%–80% confluent at this day.
Passage cells into a new 12-well plate as previously outlined in section D.
Antibiotic selection
Ensure that the appropriate puromycin concentration needed has been previously validated from an antibiotic kill curve for the cell type of interest. For the experiments outlined here, a puromycin concentration of 0.8 mg/mL was used for selection.
The day after passaging, supplement mTeSR1 with the appropriate concentration of puromycin.
Continue feeding daily with mTeSR1 supplemented with puromycin for 14 days. It is expected for the negative control cells that did not receive DNA during transfection to die within the first few days. Similarly, untransfected cells in the experimental wells should all die after 14 days. Surviving cells are expected to have integrated the CRISPRoff system into their genome to generate a stable cell line.
From this point forward, media must be supplemented with a half dose of puromycin.
Continue to maintain cells to prepare them for the second transfection.
At this stage, cryopreservation is recommended, as this stable cell line can be used for future transfections with different sgRNA pools.
sgRNA transfection
Day 1
Place 250 µL of mTeSR1 containing 1 µL of 10 mM Y27632 into a well of a Matrigel-coated 12-well plate.
Prepare a mixture with 1 µg of the Sleeping Beauty transposase plasmid [pCMV(CAT)T7-SB100] and 1 µg of the sgRNA pool in 200 µL of Opti-MEM I reduced serum medium supplemented with 6 µL of TransIT-LT1.
Make sure to include a non-targeting (NT) sgRNA pool in addition to the sgRNA pool targeting the gene of interest.
Include a mixture without DNA (DNA negative control).
After a 15-min incubation at RT, add the transfection mix into a well of a 12-well plate.
Add 1 mL of Accutase per well (for cells in a 6-well plate) to detach hESCs. Incubate for 5 min at 37 °C.
Dilute the Accutase solution by adding 2 mL of DMEM/F12 medium.
Use a cell lifter to remove cells from well and gently pipette the mixture 2–5 times with the p1000 pipette to fully dissociate the hESC cell colonies to single cells.
Transfer the cells in a conical tube.
Centrifuge the cells at 200× g for 4 min at RT.
Resuspend the cell pellet in 1 mL of mTeSR1 containing 10 µM Y27632 and count the cells with a hemocytometer.
Dilute cell suspension to 1.5 × 106 cells/mL.
Place 500 µL of this cell suspension to the well containing the transfection mix.
Incubate cells overnight at 37 °C with 5% CO2.
Day 2
Aspirate media and replace with fresh mTeSR1 without Y27632.
Day 4
Cells should be 70%–80% confluent at this day.
Passage cells into a new 12-well plate as previously outlined in section D.
Selection
Ensure that the appropriate hygromycin B concentration needed is determined from an antibiotic kill curve for the specific cell type of interest. A hygromycin B concentration of 60 mg/mL was used for selection in our experiments.
The day after passaging, supplement mTeSR1 with the appropriate concentration of hygromycin.
Continue feeding daily with hygromycin supplementation for 14 days. It is expected for the negative control cells that did not receive DNA during transfection to die within the first few days. Similarly, un-transfected cells in the experimental wells should all die after 14 days. The sgRNA transposons are expected to have integrated the chromosomes of surviving cells, giving rise to a stable cell line.
From this point forward, media must be supplemented with a half dose of puromycin and hygromycin B.
Validating pluripotency and genome integrity (Figure 3A and 3B)
Figure 3. Validation of stem cell pluripotency and genome integrity. (A) Validate stem cell properties by testing pluripotency, karyotyping, and by cryopreserving selected stem cell colonies. (B) Confirm genomic stability of NT and MCL-1 cells using the KaryoStat+ Assay (Thermo Fisher Scientific). No aberrations were detected in cells assessed for the studies presented here.
Perform colony hESC selection and continue to maintain cells.
Once clonal lines have been stabilized, use cells to test pluripotency.
Karyotype cells to validate genomic integrity of the clones. KaryoStat+ Assay Service from Thermo Fisher was used for the cell lines presented here.
Cryopreserve cells for future experiments.
Validating knockdown of embryonic lethal genes
As previously discussed, silencing of MCL-1 in stem cells is lethal. Cells were treated with Q-VD-OPh (QVD), a pan-caspase inhibitor, to prevent cell death when MCL-1 is silenced with doxycycline (DOX) treatment. Passage MCL-1 and NT cell lines onto 6-well Matrigel-coated plates. Set up plates to have four experimental groups: -DOX, +DOX, +DOX +QVD, and +DOX +DMSO (QVD vehicle control). Once cells reach 40%–50% confluency, proceed to steps below.
Day 0
Supplement mTeSR1 and antibiotic cocktail with 1 µg/mL of DOX and/or 25 µM of QVD and add to the corresponding wells.
Days 1–2
Change media daily and continue to add DOX and/or QVD for a total of 72 h.
Day 3
Once treatment reaches 72 h, collect cells for experimental endpoint assay (e.g., quantitative RT-PCR to measure gene expression of MCL-1).
Wash cells once with 1× PBS.
Add 500 µL of TRIzol reagent. Note: If using TRIzol, make sure to collect cells in a chemical fume hood.
Scrape cells with a cell lifter and collect in a 1.5 mL centrifuge tube. Samples can be stored at -80 °C or you may proceed with RNA isolation.
RNA isolation
This protocol was adapted from Invitrogen TRIzol reagent protocol. All steps using TRIzol should be done in the chemical fume hood.
Make sure to perform RNA isolation in a RNase-free area. Spray gloves, pipettes, pipette tip boxes, tubes, tube racks, and bench with RNaseZAP.
If samples were previously frozen, allow them to incubate at RT for at least 5 min.
Add 100 µL of chloroform to sample.
Shake tubes vigorously for ~15 s.
Incubate the samples at RT for 2–3 min.
Centrifuge the samples at 12,000× g for 15 min at 4 °C.
Remove the aqueous phase of the sample and place in a new tube.
Note: Make sure to dispose of TRIzol and other hazard chemicals according to institutional policies.
Add 250 µL of 2-Propanol to the aqueous phase to precipitate RNA.
Gently invert ~10 times.
Incubate at RT for 25 min.
Centrifuge at 12,000× g for 10 min at 4 °C. Keep samples on ice from this step onwards.
Remove the supernatant from the tube, leaving only the RNA pellet.
Wash the pellet with 500 μL of 75% ethanol.
Vortex the sample briefly and then centrifuge the tube at 10,000× g for 10 min at 4 °C.
Pipette away as much of the wash as possible and allow the RNA pellet to semi-dry.
Resuspend pellet in 30 μL of DEPC-treated water. Store at -80 °C.
DNase treatment of RNA
This protocol was adapted from New England Biolabs DNase I Reaction protocol.
Set up DNase I reaction on ice (Recipe 1). The volumes indicated on the Recipe are per sample.
Incubate in the thermocycler at 37 °C for 10 min.
Inactivate the DNase I by adding 0.5 µL of 0.1 M EDTA solution to the reaction mixture.
Heat in the thermocycler for 10 min at 75 °C.
The RNA sample is now ready to be used for reverse transcription.
cDNA synthesis and quantitative RT-PCR
This protocol was adapted from Thermo Fisher High-Capacity cDNA Reverse Transcription kit and SYBR Green reagent.
Set reverse transcription master mix (Recipe 2). The volumes indicated on the Recipe are per sample.
Add 10 µL of the master mix to 10 µL of RNA sample (2 µg) in a PCR tube and mix well by vortexing. Briefly spin down tubes.
Run samples in the thermal cycler using the following conditions:
25 °C for 10 min
37 °C for 120 min
85 °C for 5 min
4 °C hold
Final cDNA reaction is 20 µL. Dilute to 200 µL by adding 180 µL of nuclease-free H2O after reaction. Studies here involved 5 µL of cDNA for qPCR.
QuantStudio 3 Real-Time PCR machine, SYBR Green master mix, and manufacturer’s instructions were used to set up the assay. The primers used to measure gene expression of MCL1 are found in Table S2. Figure 4 contains results of the experimental validation in hESCs.
Figure 4. Silencing MCL-1 in human embryonic stem cells (hESCs). Gene expression of MCL-1 is significantly downregulated in MCL-1 hESCs with doxycycline and Q-VD-OPh treatment. Normalized to control (NT +DOX +QVD). Statistical test: ordinary one-way ANOVA with Dunnett’s multiple comparisons test. DOX = doxycycline. QVD = Q-VD-OPh.
Differentiation
This is a step-by-step protocol that was adapted from Douvaras and Fossati (2015) [8].
Day -3: Replating hESCs
When the cells reach 70%–90% confluency, remove the medium and add 1 mL of Accutase.
Incubate the plate in a 37 °C incubator for 5 min.
Dilute the Accutase solution by adding 2 mL of DMEM/F12 medium.
Use a cell lifter to remove cells from the well and gently pipette the mixture 2–5 times with the p1000 pipette to fully dissociate the hESC cell colonies to single cells.
Transfer the cells to a conical tube.
Centrifuge the cells at 200× g for 4 min at RT.
Resuspend the cell pellet in 1 mL of mTeSR1 containing 10 µM Y27632 and count the cells with a hemocytometer.
Plate 8 × 104 cells per well on a Matrigel-coated 6-well plate.
Day -2: Maintaining hESCs
Aspirate and add fresh mTeSR1 media to remove Y27632.
Day -1: Maintaining hESCs
Continue to maintain cells in mTeSR1.
Day 0: Neural induction
By now, colonies should have grown evenly and reached 80% confluency. When cells have reached this point, proceed with differentiation.
Aspirate medium and add NIM to induce differentiation.
Change NIM daily until cells have reached day 8.
Day 8: Oligodendrocyte lineage induction
Aspirate medium and add N2 medium to direct cells to oligodendrocyte lineage commitment.
Day 9: Replate neural precursor cells
Remove the medium and add 1 mL of Accutase.
Incubate the plate in a 37 °C incubator for 5 min.
Dilute the Accutase solution by adding 2 mL of DMEM/F12 medium.
Use a cell lifter to remove cells from the well and gently pipette the mixture 2–5 times with the p1000 pipette to fully dissociate cells.
Transfer the cells to a conical tube.
Centrifuge the cells at 200× g for 4 min at RT.
Resuspend the cell pellet in 1 mL of N2 medium containing 10 µM Y27632 and count the cells with a hemocytometer.
Plate 1.5 × 106 cells per well on a Matrigel-coated 6-well plate. Use Matrigel-coated 35 mm glass-bottom plates for immunofluorescence. Supplement medium with 10 µM Y27632.
Day 10: Doxycycline administration to inducibly silence MCL-1
Aspirate medium and add N2 medium supplemented with 1 µg/mL doxycycline.
Change N2 medium supplemented with 1 μg/mL doxycycline every 24 h.
Day 14: Collect cells for western blot, quantitative RT-PCR, and immunofluorescence
Cell lysis and protein extraction for western blotting
Prepare lysis buffer on ice. Make enough to use 100 mL per well of a 6-well plate.
Aspirate media from wells.
Wash once with 1× PBS.
Add 100 µL of lysis buffer to each well.
Scrape wells vigorously with cell lifter and transfer sample to a 1.5 µL centrifuge tube.
Place tubes on ice for 30 min. Vortex every 10 min.
Spin down samples at 14,000× g for 30 min.
Collect supernatant and transfer to a new 1.5 µL centrifuge tube.
Store sample at -20 °C.
Measuring protein expression by western blot
Prepare SDS for Tris-Gly-SDS buffer.
Dissolve 200 g of SDS in 900 mL of ddH2O.
Heat to 68 °C and stir with a magnetic stirrer to assist dissolution.
If necessary, adjust the pH to 7.2 by adding a few drops of concentrated HCl.
Adjust the volume to 1 L with ddH2O.
Determine protein concentration using the Thermo Scientific BCA Protein Assay kit.
Dilute 30 µg of protein in 1% Triton and LDS with 2-Mercaptoethanol.
Incubate samples at 95 °C for 5 min.
Run samples on a 10-well 4%–20% Mini-Protean TGX precast protein gel in Tris-Gly-SDS buffer.
Transfer gel onto polyvinylidene difluoride membranes at 4 °C overnight.
Block membrane in 5% milk diluted in TBST for one hour at RT on a benchtop rocker.
Incubate with primary antibody overnight on a benchtop rocker. The antibodies used here can be found in Table S3.
Wash membrane with TBST for 5 min on a benchtop rocker. Repeat two more times.
Incubate with HRP-conjugated secondary antibodies against mouse or rabbit IgG for 1 h at RT on a benchtop rocker.
Wash membrane with TBST for 5 min on a benchtop rocker. Repeat two more times.
Develop blots with ECL Plus reagent and image on the Chemiluminescent Imager.
Bands were quantified with Image Studio Lite. Figure 5A and 5B contain the results of the experimental validation in committed oligodendrocyte neural precursor cells.
Figure 5. Silencing MCL-1 in oligodendrocyte neural precursor cells. (A) Representative image of MCL-1 protein expression and cleaved PARP protein expression in committed oligodendrocyte neural precursor cells following 96 h of doxycycline treatment in NT and MCL-1 CRISPRoff cell lines. (B) MCL-1 is significantly downregulated in committed oligodendrocyte neural precursor cells following 96 h doxycycline treatment in MCL-1 cell line. Normalized to respective controls (NT -DOX and MCL-1-DOX). Statistical test: Student’s t-tests. (C) Cleaved PARP is significantly upregulated in oligodendrocyte neural precursor cells following 96 h doxycycline treatment in MCL-1 cell line. Normalized to respective controls (NT -DOX and MCL-1 -DOX). Statistical test: multiple t-tests. DOX = doxycycline. QVD = Q-VD-OPh.
Immunofluorescence
Fix cells with 4% PFA in 1× PBS for 20 min at RT.
Permeabilize cells with 1% Triton for 10 min at RT.
Block with 10% BSA prepared in 1× PBS for 1 h at RT.
Incubate with primary antibody diluted in 10% BSA overnight at 4 °C. The antibodies used here can be found in Table S3.
Wash with 1× PBS. Repeat two more times.
Incubate with secondary antibody diluted in 10% BSA at RT for 1 h.
Wash with 1× PBS. Repeat two more times.
Incubate with Hoechst for 10 min.
Wash with 1× PBS.
Mount with Fluoromount-G slide mounting medium.
Image cells on a confocal microscope. Silencing MCL-1 in committed oligodendrocyte neural precursor cells leads to changes in expression of OLIG2 and NKX2.2 (Figure 6).
Figure 6. Dysregulation of transcription factors that modulate oligodendrocyte lineage commitment following MCL-1 suppression. (A) Gene expression of OLIG2 was significantly increased in committed OL neural precursor cells (NPCs) following 96 h doxycycline treatment in MCL-1 cell line. Normalized to respective controls (NT -DOX and MCL-1 -DOX). Statistical test: multiple t-tests. (B) Gene expression of NKX2.2 was significantly increased in committed OL NPCs following 96 h doxycycline treatment in MCL-1 cell line. Normalized to respective controls (NT -DOX and MCL-1 -DOX). Statistical test: multiple t-tests. (C) Silencing of MCL-1 upregulates protein expression of OLIG2 and downregulates NKX2.2 in committed OL NPCs. DOX = doxycycline. Scale bar = 50 µm.
Data analysis
Detailed description of analysis can be found in the figure legends.
Validation of protocol
We validated the use of this protocol by silencing MCL-1, an anti-apoptotic protein, at an early state of oligodendrocyte differentiation in vitro. MCL-1 is required for the survival of hPSCs in vitro and thus needs to be silenced after the initiation of the differentiation process. All experiments presented have been replicated using three or more technical replicates and at least three biological replicates. Controls included NT sgRNAs.
General notes and troubleshooting
General notes
The maintenance of the stem cells should be rigorously tested to assure they are pluripotent at the start of the protocol.
Dose response of the selection drugs is essential to increase the efficiency of silencing.
Oligodendrocyte markers and primers should be rigorously validated.
Acknowledgments
This protocol was adapted from previous studies [4, 8]. This work was supported by 1R35 GM128915-01NIGMS (VG), 1RF1MH123971-01 (VG), and NSF CBET 2033800 (JMB); the Precision Medicine and Mental Health Initiative sponsored by the Vanderbilt Brain Institute (VG), HHMI Gilliam (MG). The spinning disk confocal microscopy imaging and image analysis were performed in part through the Vanderbilt Cell Imaging Shared Resource (supported by NIH grants CA68485, DK20593, DK58404, DK59637, and EY08126). Some figures in this manuscript (Figures 2, 3a, and 4a) were created using BioRender (BioRender.com).
Competing interests
The authors declare no competing financial interests.
Ethical considerations
The human embryonic stem cell line H9 (WA09) was obtained from WiCell Research Institute (Wisconsin). All experiments using hESCs were performed using the WA09 (H9) cell line under the supervision of the Vanderbilt Institutional Human Pluripotent Cell Research Oversight (VIHPCRO) Committee (Protocol IRB # 160146 to VG).
References
Tschaharganeh, D. F., Lowe, S. W., Garippa, R. J. and Livshits, G. (2016). Using CRISPR/Cas to study gene function and model disease in vivo. FEBS J 283(17): 3194–3203. doi: 10.1111/febs.13750
Valenti, M. T., Serena, M., Carbonare, L. D. and Zipeto, D. (2019). CRISPR/Cas system: An emerging technology in stem cell research. World J. Stem Cells 11(11): 937–956. doi: 10.4252/wjsc.v11.i11.937
Boettcher, M. and McManus, M. T. (2015). Choosing the Right Tool for the Job: RNAi, TALEN, or CRISPR. Mol. Cell 58(4): 575–585. doi: 10.1016/j.molcel.2015.04.028
Nuñez, J. K., Chen, J., Pommier, G. C., Cogan, J. Z., Replogle, J. M., Adriaens, C., Ramadoss, G. N., Shi, Q., Hung, K. L., Samelson, A. J., et al. (2021). Genome-wide programmable transcriptional memory by CRISPR-based epigenome editing. Cell 184(9): 2503–2519.e17. doi: 10.1016/j.cell.2021.03.025
Rinkenberger, J. L., Horning, S., Klocke, B., Roth, K. and Korsmeyer, S. J. (2000). Mcl-1 deficiency results in peri-implantation embryonic lethality. Genes Dev. 14(1): 23–27. doi: 10.1101/gad.14.1.23
Rasmussen, M. L., Taneja, N., Neininger, A. C., Wang, L., Robertson, G. L., Riffle, S. N., Shi, L., Knollmann, B. C., Burnette, D. T., Gama, V., et al. (2020). MCL-1 Inhibition by Selective BH3 Mimetics Disrupts Mitochondrial Dynamics Causing Loss of Viability and Functionality of Human Cardiomyocytes. iScience 23(4): 101015. doi: 10.1016/j.isci.2020.101015
Rasmussen, M. L., Kline, L. A., Park, K. P., Ortolano, N. A., Romero-Morales, A. I., Anthony, C. C., Beckermann, K. E. and Gama, V. (2018). A Non-apoptotic Function of MCL-1 in Promoting Pluripotency and Modulating Mitochondrial Dynamics in Stem Cells. Stem Cell Rep. 10(3): 684–692. doi: 10.1016/j.stemcr.2018.01.005
Douvaras, P. and Fossati, V. (2015). Generation and isolation of oligodendrocyte progenitor cells from human pluripotent stem cells. Nat. Protoc. 10(8): 1143–1154. doi: 10.1038/nprot.2015.075
Horlbeck, M. A., Gilbert, L. A., Villalta, J. E., Adamson, B., Pak, R. A., Chen, Y., Fields, A. P., Park, C. Y., Corn, J. E., Kampmann, M., et al. (2016). Compact and highly active next-generation libraries for CRISPR-mediated gene repression and activation. eLife 5: e19760. doi: 10.7554/elife.19760
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. doi: 10.1038/s41467-018-07901-8
Sambrook, J. and Russell, D. W. (2006). Preparation and Transformation of Competent E. coli Using Calcium Chloride. Cold Spring Harb. Protoc. 2006(1): 3932. doi: 10.1101/pdb.prot3932
Supplementary information
The following supporting information can be downloaded here:
Table S1. MCL-1 and NT sgRNAs
Table S2. MCL-1 primer sequences
Table S3. Antibodies
Article Information
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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Expansion and Polarization of Human γδT17 Cells in vitro from Peripheral Blood Mononuclear Cells
XC Xu Chen
XH Xiaoling Hu
FC Fuxiang Chen
JY Jun Yan
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4914 Views: 619
Reviewed by: Vivien J. Coulson-Thomas Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances May 2022
Abstract
γδ T cells play a critical role in homeostasis and diseases such as infectious diseases and tumors in both mice and humans. They can be categorized into two main functional subsets: IFN-γ-producing γδT1 cells and IL-17-producing γδT17 cells. While CD27 expression segregates these two subsets in mice, little is known about human γδT17 cell differentiation and expansion. Previous studies have identified γδT17 cells in human skin and mucosal tissues, including the oral cavity and colon. However, human γδ T cells from peripheral blood mononuclear cells (PBMCs) primarily produce IFN-γ. In this protocol, we describe a method for in vitro expansion and polarization of human γδT17 cells from PBMCs.
Key Features
• Expansion of γδ T cells from peripheral blood mononuclear cells.
• Human IL-17A-producing γδ T-cell differentiation and expansion using IL-7 and anti-γδTCR.
• Analysis of IL-17A production post γδ T-cell expansion.
Keywords: γδT17 cell γδ T cell IL-17 Peripheral blood mononuclear cells
Background
γδ T cells are a group of lymphocytes consisting of a γ and δ chain, being considered as a bridge linking innate and adaptive immunity (Melandri et al., 2018). γδ T cells are mainly found in cutaneous and mucosal tissues such as the skin, gut, and oral mucosa (Cai et al., 2011; Wu et al., 2014; Hovav et al., 2020), but they also circulate in peripheral blood (Davey et al., 2018). γδ T cells are commonly classified based on their TCR chains and cytokine production. In mice, γδ T cells bear different Vγ chains ranging from Vγ1 to Vγ7, according to Heilig and Tonegawa nomenclature (Heilig and Tonegawa, 1986). Murine γδ T cells are heterogenous, secrete different cytokines, and can be divided into two main functional subsets: IFN-γ-producing γδT1 cells and IL-17-producing γδT17 cells (Ribot et al., 2009). Surface markers such as CD27 and CCR6 can be used to define these two subsets (Haas et al., 2009; Ribot et al., 2009). In humans, γδ T cells can be distinguished by the δ chain, including Vδ1, Vδ2, Vδ3, and Vδ5 (Ling et al., 2022). While previous studies have reported methods for in vitro expansion of human γδ T cells (Ness-Schwickerath et al., 2010; Caccamo et al., 2011; Hur et al., 2023), the majority of peripheral blood γδ T cells in humans are Vδ2+ subsets that mainly produce IFN-γ. It is difficult to investigate human γδT17 cells as they are scarce, and little has been done to expand human γδT17 cells in vitro. A previous report from Michel et al. demonstrated that IL-7 promotes the expansion of human IL-17-producing γδ T cells (Michel et al., 2012). In this study, we describe a modified method for in vitro polarization of human γδT17 cells from peripheral blood mononuclear cells (PBMCs) (Chen et al., 2022).
Materials and reagents
24-well plate (Falcon, catalog number: 353047)
6-well plate (NEST, catalog number: 703001)
Blood-drawing tubes containing EDTA-K2 (Improve Medical, catalog number: 101680720)
12 × 75 mm plastic tubes (Falcon, catalog number: 352052)
Anti-human γδTCR Ab (Beckman, clone: IMMU510, IM1349)
Human IL-7 (R&D system, catalog number: 207-IL-005/CF)
RPMI-1640 medium (Sigma-Aldrich, catalog number: R8758-500ML)
2-Mercaptoethanol (Gibco, catalog number: 21985023)
Sterile PBS (Sangon Biotech, catalog number: E607008-0500)
Fetal bovine serum (FBS) (Atlanta Biologicals, catalog number: S11150)
Penicillin-Streptomycin liquid (100×) (Solarbio, catalog number: P1400)
Trypan Blue (STEMCELL Technologies, catalog number: 07050)
Lympholyte® cell separation media (Ficoll) (Cedarlane Laboratories, catalog number: CL5020)
Anti-human IL-17A (BioLegend, catalog number: 512306, referred to as anti-human IL-17 later in this protocol)
Anti-human CD3 (BioLegend, catalog number: 300328)
Anti-human γδTCR (Miltenyi Biotec, catalog number: 130-113-508)
Anti-human CCR6 (BioLegend, catalog number: 353412)
Viability dye (Invitrogen, catalog number: 65-0865-14)
GolgiPlug (Brefeldin A solution 1,000×) (BioLegend, catalog number: 420601)
Phorbol 12-myristate 13-acetate (PMA) (Millipore Sigma, catalog number: P8139)
Ionomycin (Millipore Sigma, catalog number: I0634)
Human TruStain FcXTM (BioLegend, catalog number: 422302)
Fixation buffer (BioLegend, catalog number: 420801)
Intracellular staining perm 10× wash buffer (BioLegend, catalog number: 421002, referred to as wash buffer later in this protocol)
Equipment
CO2 incubator (Thermo Fisher Scientific, model number: 3543 or 3111)
Centrifuge (Beckman Coulter, model: Allegra® X-15R, catalog number: 392934)
Laminar flow hood (Scitech Equipments Ltd., model: EVL-5S, catalog number: ZX0907-04)
Flow cytometry (BD Bioscience, model: FACSCantoTM II, catalog number: 338962)
Pipettes (multi-channel, Eppendorf)
Software and Datasets
FlowJo v10.8.1 (BD Biosciences)
Procedure
Recipes for preparation
Make complete RPMI-1640 medium: RPMI-1640 medium + 10% FBS + 1% Penicillin-Streptomycin liquid + 2-Mercaptoethanol (0.1 mL/L).
Make anti-human γδTCR solution for coating: 0.1 mg of human γδTCR Ab (clone: IMMU510, IM1349) + 500 μL of PBS = 0.2 μg/μL human γδTCR Ab.
Make working stock of viability dye: 1 μL of viability dye + 49 μL of PBS (Viability dye:PBS = 1:49).
Make 1× wash buffer: 1 mL of 10× wash buffer + 9 mL of H2O (10× wash buffer:H2O = 1:9).
PBMC isolation from peripheral blood (see Note 1)
This section provides PBMCs for polarization of human γδ T cells in Procedure C.
Draw peripheral blood from healthy donors and collect approximately 15–20 mL of blood with EDTA-K2-containing tubes in accordance with institutional ethics and safety protocols.
Dilute blood samples with an equal volume of RPMI-1640 medium.
Carefully layer the diluted blood suspension (5 mL) over 5 mL of Ficoll in a 15 mL conical tube.
Centrifuge at 931× g for 15 min at room temperature without brakes.
Carefully transfer the mononuclear cell layer to a new 15 mL conical tube [the mononuclear cell layer is the white layer between topside plasma layer and Ficoll layer (Figure 1A); the colors from top to down are light yellow, white, clear, and red]. The white layer must be collected.
Fill the conical tube with complete RPMI-1640 medium to 10 mL.
Mix gently by hand (up and down mixing) and centrifuge at 524× g for 10 min at 4 °C.
Carefully remove and discard completely the supernatant by pipetting. The cells are at the bottom of the 15 mL conical tube.
Wash the cells with 5 mL of complete RPMI-1640 medium. Specifically, resuspend the cells in 5 mL of complete RPMI-1640 medium and centrifuge at 524× g for 10 min at 4 °C. Then, completely remove the supernatant by pipetting.
Resuspend the cell pellet in 5 mL of complete RPMI-1640 medium and count the cell numbers directly with a hemocytometer under a microscope or using Trypan blue staining (see Note 2).
In vitro polarization
This section provides polarization steps for human γδ T cells.
Coat a 24-well plate with PBS containing anti-human γδTCR Ab. Please note that plates need to be coated one day before cell isolation and polarization. Add 245 μL of PBS per well and then add 5 μL of 0.2 μg/μL anti-human γδTCR Ab per well to achieve a final concentration of 1 μg anti-human γδTCR Ab/250 μL of PBS. Incubate overnight (18–24 h) in a 37 °C incubator with 5% CO2.
Aspirate and discard the coating antibody solution from the 24-well plate on the second day (see Note 3).
Isolate PBMCs from healthy donors’ peripheral blood (Procedure B) and dilute PBMC prepared in Procedure B with complete RPMI-1640 medium to an appropriate concentration.
Seed PBMCs (1–4 million per well in 1 mL of complete RPMI-1640 medium containing 20 ng/mL human IL-7) into the 24-well plate, which has been pre-coated with anti-human γδTCR Ab.
Check the status of cells every day under a microscope to exclude contamination. Healthy cells will appear bright (see Note 4). If cells are contaminated with bacteria, the medium will appear cloudy. Sudden drops in the pH of the culture medium (the color of the culture medium turns yellow) are also signs of contamination.
According to the cell density in the plate, transfer and passage the cells to a new 6-well plate (not coated with anti-human γδTCR Ab) on the third to fourth day using complete RPMI-1640 medium containing 20 ng/mL human IL-7. Transfer and passage the cells before two-thirds of plate confluency (see Note 5). Please note that γδ T cells are suspension cells, and passage of cells can be performed by aspirating 1 mL of cells medium by pipetting into a new well in a 6-well plate and adding 2 mL of complete RPMI-1640 medium containing 20 ng/mL human IL-7 into this new well.
Check the status of cells every day under a microscope to exclude any contaminations and make sure the cells appear healthy (bright).
Passage the cells using complete RPMI-1640 medium containing 20 ng/mL human IL-7 when necessary.
Culture the cells for 2–3 weeks (approximately 4–5 passages).
Collect the cells by aspirating 1 mL of cells medium by pipetting and perform intracellular cytokine staining of IL-17 with the cells (Procedure D)
Data acquisition and analysis
This procedure provides an analysis of IL-17 production from human γδ T cells after polarization.
Stimulate approximately 100,000 cells that have been polarized in Procedure C with PMA (final concentration: 50 ng/mL) and ionomycin (final concentration: 1 μg/mL) in the presence of GolgiPlug (final concentration: 1× in the medium) in a total 250 μL of complete RPMI-1640 medium per well using 24-well plate.
Incubate for 4–6 h in a 37 °C incubator with 5% CO2.
Collect the cell suspension (T cells are non-adherent cells) into a 12 × 75 mm plastic tube by pipetting from the wells in the 24-well plate and wash the wells with 1 mL of PBS (also collect by pipetting).
Centrifuge at 524× g for 5 min at 4 °C and discard the supernatant.
Add human TruStain FcXTM (Fc receptor blocking solution, 5 μL/test) and incubate at 4 °C for 10 min.
Add commercial antibodies for surface staining, including working stock of viability dye (5 μL/test), anti-human CD3 (5 μL/test), anti-human γδTCR (3 μL/test), and anti-human CCR6 (5 μL/test) according to the instructions. Incubate at 4 °C for 30 min.
Wash the cells with 1 mL of PBS and centrifuge at 524× g (1,500 rpm) for 5 min at 4 °C.
Completely discard the supernatant by pipetting.
Add 400 μL of fixation buffer per tube and vortex.
Fix the cells at room temperature for 20 min in the dark.
Wash the cells with 1 mL of 1× wash buffer and centrifuge at 524× g for 5 min at 4 °C.
Discard the supernatant and wash the cells with 1 mL of 1× wash buffer again.
Centrifuge at 524× g for 5 min at 4 °C and discard the supernatant.
Add anti-human IL-17 antibody (5 μL/test) and incubate at 4 °C overnight (see Note 6).
Wash the cells with 1 mL of 1× wash buffer and centrifuge at 524× g for 5 min at 4 °C.
Discard the supernatant and resuspend the cells.
Acquire the cells on FACSCantoTM II flow cytometer (see Note 7).
Analyze data with FlowJo software (see Notes 8 and 9, Figures 1B, C, and 2).
Figure 1. Human γδT17 expansion in vitro from peripheral blood mononuclear cells (PBMCs). A. Overall workflow of this protocol. B. Percentage of human γδ T cells after expansion. Cells were gated on CD3+ cells. C. Production of IL-17 by human γδ T cells after polarization from two different donors. Cells were gated on γδ TCR+ cells.
Figure 2. Analysis of IL-17 production in γδ T cells from peripheral blood mononuclear cells (PBMCs) before and after polarization. A. Representative dot plot and summarized data of IL-17 production in primary γδ T cells from PBMCs. B. Representative dot plot and summarized data of IL-17 production in polarized γδ T cells (Chen et al., 2022). Each dot represents one individual donor, and student t-test was used for statistical analysis; *** p < 0.001.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Chen et al. (2022). Differential metabolic requirement governed by transcription factor c-Maf dictates innate γδT17 effector functionality in mice and humans. Science Advances (Figure 6, panel A).
Notes
A 24-well plate should be coated with PBS containing anti-human γδTCR Ab on the day before PBMC isolation; details can be found in the first step of Procedure C.
Count the cell numbers using a hemocytometer under a microscope directly or use Trypan blue staining. Either method is suitable since the cells are mostly viable. When counting cells directly, healthy cells appear bright; when counting cells using Trypan blue staining, live cells remain unstained and bright while dead cells are stained with the blue dye. If 10 μL of cell suspension is mixed with 90 μL of Trypan blue staining solution, the dilution factor is 10 when calculating the total cell numbers.
There are no washing steps after discarding the coating solution.
Healthy cells under the microscope appear bright, as shown in Figure 3.
Figure 3. Morphology of the cells under a microscope. Different densities of the cells are shown: low density (A) and high density (B).
Passage the cells according to the cell density; two-thirds of the plate confluency is considered a target density, as shown in Figure 3B.
Incubate anti-human IL-17 according to the manufacturer’s instructions. We usually incubate anti-human IL-17 Ab overnight.
The analysis of IL-17 production in γδ T cells (gating strategy) is as follows: 1) gating live cells (viability dye negative), 2) gating CD3-positive cells, 3) gating CD3-positive and γδTCR-positive cells (Figure 1B), 4) gating IL-17-positive cells on γδ T cells (Figure 1C).
The percentage of γδ T cells increased to 64.8% of CD3+ cells after expansion (Figure 1B). However, the percentage of IL-17-producing γδ T cells varies depending on different donors. As shown in Figure 1C after polarization, the percentages of IL-17+ γδ T cells were 7.59% and 1.03% in Donor A and Donor B, respectively.
The percentage of γδ T cells in CD3+ T cells post polarization is approximately a 10-fold increase. Primary γδ T cells from PBMCs produce scarce IL-17 (Figure 2A). After polarization, IL-17+ γδ T cells are predominantly CCR6+ γδ T cells, and the percentage of IL-17+ γδ T cells in the total γδ T-cell population is approximately 10% on average (Figure 2B) (Chen et al., 2022).
Acknowledgments
This work was supported by the NIH R01AI128818 and R01CA213990. Xu Chen was supported by the Shanghai Pujiang Program (23PJD047) and on-job postdoctoral program.
Competing interests
The authors declare no conflicts of interest.
Ethical considerations
All experimental procedures have been approved by the Ethics Committee of the Shanghai Ninth People’s Hospital, Shanghai, China.
References
Caccamo, N., La Mendola, C., Orlando, V., Meraviglia, S., Todaro, M., Stassi, G., Sireci, G., Fournié, J. J. and Dieli, F. (2011). Differentiation, phenotype, and function of interleukin-17–producing human Vγ9Vδ2 T cells. Blood 118(1): 129–138.
Cai, Y., Shen, X., Ding, C., Qi, C., Li, K., Li, X., Jala, V. R., Zhang, H. g., Wang, T., Zheng, J., et al. (2011). Pivotal Role of Dermal IL-17-Producing γδ T Cells in Skin Inflammation. Immunity 35(4): 596–610.
Chen, X., Cai, Y., Hu, X., Ding, C., He, L., Zhang, X., Chen, F. and Yan, J. (2022). Differential metabolic requirement governed by transcription factor c-Maf dictates innate γδT17 effector functionality in mice and humans. Sci. Adv. 8(21): eabm9120.
Davey, M. S., Willcox, C. R., Hunter, S., Kasatskaya, S. A., Remmerswaal, E. B. M., Salim, M., Mohammed, F., Bemelman, F. J., Chudakov, D. M., Oo, Y. H., et al. (2018). The human Vδ2+ T-cell compartment comprises distinct innate-like Vγ9+ and adaptive Vγ9- subsets. Nat. Commun. 9(1): e1038/s41467–018–04076–0.
Haas, J. D., González, F. H. M., Schmitz, S., Chennupati, V., Föhse, L., Kremmer, E., Förster, R. and Prinz, I. (2009). CCR6 and NK1.1 distinguish between IL-17A and IFN-γ-producing γδ effector T cells. Eur. J. Immunol. 39(12): 3488–3497.
Heilig, J. S. and Tonegawa, S. (1986). Diversity of murine gamma genes and expression in fetal and adult T lymphocytes. Nature 322(6082): 836–840.
Hovav, A. H., Wilharm, A., Barel, O. and Prinz, I. (2020). Development and Function of gammadeltaT Cells in the Oral Mucosa. J. Dent. Res. 99(5): 498–505.
Hur, G., Choi, H., Lee, Y., Sohn, H. J., Kim, S. Y. and Kim, T. G. (2023). In Vitro Expansion of Vδ1+ T Cells from Cord Blood by Using Artificial Antigen-Presenting Cells and Anti-CD3 Antibody. Vaccines 11(2): 406.
Ling, S., You, Z., Li, Y., Zhang, J., Zhao, S., He, Y. and Chen, X. (2022). The role of γδ T17 cells in cardiovascular disease. J. Leukoc. Biol. 112(6): 1649–1661.
Melandri, D., Zlatareva, I., Chaleil, R. A. G., Dart, R. J., Chancellor, A., Nussbaumer, O., Polyakova, O., Roberts, N. A., Wesch, D., Kabelitz, D., et al. (2018). The γδTCR combines innate immunity with adaptive immunity by utilizing spatially distinct regions for agonist selection and antigen responsiveness. Nat. Immunol. 19(12): 1352–1365.
Michel, M. L., Pang, D. J., Haque, S. F. Y., Potocnik, A. J., Pennington, D. J. and Hayday, A. C. (2012). Interleukin 7 (IL-7) selectively promotes mouse and human IL-17–producing γδ cells. Proc. Natl. Acad. Sci. U.S.A. 109(43): 17549–17554.
Ness-Schwickerath, K. J., Jin, C. and Morita, C. T. (2010). Cytokine Requirements for the Differentiation and Expansion of IL-17A– and IL-22–Producing Human Vγ2Vδ2 T Cells. J. Immunol. 184(12): 7268–7280.
Ribot, J. C., deBarros, A., Pang, D. J., Neves, J. F., Peperzak, V., Roberts, S. J., Girardi, M., Borst, J., Hayday, A. C., Pennington, D. J., et al. (2009). CD27 is a thymic determinant of the balance between interferon-γ- and interleukin 17–producing γδ T cell subsets. Nat. Immunol. 10(4): 427–436.
Wu, P., Wu, D., Ni, C., Ye, J., Chen, W., Hu, G., Wang, Z., Wang, C., Zhang, Z., Xia, W., et al. (2014). δγT17 cells promote the accumulation and expansion of myeloid-derived suppressor cells in human colorectal cancer. Immunity 40(5): 785–800.
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Development of Recombinase Polymerase Amplification–Lateral Flow Dipstick (RPA–LFD) as a Rapid On-Site Detection Technique for Fusarium oxysporum
SH Shuodan Hu
HY Hong Yu
CZ Chuanqing Zhang
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4915 Views: 667
Reviewed by: Shweta PanchalMichael EnosSoumya Moonjely
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Original Research Article:
The authors used this protocol in Plant Disease Feb 2023
Abstract
Fusarium oxysporum can cause many important plant diseases worldwide, such as crown rot, wilt, and root rot. During the development of strawberry crown rot, this pathogenic fungus spreads from the mother plant to the strawberry seedling through the stolon, with obvious characteristics of latent infection. Therefore, the rapid and timely detection of F. oxysporum can significantly help achieve effective disease management. Here, we present a protocol for the recombinase polymerase amplification– lateral flow dipstick (RPA–LFD) detection technique for the rapid detection of F. oxysporum on strawberry, which only takes half an hour. A significant advantage of our RPA–LFD technique is the elimination of the involvement of professional teams and laboratories, which qualifies it for field detection. We test this protocol directly on plant samples with suspected infection by F. oxysporum in the field and greenhouse. It is worth noting that this protocol can quickly, sensitively, and specifically detect F. oxysporum in soils and plants including strawberry.
Key features
• This protocol is used to detect whether plants such as strawberry are infected with F. oxysporum.
• This protocol has potential for application in portable nucleic acid detection.
• It can complete the detection of samples in the field within 30 min.
Graphical overview
Keywords: Fusarium oxysporum Strawberry Crown rot RPA-LFD Rapid detection
Background
Strawberry crown rot caused by Fusarium oxysporum has seriously affected strawberry yield [1]. The pathogen is transmitted from the mother plant to the strawberry seedling through the stolon [2]. Due to the particularities of pathogen transmission, the control of this disease is difficult. Therefore, rapid and accurate detection of F. oxysporum in the early stages is of utmost importance for preventing and controlling crown rot disease.
Koch’s postulates are the traditional method for pathogen detection through isolation, culture, and pathogenicity testing, being time-consuming and requiring professional training. Molecular detection and identification of the pathogen are highly accurate and sensitive alternative methods for disease diagnosis. Polymerase chain reaction (PCR), quantitative PCR, and other molecular detection techniques have been invented to detect pathogens [3, 4]. However, these techniques require professional experience in performing the tests and expensive equipment, which may not be possible in on-site detection. Altogether, these create a bottleneck for early-disease diagnosis. Fortunately, other molecular biological methods can overcome these limitations, such as the recombinase polymerase amplification (RPA) detection technique [5]. RPA is performed at constant temperature, and its most significant advantage is that the detection results can be visualized by lateral flow dipstick (LFD) [6]. This method is simple to operate and requires little laboratory equipment. Therefore, it has potential for application in portable nucleic acid detection.
Here, we develop and evaluate the RPA–LFD detection technique that can quickly, sensitively, and specifically detect F. oxysporum infection in strawberry [7]. By comparing the genomes of F. oxysporum with other strains common in soil and strawberry plants, we found that the sequence of CYP51C gene was unique and highly variable in Fusarium spp.; therefore, we chose the CYP51C gene to design primers and probes for F. oxysporum detection. We demonstrate how F. oxysporum can be accurately detected on-site. This protocol requires no professional experience, is easy to operate, and the results can be observed within 8 min at 39 °C. This advantage makes it a valuable diagnostic tool, especially in the absence of experienced microscopists. The sensitivity of this protocol is consistent with PCR, and it is accurate enough to detect F. oxysporum. In summary, this protocol is validated on the early field diagnosis of strawberry crown rot. Additionally, it can be applied to detect F. oxysporum in soils and other plants that are damaged by this fungus and can be adjusted to detect other pathogens, only by redesigning specific primers and probes according to the genome sequence of the pathogen to be detected.
Materials and reagents
Biological materials
DNA of F. oxysporum, F. tricinctum, F. proliferatum, F. fujikuroi, F. graminearum, F. solani, F. equiseti, Colletotrichum siamense, C. gloeosporioides, C. aenigma, C. fructicola, Botrytis cinerea, Phoma sp., and Stagonosporopsis sp.
Strawberry plant
Potato
Reagents
KOH (SangonBiotech, catalog number: A610441)
Polyethylene glycol (PEG200) (SangonBiotech, catalog number: A601780-0500)
EDTA disodium salt dihydrate (SangonBiotech, catalog number: A610185-050)
CTAB, cetyltrimethylammonium bromide (SangonBiotech, catalog number: A600108-0500)
Sodium chloride (NaCl) (SangonBiotech, catalog number: A610476-0001)
Tris (SangonBiotech, catalog number: A610195-0500)
Ethanol absolute (SangonBiotech, catalog number: A500737-0500)
Isopropanol (Shanghai Hushi, catalog number: 180109218)
Trichloromethane (Shanghai Hushi, catalog number: 10006818)
Dextrose (SangonBiotech, catalog number: A610219-0500)
Agar (SangonBiotech, catalog number: A505255-025)
Primer and probe:
F91:5′-TCAACTGGCATCGTCAACATCACCGAAGTAA-3′
R91:5′ Biotin-CCAGGCATGACGAAGTTGATAGGTTGAAAGC-3′
Probe:5′ 6-FAM-AGGCTCCCTCCTCGGTAACGAAGTCCGCTCCA/idSp/GTTTGACAGCACATT-3′ C3 Spacer
Solutions
CTAB extracting solution (see Recipes)
70% Ethanol absolute (see Recipes)
60% PEG200 with pH 13.2–13.5 (see Recipes)
2 M KOH (see Recipes)
Potato dextrose agar (PDA) (see Recipes)
Recipes
CTAB extracting solution
*Note: The solute is first mixed and then ddH2O is added to the constant volume of 1,000 mL.
Reagent Final concentration Quantity or Volume
CTAB
Tris
EDTA
NaCl
H2O
2%
100 mM
20 mM
1.4 M
n/a
20 g
12.114 g
7.448 mL
81.816 g
to 1,000 mL
Total n/a 1,000 mL
70% ethanol absolute
Reagent Final concentration Quantity or Volume
Ethanol absolute
H2O
70% (v/v)
n/a
70 mL
30 mL
Total n/a 100 mL
2 M KOH
*Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
KOH
H2O
2 M
n/a
1.12 g
10 mL
Total n/a 10 mL
60% PEG200 with pH 13.2–13.5
*Note: This solution needs to be prepared and used immediately.
Reagent Final concentration Quantity or Volume
KOH (Recipe 3)
PEG200
H2O
n/a
60% (v/v)
n/a
0.33 mL
8 mL
11.67 mL
Total n/a 20 mL
Potato dextrose agar (PDA)
*Note: Boil the potato in 1,000 mL of H2O using an induction cooker and pan and filter. Then, add the dextrose and agar to the filtrate.
Reagent Final concentration Quantity or Volume
Potato
Dextrose
Agar
H2O
200 g/L
20%
15%
n/a
200 g
20 g
15 g
1,000 mL
Total n/a 1,000 mL
Laboratory supplies
1.5 mL microcentrifuge tubes (Axygen, catalog number: MCT-150-C)
HybriD DNA thermostat rapid amplification kit (APWL, catalog number: WLN8203KIT)
HybriDetect lateral flow dipstick (APWL, catalog number: WLFS8201)
Forceps (Shanghai Leigu, catalog number: W-002903)
Sterile blade (VWR International, catalog number: 55411-050)
Beakers (Shanghai Leigu, catalog number: B-000103)
Stir bars (Shanghai Leigu, catalog number: B-040315)
Graduated cylinders (Shanghai Leigu, catalog number: B-031603)
Glass bottles (Shanghai Leigu, catalog number: B-W00323)
90 mm Petri dish (Nantong Baiyao, catalog number: YJ-90-12g)
Inoculating needle (Shanghai Leigu, catalog number: W-006001)
Forceps (Shanghai Leigu, catalog number: W-002806)
Quartz sand (SangonBiotech, catalog number: A500823-0001)
Equipment
Micropipettes 0.1–2.5 µL, 2–20 µL, 20–200 µL (Eppendorf, model: 3123000217, 3123000233, 3123000250)
Incubator, 25 °C (Shanghai Boxun, model: MJX-250B-Z)
4 °C refrigerator (Haier, model: SC-339JN)
Grinding machine (Shanghai Jingxin, model: JXFSTPRP-24L)
Ultramicro accounting protein analyzer (BioDrop, model: BioDrop µlite+)
Centrifuge (ThermoFisher, model: 75002440)
Thermostat (Hangzhou Aosheng, model: 028-14484-20110001)
Balance (Shanghai Haosheng, model: JA303A)
Induction cooker (Midea, model: RT21E0105)
Pan (Supor, model: ST22P1)
Vortex shaker (SangonBiotech, model: WH-861)
Software and datasets
DNAMAN v6.0.3.99
Snapgene v.5.2.4
Procedure
Design of primers and probes
The sequence of the CYP51C gene was unique and highly variable in Fusarium spp. Therefore, we used DNAMAN v6.0.3.99 to design primers based on the CYP51C sequence of the common Fusarium species in soil and plants, such as F. tricinctum, F. proliferatum, F. fujikuroi, F. graminearum, F. solani, and F. equiseti. The specific sequence of F. oxysporum was found by CYP51C gene sequence to design specific primers (Figure 1).
Recommended primers length was between 31 and 32 bp. Shorter primers can affect amplification speed and detection sensitivity, while longer primers can form secondary structures that affect amplification. The pairs of primers will be as follows:
F91: 5′-TCAACTGGCATCGTCAACATCACCGAAGTAA-3′
R91: 5′-CCAGGCATGACGAAGTTGATAGGTTGAAAGC-3′
Biotin label was applied to the 5′ end of the screened downstream primers R91. Tailed primers will be as follows:
5′ Biotin-CCAGGCATGACGAAGTTGATAGGTTGAAAGC-3′.
The probe was designed with an antigen marker (6-FAM) modification at the 5′ end, a modified group (C3 Spacer) at the 3′ end, and a dSpacer (tetrahydrofuran, THF) labeled at the middle of the 5′ and 3′ ends as the recognition site for NFO. C3 Spacer was blocking groups that prevent the DNA strand from extending, FAM was a fluorophore. Exonuclease NFO (buffer) recognized dSpacer (tetrahydrofuran, THF) and cut it to form a free hydroxyl end extending at the 3′ end. Finally, the amplification product of 5′ FAM-RPA-3′ Biotin was obtained. Gold particle of biotin antibody and FAM antibody fixed on LFD (T line), 5′ FAM-RPA-3′ biotin will be captured to appear bands.
The labeled probe will be as follows:
5′ 6-FAM-AGGCTCCCTCCTCGGTAACGAAGTCCGCTCCA/idSp/GTTTGACAGCACATT-3′ C3 Spacer.
Figure 1. Screenshot of Snapgene with Fusarium sp. genome sequence comparison. Specific primers were designed at locations with large sequence differences. The red boxes indicated the locations of the primers and probes.
DNA extraction
Culture the F. tricinctum, F. proliferatum, F. fujikuroi, F. graminearum, F. solani, F. equiseti, C. siamense, C. gloeosporioides, C. aenigma, C. fructicola, Botrytis cinerea, Phoma sp., and Stagonosporopsis sp. isolates, stored in a 4 refrigerator, in a PDA plate using an inoculating needle. Culture at 28 until colony diameter reaches 6 cm.
Scrape the mycelium into a 1.5 mL tube, add 0.1 g of quartz sand and 300 μL of CTAB extracting solution, and grind for 120 s at 85 Hz in a grinding machine.
Add 400 μL of CTAB extracting solution and shake for 15 s using the vortex.
Add 700 μL of trichloromethane, shake for 15 s using the vortex, and centrifuge at 13,800× g for 10 min.
Remove 500 μL of supernatant and transfer it to a new 1.5 mL tube. Add 500 mL of isopropanol and centrifuge at 13,800× g for 10 min.
Discard the supernatant, add 500 μL of 70% ethanol, and centrifuge at 13,800× g for 5 min.
Discard the supernatant, invert the tube for 2 h to dry, and add 50 μL of ddH2O. Store at 4°C.
Specific detection of primers and probes
Set up the reaction system as follows:
Buffer A 29.4 μL
10 μM upstream F91 2 μL
10 μM downstream R91 2 μL
10 μM probe 0.6 μL
ddH2O 11.5 μL
Template DNA 2 μL
Buffer B 2.5 μL
Buffer B needs to be added last. Buffer A and Buffer B are supplied with the HybriD DNA thermostat rapid amplification kit.
Incubate at 38 °C for 10 min.
Dilute the reaction product 20 times using ddH2O and add 50 μL of the diluted reaction product to the sample hole of HybriDetect LFD. Interpretation of test results is available within 5 min. If the test results are not read within 5 min, the test is not successful.
If the control line is visible and the detection line is not, this means a negative result. If both the detection and control lines are visible, this means a positive result. If neither the detection and control lines are visible, this means that the experimental operation was performed incorrectly or that the kit was damaged (Figure 2).
Figure 2. Assessment of specificity of primers pair F91/R91 and probe for recombinase polymerase amplification–lateral flow dipstick (RPA–LFD) detection of F. oxysporum
RPA–LFD reaction condition optimization
According to kit instructions, set the reaction temperature to 33, 36, 39, 42, and 45 °C and the reaction time to 10 min.
After determining the optimal reaction temperature, set the reaction time to 4, 6, 8, 10, and 12 min.
Visualize the amplification results according to the above method.
The best reaction condition was determined according to the color depth of the detection line (Figure 3).
Figure 3. Optimization of recombinase polymerase amplification–lateral flow dipstick (RPA–LFD) detection for F. oxysporum. (A) Optimization of the RPA reaction temperature (33, 36, 39, 42, 45, 48, and 51 °C). (B) Optimization of the RPA reaction time (4, 6, 8, 10, 12, and 14 min).
Detection system sensitivity verification
Set the DNA concentration to 10 ng/μL, 1 ng/mL, 100 pg/μL, 10 pg/μL, 1 pg/μL, and 100 fg/μL. DNA concentration was determined using Ultramicro accounting protein analyzer.
Add different concentrations of DNA templates to the reaction system.
Incubate at 39 °C for 8 min.
Visualize the amplification results according to step C3.
The detection limit was determined by the presence or absence of the detection line.
Field sample detection
Take 100 mg of strawberry crown tissue (approximately 125 mm3 of crown tissue) and place in a 1.5 mL tube.
Add 100 μL of 60% PEG200 .
Incubate at room temperature for 15 min to lyse plant tissue and release DNA.
Add the lysate supernatant as a template into the RPA reaction system. Set up the reaction system as follows:
Buffer A 29.4 μL
Upstream F91 2 μL
Downstream R91 2 μL
Probe 0.6 μL
ddH2O 11.5 μL
Template DNA 2 μL
Buffer B 2.5 μL
Buffer B needs to be added last. Buffer A and Buffer B are supplied with the kit.
Incubate at 39 °C for 8 min.
Visualize the amplification results according to step C3.
If the control line is visible and the detection line is not, this indicates that the strawberry was not infected by F. oxysporum. If both the detection and control lines are visible, this indicates that the strawberry was infected by F. oxysporum (Figure 4).
Figure 4. Symptoms of strawberry crown rot and the result of recombinase polymerase amplification–lateral flow dipstick (RPA–LFD) detection. (a–a2) Plant with crown rot symptoms and positive test results. (b–b2) Plant without crown rot symptoms and positive test results. (c–c2) Plant without crown rot symptoms and negative test results. The tissue indicated by the arrow is used for detection.
Data analysis
Determination of primers and probes specificity:
Judge the specificity of primers and probes by the presence or absence of detection lines of HybriDetect LFD. The detection line was visible only for F. oxysporum and the detected result is positive; the detection lines were not visible for other genomic DNA, and the detected result is negative (Table 1). These results indicate that the primers F91/R91 and probe had good specificity for F. oxysporum.
Table 1. Results of other isolates detected by RPA-LFD
Species Detection result
F. oxysporum +
F. tricinctum -
F. proliferatum -
F. fujikuroi -
F. graminearum -
F. solani -
F. equiseti -
C. siamense -
C. gloeosporioides -
C. aenigma -
C. fructicola -
Botrytis cinerea -
Phoma sp. -
Stagonosporopsis sp. -
“+” represents positive result, “-” represents negative result.
Determination of optimal reaction conditions:
According to the manufacturer’s instructions, the optimal reaction temperature was determined with 10 min as the reaction time. The result showed that 39 °C is the optimal reaction temperature, and the optimal reaction time was 8 min.
Determination of detection limits:
The detection limit was determined by the presence or absence of the detection line. The HybriDetect LFD still had bands in the detection line at a template concentration of 1 pg/μL, and we concluded that the detection limit of the RPA-LFD detection was 1 pg/μL.
Field application of RPA-LFD detection:
The appearance of the test line indicated a positive result, suggesting that the plant was infected with F. oxysporum.
Validation of protocol
Hu et al. (2023) [7]. Establishment of the recombinase polymerase amplification-lateral flow dipstick (RPA-LFD) detection technique for Fusarium oxysporum. Plant Disease. (Figure 4).
General notes and troubleshooting
General notes
The length of primers for RPA detection was 31–32 bp; too-short primers will affect the amplification speed and detection sensitivity.
Before the positive test, the negative test was performed to rule out the formation of false positives due to the primers probe.
Care should be taken to avoid cross contamination during the experimental operation.
This operation was effective only when the HybriDetect LFD control line is striped.
HybriDetect LFD interpretation results were completed within 5 min.
Troubleshooting
Problem 1: The combination of primers and probe could not specifically detect the target fungus.
Possible cause(s): Poor specificity of primers and probe.
Solution(s): The specific primers need to be screened again, and the probes need to be screened after the specific primers are obtained.
Problem 2: The test line of negative control showed banding.
Possible cause(s): The primers bound to the probe itself.
Solution(s): Re-screen primers and probe.
Problem 3: The color of the detection line was weak.
Possible cause(s): The reaction conditions were not optimal, or the DNA concentration is low.
Solution(s): Optimize reaction conditions and increase DNA concentration.
Acknowledgments
This research was funded by Agriculture and Social Development Research Project of Hangzhou (202203A07), Guizhou Research academy [110202101048(LS-08)], and Key Research and Development Project of Zhejiang Province, China (No. 2020C02005).
The protocols described here are adapted from our previous work [7].
Competing interests
The authors declare no competing interests.
References
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4,916 | https://bio-protocol.org/en/bpdetail?id=4916&type=0 | # Bio-Protocol Content
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Generation of Mature Toxoplasma gondii Bradyzoites in Human Immortalized Myogenic KD3 Cells
DM Deborah Maus
BC Blake Curtis
DW David Warschkau
EB Estefanía Delgado Betancourt
FS Frank Seeber
MB Martin Blume
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4916 Views: 535
Reviewed by: Marcelo S. da Silva Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Communications Mar 2022
Abstract
Toxoplasma gondii is a zoonotic protozoan parasite and one of the most successful foodborne pathogens. Upon infection and dissemination, the parasites convert into the persisting, chronic form called bradyzoites, which reside within cysts in muscle and brain tissue. Despite their importance, bradyzoites remain difficult to investigate directly, owing to limited in vitro models. In addition, the need for new drugs targeting the chronic stage, which is underlined by the lack of eradicating treatment options, remains difficult to address since in vitro access to drug-tolerant bradyzoites remains limited. We recently published the use of a human myotube-based bradyzoite cell culture system and demonstrated its applicability to investigate the biology of T. gondii bradyzoites. Encysted parasites can be functionally matured during long-term cultivation in these immortalized cells and possess many in vivo–like features, including pepsin resistance, oral infectivity, and antifolate resistance. In addition, the system is scalable, enabling experimental approaches that rely on large numbers, such as metabolomics. In short, we detail the cultivation of terminally differentiated human myotubes and their subsequent infection with tachyzoites, which then mature to encysted bradyzoites within four weeks at ambient CO2 levels. We also discuss critical aspects of the procedure and suggest improvements.
Key features
• This protocol describes a scalable human myotube-based in vitro system capable of generating encysted bradyzoites featuring in vivo hallmarks.
• Bradyzoite differentiation is facilitated through CO2 depletion but without additional artificial stress factors like alkaline pH.
• Functional maturation occurs over four weeks.
Graphical overview
Keywords: Toxoplasma gondii Tissue cysts Bradyzoites Cell culture Human myotubes
Background
The obligate intracellular parasite Toxoplasma gondii infects nearly all warm-blooded animals and an estimated quarter of the human global population (Molan et al., 2019). The World Health Organization ranked T. gondii to be an important and common food-borne protozoan, with one of the highest disease burdens worldwide (Torgerson et al., 2015). It commonly manifests as an opportunistic pathogen in AIDS patients and other immunocompromised individuals (Luft and Remington, 1992). In contrast, most healthy individuals can effectively control the acute infection through their immune system (Denkers and Gazzinelli, 1998). Semi-dormant bradyzoites can persist for the duration of the host’s life in cysts within mainly the brain or muscle tissue, being resistant to current treatment options (Dubey, et al., 1998; Watts et al., 2015; Barrett et al., 2019). Bradyzoites can reactivate if the immune pressure abates (e.g., via AIDS, transplantation, etc.) leading to the emergence of severe clinical manifestations. Tissue cysts are a major source of transmission via the consumption of undercooked meat from infected animals (Gabriël et al., 2022). Hence, the investigation of bradyzoite biology including chemotherapy development is much needed.
In the past, corresponding research efforts were hindered by the lack of an appropriate in vitro model. Either isolated cysts from mice brain homogenates or functionally immature bradyzoites derived from fibroblast cell cultures were used. Many traits of bradyzoites, such as drug-tolerance mechanisms, remain challenging to study in vitro. Spontaneous conversion of tachyzoites to bradyzoites has been characterized in murine C2C12 myotubes (Guimarães et al., 2008; Ferreira-da-Silva et al., 2009; Swierzy and Lüder, 2015). We recently showed that immortalized human myoblasts (called KD3) are also suitable for long-term bradyzoite culture (Christiansen et al., 2022). KD3 myoblasts harbor transgenes for mutated cyclin-dependent kinase 4, cyclin D1, and a human telomerase reverse transcriptase (hTERT), resulting in the immortalization of these primary subcutaneous muscle cells (Shiomi et al., 2011). In differentiated KD3 myotubes, cystogenic strains of T. gondii can be matured over a period of four weeks, during which the cysts develop structural and functional traits of in vivo cysts (Christiansen et al., 2022) and express bradyzoite and cyst markers, such as BAG1, CC2, glycans, a discernible cyst wall, as well as amylopectin granules. In addition, the cysts become tolerant to the antifolates pyrimethamine and sulfadiazine and are orally infectious to mice. They gradually develop resistance to pepsin digestion and survive storage at 4 °C and exposure to 55 °C (Christiansen et al., 2022).
With this protocol, we aim to facilitate the use of in vitro T. gondii cysts in studies to reduce animal experiments and to increase the exploration of this comparatively understudied form of T. gondii.
Materials and reagents
Cell lines
BJ-5ta human foreskin fibroblast (HFF) cells (ATCC CRL-4001)
Toxoplasma gondii Prugniaud- tdTomato (Chtanova et al., 2008) or any other cystogenic strain
KD3 myoblasts (Shiomi et al., 2011) or other immortalized human myogenic cell lines with similar properties
Note: We know of four human immortalized myogenic cell lines (and derivatives thereof) described in the literature (Zhu et al., 2007; Shiomi et al., 2011; Thorley et al., 2016; Massenet et al., 2020). KD3 cells used here were initially obtained from N. Hashimoto. Inquiries for KD3 cells should now be addressed to Dr. Tohru Hosoyama, Dept. of Musculoskeletal Disease, The Geroscience Research Center, National Center for Geriatrics and Gerontology, 7-430 Morioka-cho, Obu City, Aichi Prefecture, Japan ([email protected]). Upon completion of an MTA, the cells might be obtained in Europe from the authors, or in the US from Dr. L. Weiss, Albert Einstein College of Medicine, 1300 Morris Park Avenue, Forchheimer Building, Bronx, NY 10461, USA ([email protected]). LHCN-M2 cells (Zhu et al., 2007) are commercially available from Evercyte. Cells from Massenet et al. might be obtained from the respective authors.
Reagents
Dulbecco’s modified Eagle’s medium (DMEM) (Gibco Life Technologies, catalog number: 12800-082)
Glucose (Carl-Roth, catalog number: X997.2)
L-glutamine (Thermo Fisher Scientific, catalog number: 25030081)
Sodium pyruvate (Capricorn Scientific, catalog number: NPY-B)
Penicillin/streptomycin (Capricorn Scientific, catalog number: PS-B)
Bovine serum, heat inactivated, iron fortified (Capricorn Scientific, catalog number: CS-IF-1A)
Fetal bovine serum, heat inactivated (Capricorn Scientific, catalog number: FBS-12A)
Fetal bovine serum, heat inactivated low endotoxin (Capricorn Scientific, catalog number: FBS-LE-12A)
Ultroser® G (Cytogen, catalog number: 15950-017)
Donor horse serum (Capricorn Scientific, catalog number: DHS-1A)
Human recombinant insulin (PAN-Biotech, catalog number: P-2701001)
Insulin Lispro (Sanofi, catalog number: 12910612)
Human holo-transferrin (Sigma, catalog number: TO665)
Na2SeO3 (Sigma-Aldrich, catalog number: 214485)
RPMI 1640 pH 7.4 (containing 4 mM L-glutamine) (Sigma-Aldrich, catalog number: R1383)
HEPES (Sigma-Aldrich, catalog number: H3375)
DMSO (AppliChem, catalog number: A3672)
Collagen I from rat tail (Corning, catalog number: 354236)
NaCl (Merck catalog number: 1.06404)
KCl (Merck, catalog number: 1.04936)
KH2PO4 (Merck, catalog number: 1.04877)
Na2HPO4·2H2O (Merck, catalog number: 1.06580)
Acetic acid (Carl Roth, catalog number: 3738.4)
0.05% trypsin/0.53 mM EDTA (Capricorn Scientific, catalog number: TRY-1B10)
Media and solutions
70% ethanol for sterilization
HFF growth medium (see Recipes)
Tachyzoite medium (see Recipes)
Myoblast growth medium (see Recipes)
Myoblast differentiation medium (see Recipes)
Cyst medium (see Recipes)
Freezing medium (see Recipes)
Rat tail collagen type I (see Recipes)
1× Trypsin-EDTA solution in PBS (w/o Ca2+ and Mg2+)
Phosphate-buffered saline (PBS) (w/o Ca2+and Mg2+)
Note: All media have to be sterile.
Material
T-25 or T-75 cell culture flasks with filter caps (TPP, catalog number: 90026 or 90076)
Microtiter plates 6 well or 96 well (TPP, catalog number: 92006 or 92048)
15 mL and 50 mL conical centrifuge tubes (TPP, catalog number: 91015 and 91050)
1.5 mL cryovials (Nalgene, catalog number: 5000-1020)
Pipette tips [10, 200, 1,000 µL (Various brands)]
Sterile syringe filters (0.22 μm) (TPP, catalog number: 99722)
C-Chip disposable hemocytometer (NanoEnTek Inc., DHC-N01) or Neubauer hemocytometer
Syringes (20 mL) (Luer Lock, Injekt, catalog number: 4606736V)
Blunt cannula (27 G) (Sterican, catalog number: 9180117)
Note: Examples for providers/manufacturers are given; similar material from others will probably work as well.
Recipes
HFF growth medium
DMEM pH 7.2 (containing 3.75 g/L NaHCO3, 25 mM glucose, 4 mM L-glutamine, and 1 mM sodium pyruvate), supplemented with 1× penicillin/streptomycin and 10% bovine serum. Store at 4 °C.
Tachyzoite medium
HFF growth medium but with only 1% fetal bovine serum. Store at 4 °C.
Myoblast growth medium
DMEM pH 7.2 (containing 3.75 g/L NaHCO3, 25 mM glucose, 4 mM L-glutamine, and 1 mM sodium pyruvate), supplemented with 1× penicillin/streptomycin, 20% fetal bovine serum, and 2% Ultroser® G. Store at 4 °C.
Myoblast differentiation medium
DMEM pH 7.2 (containing 3.75 g/L NaHCO3, 25 mM glucose, 4 mM L-glutamine, and 1 mM sodium pyruvate), supplemented with 1× penicillin/streptomycin, 2% donor horse serum, 10 µg/mL human recombinant insulin (alternatively, Insulin Lispro), 5 µg/mL human holo-transferrin, and 10 nM Na2SeO3. Store at 4 °C.
Note: Lower concentrations of insulin might also work, according to literature.
Cyst medium
RPMI 1640 pH 7.4 (containing 4 mM L-glutamine), supplemented with 5 mM glucose, 1× penicillin/streptomycin, 50 mM HEPES, 2% donor horse serum, 10 µg/mL human recombinant insulin (alternatively, Insulin Lispro), 5 µg/mL human holo-transferrin, and 10 nM Na2SeO3. Store at 4 °C.
Freezing medium
90% heat-inactivated fetal bovine serum low endotoxin and 10% DMSO. Store at 4 °C.
Rat tail collagen
Collagen I from rat tail dissolved at a stock concentration of 0.1 mg/mL in 100 mM acetic acid. Use 1:6.67 in water. Store at 4 °C.
1× Trypsin-EDTA solution
0.05% trypsin/0.53 mM EDTA in PBS (w/o Ca2+ and Mg2+)
Phosphate-buffered saline (PBS) without calcium and magnesium
MilliQ water supplemented with 137 mM NaCl, 2.7 mM KCl, 1.5 mM KH2PO4, and 6.5 mM Na2HPO4·2H2O at pH 7.4.
Equipment
Humidified CO2 cell incubator at 36.7 °C, 10% CO2 (Thermo Fisher Scientific, model: Heracell 150i CO2 incubator)
Incubator at 36.7 °C, ambient CO2 (Binder Inkubator, model: KB 240)
Safety cabinet for BSL-2 work (Luft- und Reinraumtechnik GmbH, model: BDK S1200 laminar flow workbench)
Water bath at 37 °C (Memmert GmbH & Co. KG, model: WNB 14)
Adjustable micropipettes 2–20 µL, 20–200 µL, 100–1,000 µL (Eppendorf GmbH, model: Research Plus micropipette)
Pipetting aid for serological pipettes (Brand GmbH, model: Accu-Jet Pro pipette)
Vacuum aspirator system, Integra Biosciences Vacusafe Comfort Aspiration system (Integra, model: 10590273)
Inverted light microscope with phase contrast, differential interference contrast, or similar optics (Nikon, model: Eclipse Ts2 inverted microscope)
Centrifuge with swing-out rotor and refrigeration (Hettich GmbH & Co. KG, model: Rotina 380R centrifuge 1706)
CoolCell LX Cell Freezing Vial Containers (Fisher Scientific, Corning, catalog number: 15572771) or similar cryo-freezing container
Refrigerator 4 °C (AEG, model: RTB415E2AW)
-20 °C freezer (Liebherr, model: Mediline LGex 3410)
-70 °C freezer (SANYO, model: VIP Series MDF-U53V ultra-low temperature freezer)
Liquid nitrogen storage capabilities
Note: Examples for providers/manufacturers are given; similar material from others will probably work as well.
Procedure
General description of the procedure and notes
Generating mature T. gondii cysts in KD3 human myotubes involves three steps (Figure 1): (i) set up a KD3 myoblast culture, (ii) differentiation of these cells into KD3 myotubes, and (iii) infection of myotubes with tachyzoites and the differentiation of parasites to bradyzoites. We also suggest experiments to confirm the maturation of cysts.
Figure 1. Overview of the protocol steps. Timing of the cultivation of in vitro T. gondii cysts. KD3 myoblasts are seeded into desired cell culture vessels. When they reach a maximum confluency of 50%, differentiation is induced. After five days, the cells feature longer tube-like multinucleated cells, which are hallmarks of the myotube stage. Myotubes are infected with cystogenic T. gondii parasites. Over the course of four weeks of CO2 depletion, tachyzoites differentiate into bradyzoites and mature to in vitro cysts. (B) Representative pictures of the same culture at relevant developmental stages. Prugniaud parasites express the red fluorescent protein tdTomato. Scale bar indicates 100 µm.
(i) Myoblast culture
Handling of KD3 myoblasts is adapted from the procedure described by Shiomi et al. (2011). KD3 myoblasts appear similar to endothelial cells (Figure 2A) and are slightly heat-sensitive (avoid prolonged temperatures higher than 37 °C during cultivation). During maintenance of the myoblasts, it is imperative to keep the main culture subconfluent (<70%), as cell-to-cell contact appears to act as a trigger for differentiation into myotubes. Before the monolayer reaches this density, the cells are subcultured (split). The medium is exchanged for warm phosphate-buffered saline without magnesium and calcium. Pre-warmed trypsin-EDTA solution is added to cover the monolayer and the culture is incubated at 36.7 °C until cells detach. The trypsinization reaction is stopped by adding myoblast growth medium, and a homogenous cell suspension is generated by pipetting. At this point, the cell density can be determined by counting. The suspension is distributed to suitable cell culture vessels, which can optionally be coated with collagen to enhance myotube attachment. Do not aim for a confluency below 5%, as this suppresses myoblast growth. Incubate cultures at 36.7 °C at 5%–10% CO2.
Since cell-to-cell contact cannot be prevented even at low confluency subculturing, KD3 myoblasts may unintentionally start to differentiate to myotubes, and the culture may exhibit a mixture of both cell types (Figure 2C). We often observe this with increased confluency; the growth rate drastically declines, and the culture becomes unsuitable for further use. The number of usable passages of the culture thus highly depends on handling, and we recommend monitoring cell behavior in correlation with the respective passage number. Generally, the number of passages should not exceed 16 of a freshly thawed vial. An occasional selection for the presence of the transgenes might be advisable. The cells are resistant to G418 (400 µg/mL) and puromycin (0.5 µg/mL) (Shiomi et al., 2011).
Long-term KD3 myoblast storage is done in liquid nitrogen in freezing medium. Use a controlled freezing rate apparatus or an isopropanol chamber to ensure a slow and steady temperature drop. For thawing, quickly thaw in a 37 °C water bath, replace the freezing medium with growth medium, and distribute cells to a cell culture vessel. Incubate at 36.7 °C at 5%–10% CO2 until the culture is ready for subcultivation. We usually use 0.3 mL of medium per cm2 cell culture vessel.
Figure 2. KD3 culture at various stages. (A) Healthy KD3 myoblast culture at the right confluency for myotube induction. Scale bar indicates 500 µm. (B) KD3 myoblast cultures at the maximum density allowed before splitting. (C) Improper handling or high passage number of the KD3 culture creates a mixture of myoblasts and emerging myotubes. Scale bar indicates 100 µm. (D) KD3 myotubes five days post induction. Scale bar indicates 100 µm.
(ii) Myoblast differentiation
KD3 myoblasts are progenitor cells that undergo myogenesis when induced by serum starvation (Shiomi et al., 2011). Growth medium is replaced with myoblast differentiation medium at a confluency of 20%–50% (Figure 2A). Cells fuse and form multinucleated tubes over the course of five days. Often, myoblasts replicate once or twice before they fuse (Figure 2B). The differentiation efficiency depends on the general potency and passage number of the myoblast culture.
(iii) Tachyzoite infection and bradyzoite maturation
In general, every cystogenic strain of T. gondii can be used for infection, and a selection of type I, type II, and type III strains were tested directly for their ability to generate DBA-positive and SAG1-negative cysts (Christiansen et al., 2022). For a fully matured, in vivo–like cyst culture, we recommend the use of type II or type III strains. We frequently cultivate a Pru-∆hxgprt tdTomato (Chtanova et al., 2008) strain. Furthermore, we recommend using a fluorescent protein–expressing strain to easily monitor the cyst development, since the cysts may be difficult to find using brightfield or phase-contrast microscopy. It is also highly beneficial to use a tachyzoite strain that has been recently re-differentiated from either a cyst-containing tissue homogenate (Dubey, 1998) or a previous in vitro cyst culture, as long-term tachyzoite cultures lead to low differentiation efficiencies (Colos-Arango et al., 2023).
The myotube culture is infected with 7.2 × 103 tachyzoites per cm2 surface area in cyst medium. The culture is incubated at 36.7 °C and ambient CO2 concentration. Ambient CO2 levels limit the rate of parasite pyrimidine biosynthesis and facilitate stage conversion (Bohne and Roos, 1997). After infection, the culture needs to be monitored daily and the medium is replaced every second to third day. Parasite stage-conversion depends on the quality of the myotubes and the cystogenicity of the strain and will start immediately post-infection. However, long-term tachyzoite cultures with a decreased cystogenic potential may require additional washing steps if parasite egress is apparent to prevent overgrowth of the culture with tachyzoites. After four weeks, the cysts are largely functionally mature (Christiansen et al., 2022).
Throughout the experiment, myotubes move and change their shape, which can lead to cell detachment. Furthermore, throughout the culture period, cysts grow in size, new cysts will emerge from egressed bradyzoites, and cysts may also dissipate.
Bench protocol
All parasite and muscle cell handling should be done in a sterile laminar flow BSL-2 workbench.
Thawing KD3 myoblasts
Warm myoblast growth medium to 37 °C in a water bath.
Sterilize all involved surfaces of the laminar flow BSL-2 workbench with 70% ethanol.
Place cryovial containing frozen KD3 myoblasts on ice.
In the laminar flow BSL-2 workbench, fill a 15 mL centrifuge tube with warm myoblast growth medium.
Place the cryovial in the 37 °C warm water bath until the outer layer of the frozen cell suspension starts to melt.
Sterilize the cryovial with 70% ethanol.
In the laminar flow BSL-2 workbench, transfer the cell suspension from the cryovial to the centrifugation tube.
Centrifuge the suspension at 500× g for 5 min at room temperature.
In the laminar flow BSL-2 workbench, open the centrifuge tube and discard the supernatant.
Gently resuspend the pellet in 15 mL of myoblast growth medium. Transfer the suspension to a T-75 cell culture flask with a filter cap (choose flask size depending on cell density).
Incubate in a humidified incubator at 5%–10% CO2 at 36.7 °C.
The next day, visually inspect confluency and cell adhesion under a microscope.
Subculturing KD3 myoblasts
Critical: Before the monolayer reaches a confluency of 70%, subculture the myoblasts.
Warm PBS (without Ca2+ and Mg2+) and 0.05% trypsin/EDTA in PBS to room temperature.
Warm myoblast growth medium in a water bath to 37 °C.
Sterilize all involved surfaces of the laminar flow BSL-2 workbench with 70% ethanol.
Aspirate the culture medium.
Wash the monolayer with at least 10 mL of pre-warmed PBS (without Ca2+ and Mg2+).
Add 0.02 mL/cm2 trypsin/EDTA in PBS to the monolayer.
Incubate at 36.7 °C and visually inspect for the detachment of cells (approximately 2 min).
Add sufficient myoblast growth medium to neutralize trypsin activity and mix the suspension by pipetting. Transfer the suspension into a centrifuge tube and centrifuge at 500× g for 5 min at room temperature.
Discard the trypsin-containing supernatant and add myoblast growth medium (0.3 mL per cm2 culture area).
Optional: Use a hemocytometer to determine the cell density and distribute 6 × 103 cells per cm2 culture area into desired cell culture vessels after the trypsinized cells are spun down and resuspended in fresh growth medium.
Incubate in a humidified incubator at 5%–10% CO2 at 36.7 °C. Typically, 70% confluency is reached after two to three days.
Freezing KD3 myoblasts
Critical: Use a culture with a low passage number for cryopreservation.
Critical: Freeze myoblasts before the monolayer reaches a confluency of 70%.
Equilibrate PBS (without Ca2+ and Mg2+) and trypsin/EDTA in PBS to room temperature.
Warm freezing medium in a water bath to 37 °C.
Sterilize all surfaces involved with 70% ethanol.
Discard the medium.
Wash the monolayer with at least 10 mL of PBS (without Ca2+ and Mg2+).
Add 0.02 mL/cm2 trypsin/EDTA in PBS to the monolayer.
Incubate at 37 °C and visually inspect for the detachment of cells (approximately 2 min).
Add freezing medium to stop trypsin digestion and gently mix the suspension by pipetting.
Transfer the suspension into a centrifuge tube and centrifuge the cell suspension at 500× g for 5 min at room temperature. Discard the trypsin-containing supernatant and add myoblast growth medium.
Aliquot 75 cm2 worth of cells per cryovial (approximately 1.5–1.8 × 106 cells).
Place cryovials in a cryo-freezing container and freeze them overnight at -70 °C.
Store cryovials in liquid nitrogen.
Differentiating KD3 myoblasts into myotubes
Optional: Coat the desired cell culture vessel with 15 μg/mL rat tail collagen type I for 1 h at room temperature. Wash with PBS three times or leave until completely dry in the laminar flow BSL-2 workbench. Coated plasticware may be stored at 4 °C for a few weeks.
Follow the instructions of Section B, steps 1–10.
Plate 6 × 103 cells per cm2 culture area into the desired culture vessel containing myoblast growth medium.
Incubate in a humidified incubator at 5%–10% CO2 at 36.7 °C.
Critical: Visually inspect confluency. Begin differentiation at 20%–50% monolayer confluency (usually after one or two days).
Warm myoblast differentiation medium to 37 °C in a water bath.
Sterilize all surfaces involved with 70% ethanol.
Aspirate growth medium. Make sure to completely remove the myoblast growth medium by washing the cells with warm PBS at least once.
Add myoblast differentiation medium.
Incubate in a humidified incubator at 5%–10% CO2 at 36.7 °C.
After five days, the cells have fused and formed multinucleated myotubes.
Optional: Confirm cell fusion by appropriate staining protocols (e.g., immunofluorescence staining for myosin heavy chain and nuclei) and calculate myogenic index (Noë et al., 2022).
Infection of KD3 myotubes and cyst maturation
Tachyzoites are maintained in HFF host cells using standard procedures (Khan and Grigg, 2017).
Determine the parasite count in a freshly egressed or syringe-released tachyzoite suspension.
The myotubes are infected with 7.2 × 103 tachyzoites per cm2. Prepare an appropriate dilution of the parasite suspension in cyst medium.
Wash the myotube monolayer with cyst medium once and then apply the parasite suspension.
Critical: Incubate in an incubator at 36.7 °C at ambient CO2 concentration.
Change cyst medium gently every second to third day. Leave 25% of the volume in the cell culture vessel and slowly replace only 75% to avoid perturbation of the monolayer.
Critical: Visually monitor the culture using an inverted microscope. Inspect for possible tachyzoite overgrowth, myotube delamination, or cyst loss (see troubleshooting section).
After four weeks, the culture contains matured cysts.
Optional: To induce tachyzoite re-differentiation, replace cyst medium with tachyzoite medium and incubate in a humidified incubator at 5%–10% CO2 at 36.7 °C.
Validation of protocol
Several markers are known to indicate bradyzoite development, including the presence of proteins BAG1, LDH2, and p21, and the absence of the SAG1 surface protein. The formation of the cyst wall may be monitored using antibodies against CST1 and CC2 or Dolichos biflorus agglutinin (DBA) staining (Yang and Parmley, 1995; Gross et al., 1996; Dubey et al., 1998; Knoll and Boothroyd, 1998; Zhang et al., 2001). All these markers start to appear within the first week of cyst maturation and have been shown to develop even using fibroblast-based methods. In contrast, functional cyst maturation requires more time. Resistance to pepsin, temperature stress, and drugs, as well as oral infectivity gradually emerge after 14 days and continue to develop until at least 28 days (Christiansen et al., 2022). Drug resistance can be easily monitored by exposing cysts for one week to high doses of pyrimethamine (20 µM) and sulfadiazine (20 µM) and monitoring re-differentiation.
General notes and troubleshooting
Application of the protocol
The method can be helpful in many applications, considering the different characteristics to be studied and the scale required for their analysis. While the cyst wall starts to develop within the first week of differentiation as seen by DBA staining (Christiansen et al., 2022), antifolate tolerance fully developed only after four weeks. Further, resistance to pepsin digestion is increasing throughout four weeks of maturation. Myotubes are a natural host cell type for T. gondii and thus may provide a more relevant system to investigate parasite interactions with the host cell than, for instance, HFF.
Downstream applications require suitable cyst isolation protocols. We performed an LC-MS-driven metabolic characterization of in vitro bradyzoites. To this end, we isolated tissue cysts using DBA- and streptavidin-coated magnetic beads from syringe-passaged infected KD3 myotubes (Christiansen et al., 2022). This isolation procedure can be performed at 4 °C and does not require detergents, polymers, or other MS-incompatible chemicals.
For other applications such as imaging, intact cysts can also be isolated using a percoll gradient (Watts et al., 2017). Single bradyzoites have been obtained using either pepsin (Dubey, 1998) or trypsin digestion (Jacobs et al., 1960; Fu et al., 2021). Enzymatic treatment may be directly applied to the monolayer and may provide a means of removing immature bradyzoites before purification.
Given the need for new drugs and drug targets, research efforts in the field of drug discovery have intensified over the last years (Dittmar et al., 2016; Adeyemi et al., 2018; Spalenka et al., 2018; Han et al., 2020). However, none of these studies integrated matured cysts into the screening process. Our method can be adapted into a 96-well plate format suitable for drug screens. Compounds that are cidal to bradyzoites would prevent regrowth of re-differentiated tachyzoites, which are easily detectable by fluorescence or plaque assay. In vitro bradyzoites develop tolerance towards antifolates and other antiparasitics in a time-dependent manner (Christiansen et al., 2022).
In the last 10 years, more than 135 studies in PubMed have mentioned the use of mice in the context of T. gondii bradyzoites. Assuming an average of 100 mice per study and an unknown number of mice that are routinely infected to have a constant supply of tissue cysts, this adds up to several thousand animals. A significant number of those could have been replaced if adequate in vitro systems like this one would have been applied. We hope that this protocol encourages the community to consider its usefulness instead of mouse experiments, where appropriate, thereby contributing to the 3R principle.
Limitations and further development of the procedure
Improving the myotube culture
Terminally differentiated myotubes are a suitable cell type for obtaining mature tissue cysts since they are long-lived under in vitro conditions. However, occasionally we observe the loss and morphological changes of myotubes, which may have a negative impact on cyst yield. Infected and detached myotubes might be rinsed off with media change. This problem may be addressed by improving host cell attachment to the culture vessel using various methods (Madden et al., 2015; Bettadapur et al., 2016; Denes et al., 2019; Martins et al., 2022). Finally, cellular senescence may also pose a limit on the maximal passages of KD3 cells and may be ameliorated by reversine treatment (Rajabian et al., 2023). Mild heat stress from temperature fluctuations of cell culture incubators may also induce differentiation and needs to be avoided (Yamaguchi et al., 2010). Whether any of these strategies provide improvements for in vitro cyst generation needs to be tested.
Improving the cyst culture
A healthy, long-lasting myotube culture is required but not sufficient for in vitro cyst maturation. In particular, parasite strains that have been passaged over extended periods of time as tachyzoites can easily overgrow in the first week of infection (Colos-Arango et al., 2023). Adjusting the multiplicity of infection can ameliorate this problem. Ideally, parasites intended for in vitro bradyzoite generation that have been recently isolated from an infected mouse brain or a previous long-term bradyzoite culture should be used.
Over the course of a four-week culture, the medium is changed 9–12 times, and associated shearing forces may lead to the loss of some cysts. This can be prevented by careful handling and minimizing medium changes in general. We have not experimented with chemostat culture or automated pipetting systems (Pazdzior and Kubicek, 2021). They may provide a more stable and predictable environment during the whole culturing period and could reduce workforce for large-scale approaches.
Cyst maturity can be assessed by a variety of assays relevant to the application. To date, there is a lack of very late reporter strains that allow monitoring the expression of late (>2 weeks) bradyzoite markers. However, bradyzoites within tissue cysts are considered to form a heterogeneous population and may thus display different proteins (Watts et al., 2015). The reported proteomic datasets from up to 5-month-old bradyzoites from infected mouse brains could provide a starting point for the construction of such reporter strains (Garfoot et al., 2019).
Reduction of animal ingredients and cost-effectiveness
To satisfy 3R principles and reduce costs, it is preferable to reduce both the use of animal experiments and the use of animal-derived products in cell culture. In particular, serum components in media formulations used in this protocol may be replaced or reduced. Serum-free medium maintains murine C2C12 cells in both their myoblasts and myotube stage, respectively (Jang et al., 2022). In particular, myotube formation can also be induced in serum-free, BSA-supplemented media for human skeletal muscle cell isolates (Rajabian et al., 2021). The impact of these serum-reduction strategies on cyst maturation and their resemblance to in vivo cysts, however, remains to be established.
Troubleshooting
KD3 myoblasts do not completely differentiate into myotubes
KD3 myoblasts may be used up to 16 passages upon thawing a new vial. However, we observed that with increased passage numbers their potential to differentiate into myotubes decreases and myotube formation may take longer. Also, myotube formation is effectively induced by serum starvation, and high-serum myoblast growth medium has to be removed completely. Myoblasts exhibit the potential to differentiate into other cell types besides myotubes such as adipocytes or osteocytes depending on the media composition (Shiomi et al., 2011). If the differentiation of myotubes was only partially successful, and, for example, a subsequent treatment of the cyst culture involves fatty acids, uncommitted myoblasts will differentiate to adipocytes.
KD3 myotube monolayer has gaps
Delamination of myotube monolayers can occur; however, not to an obstructive extent. Clusters of detaching myotubes may leave empty patches and incompletely differentiated myotubes behind. We found this phenomenon to be more prevalent when myoblasts were grown too dense prior to myogenesis induction. Seed fewer myoblasts or induce myogenesis earlier.
Parasites do not differentiate properly (tachyzoite overgrowth)
After infection, the parasitophorous vacuole forms normally, but tachyzoite growth is not quenched. Parasites continue their lytic cycle, thus overgrowing the whole culture. We encountered such behavior in particular with lab-adapted strains. To an extent, washing the monolayer more thoroughly in the first week post-infection and lowering the tachyzoite inoculum helps to avoid excess tachyzoites. If parasites recently isolated from mouse brain tissue are not available, gradual in vitro adaptation to bradyzoites enhances their ability to form cysts. These bradyzoites can either be used for infection directly or be re-differentiated into tachyzoites with higher cyst-forming capacity. It is advisable to generate freezer stocks of suitable tachyzoite cultures to ensure reliable and stable differentiation behavior. If parasite overgrowth cannot be avoided by the aforementioned measures, pharmaceutical inhibition of tachyzoite growth might help to allow bradyzoites to mature for longer. In that case, we recommend the use of 10 nM buparvaquone, a bc1 complex inhibitor.
Cyst numbers vary
Many variable host- and parasite-related factors influence this in vitro system. The comparatively long culture periods amplify these effects and lead to significant differences in total cyst numbers. Try to work as consistently as possible. Ensure that the myoblasts passage number is not too high, the myoblast confluency does not exceed 70% before differentiating them to myotubes, the monolayer has no gaps, the parasites are freshly egressed or released via syringing before infection, and that the parasites have not been cultured as tachyzoites for extended times. The latter is especially important when comparing bradyzoite formation efficiencies of different parasite strains. Using this protocol and a cystogenic parasite strain, approximately 106 cysts can be obtained from a T150 dish (Christiansen et al., 2022). After maturation, we occasionally observe multiple cysts per cell but few infected cells.
Myotubes detach from glass coverslips
KD3 myotubes can be grown and infected on glass coverslips for microscopy applications. We observe an increased rate of detachment of myotubes in these cultures. This may be compensated by coating the coverslips with rat collagen (see above), low levels of Cell-Tak, or gelatine.
Acknowledgments
We are grateful to Naohiro Hashimoto for initially sharing KD3 skeletal muscle cells, and Tohru Hosoyama, Tadakimi Tomita, and Louis Weiss for help in providing KD3 cells to the community. We thank Boris Striepen for providing strain Pru tdTomato. M.B. and D.M. were partially funded by the Federal Ministry of Education and Research (BMBF) under project number 01KI1715 as part of the “Research Network Zoonotic Infectious Diseases”. B.C. received funding from the international graduate school IRTG 2290, co-funded by the German Research Council (DFG). D.W. and E.D.B. received funding from the DFG via the graduate school GRK 2046. E.D.B., F.S., and M.B. also receive support from the NRP79 program of the Swiss National Science Foundation (SNSF), project number 407940. F.S. is a senior member of graduate schools GRK 2046 and IRTG 2290. M.B., D.M., D.W., and F.S. receive internal support from the Robert Koch-Institute. We also thank Celine Christiansen (Christiansen et al., 2022) for her contributions establishing the KD3 system.
Competing interests
The authors declare no competing interests.
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Real-Time Monitoring of ATG8 Lipidation in vitro Using Fluorescence Spectroscopy
WZ Wenxin Zhang
TN Taki Nishimura
ST Sharon A. Tooze
Published: Vol 14, Iss 1, Jan 5, 2024
DOI: 10.21769/BioProtoc.4917 Views: 650
Reviewed by: David PaulWeidong An Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Jun 2023
Abstract
Autophagy is an essential catabolic pathway used to sequester and engulf cytosolic substrates via a unique double-membrane structure, called an autophagosome. The ubiquitin-like ATG8 proteins play an important role in mediating autophagosome membrane expansion. They are covalently conjugated to phosphatidylethanolamine (PE) on the autophagosomes via a ubiquitin-like conjugation system called ATG8 lipidation. In vitro reconstitution of ATG8 lipidation with synthetic liposomes has been previously established and used widely to characterise the function of the E1 ATG7, the E2 ATG3, and the E3 complex ATG12–ATG5-ATG16L1. However, there is still a lack of a tool to provide kinetic measurements of this enzymatic reaction. In this protocol, we describe a real-time lipidation assay using NBD-labelled ATG8. This real-time assay can distinguish the formation of ATG8 intermediates (ATG7~ATG8 and/or ATG3~ATG8) and, finally, ATG8-PE conjugation. It allows kinetic characterisation of the activity of ATG7, ATG3, and the E3 complex during ATG8 lipidation. Furthermore, this protocol can be adapted to characterise the upstream regulators that may affect protein activity in ATG8 lipidation reaction with a kinetic readout.
Key features
• Preparation of ATG7 E1 from insect cells (Sf9 cells).
• Preparation of ATG3 E2 from bacteria (E. coli).
• Preparation of LC3B S3C from bacteria (E. coli).
• Preparation of liposomes to monitor the kinetics of ATG8 lipidation in a real-time manner.
Graphical overview
Experimental design to track the full reaction of ATG8 lipidation, described in this protocol
Keywords: Autophagy In vitro ATG8 lipidation Ubiquitin-like conjugation Real-time ATG8 lipidation assay Liposomes Site-directed fluorescence NBD Fluorescence spectroscopy
Background
Autophagy is a well-conserved bulk degradation pathway from yeast to mammals. It occurs ubiquitously in response to metabolic demands and plays an important role in maintaining cellular homeostasis and cell survival (Mizushima and Komatsu, 2011). Upon autophagy induction, cytosolic materials are sequestered and enclosed by a double-membrane structure, called an autophagosome, leading to the degradation of the content after fusion with lysosomes. One of the key discoveries in autophagy field is ATG8 lipidation via the ubiquitin-like conjugation systems (Mizushima, 2020; Nishimura and Tooze, 2020). The ubiquitin-like ATG8 is first primed by a cysteine protease ATG4 to expose its C-terminal glycine. Then, the activated ATG8 is conjugated to ATG7 (E1) in an ATP-dependent manner and transferred to ATG3 (E2). It is finally covalently conjugated to the headgroup of phosphatidylethanolamine (PE), catalysed by ATG12–ATG5-ATG16L1 complex (hereafter, the E3 complex). Lipidated ATG8 is commonly used as autophagosomal membrane marker, and ATG8 lipidation is monitored to assess the autophagy activity in cells (Mizushima et al., 2010).
In vitro reconstitution of ATG8 lipidation with synthetic membrane models, such as large unilamellar vesicles (LUVs), is also well established (reviewed in Huang et al., 2022). The end-point level of lipidated ATG8, which is usually examined by SDS-PAGE, has been used to assess the function of ATG7 (Taherbhoy et al., 2011; Noda et al., 2011), ATG3 (Nath et al., 2014), and the E3 complex (Lystad et al., 2019). However, this end-point readout cannot track ATG8 conjugation reactions in a real-time manner or provide more information on the reaction rate.
Recently, we designed a real-time lipidation assay using human ATG8 proteins (LC3B/GABARAP) N-terminally labelled with 7-nitrobenz-2-oxa-1,3-diazol-4-yl (NBD) (Zhang et al., 2023). As NBD fluoresces brightly around 535 nm when it is located in a hydrophobic environment, we use it to dynamically track the hydrophobic environment that ATG8 N-terminus encounters during the lipidation reaction (provided by the ATG7 or ATG3 interfaces and/or the membrane). Interestingly, the increased NBD signals in the real-time assay, in addition to responding to membrane environments, also reflect the enzymatic activity of ATG7, ATG3, and the E3 complex, which can be readily adapted to evaluate the function of ATG7, ATG3, the E3 complex, or other upstream regulators, and provide a kinetic measurement in future studies. This protocol provides the setup for the real-time lipidation assay using NBD-labelled LC3B, as an example.
Materials and reagents
Sf9 cells (ATCC, catalog number: CRL-1711)
BL21 (DE3) E. coli (New England BioLabs, catalog number: C2527H)
Sf-900TM II SFM medium (Thermo Fisher Scientific, catalog number: 10902104)
Fugene HD transfection reagent (Promega, catalog number: E2311)
flashBAC GOLD (Oxford Expression Technologies, catalog number: 100202)
Gentamicin (10 mg/mL) (Thermo Fisher Scientific, catalog number: 15710049)
Amphotericin B (Thermo Fisher Scientific, catalog number: 15290018)
Centrifugal filters (0.2 μm) (VWR, catalog number: 516-0233)
Ampicillin (Sigma-Aldrich, catalog number: A9518-25G)
Kanamycin (Sigma-Aldrich, catalog number: K4000-25G)
Isopropyl-β-D-thio-galactopyranoside (IPTG) (Melford, catalog number: I56000-25.0)
Tris Base (Fisher Scientific, catalog number: BP1521)
Hydrochloric acid (HCl), 37% (Fisher Scientific, catalog number: 10053023)
Sodium chloride (NaCl) (Fisher Scientific, catalog number: 15526005)
Tris(2-carboxyethyl)phosphine hydrochloride solution (TCEP) (Sigma-Aldrich, catalog number: 646547-10X1ML)
AEBSF hydrochloride (AppliChem, catalog number: A1421)
Benzamidine (Sigma-Aldrich, catalog number: B6506-25G)
cOmpleteTM, EDTA-free protease inhibitor cocktail (Sigma-Aldrich, catalog number: 11873580001)
Glutathione Sepharose® 4B beads (Cytiva, catalog number: 17075605)
IANBD (Invitrogen, catalog number: D2004)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D2650)
L-cysteine (Sigma-Aldrich, catalog number: 168149)
Glycerol (Sigma-Aldrich, catalog number: 2025)
100 mM MgATP (R&D Systems, Inc, catalogue number B-20)
Lipids
16:0-18:1 PC (POPC) (Avanti Polar Lipids, catalog number: 850457C)
18:1 (Δ9-Cis) PE (DOPE) (Avanti Polar Lipids, catalog number: 850725C)
Solutions
Lysis buffer 1 (see Recipes)
Lysis buffer 2 (see Recipes)
Wash buffer (see Recipes)
Equilibration buffer (see Recipes)
Labelling buffer (see Recipes)
Assay buffer (see Recipes)
Recipes
Lysis buffer 1
50 mM Tris-HCl, pH 8.0
500 mM NaCl
0.5 mM TCEP
0.4 mM AEBSF
15 μg/mL benzamidine
Store at 4 °C, use within three days.
Lysis buffer 2
50 mM Tris-HCl, pH 8.0
500 mM NaCl
0.5 mM TCEP
1× cOmpleteTM, EDTA-free protease inhibitor
Store at 4 °C, use within three days.
Wash buffer
50 mM Tris-HCl, pH 8.0
500 mM NaCl
0.5 mM TCEP
Store at 4 °C, use within two weeks.
Equilibration buffer
25 mM Tris-HCl, pH 8.0
150 mM NaCl
0.5 mM TCEP
Store at 4 °C, use within two weeks.
Labelling buffer
25 mM Tris-HCl, pH 7.5
150 mM NaCl
Store at 4 °C, use within two weeks.
Assay buffer (the same as equilibration buffer)
25 mM Tris-HCl, pH 8.0
150 mM NaCl
0.5 mM TCEP
Store at 4 °C, use within two weeks.
Equipment
Sonicator (MSE, model: Soniprep 150)
Vivaspin 20 concentrator column (MWCO 10K) (Cytiva, catalog number: 28932360)
Vivaspin 6 concentrator column (MWCO 30K) (Cytiva, catalog number: 28932317)
ӒKTA pureTM equipped with Superdex 200 16/60 GL column (Cytiva, catalog number: 28-9893-35) and Superdex 200 10/300 GL column (Cytiva, catalog number: 17-5175-01)
PD MinitrapTM G-25 (Sigma-Aldrich, catalog number: GE28-9180-07)
Extruder set with holder/heating block including mini-extruder, two syringes, polycarbonate membranes 0.1 µm 19 mm, 100 filter supports, and one holder/heating block (Avanti Polar Lipids, catalog number: 610000-1EA)
Polycarbonate membranes 0.2 μm 19 mm (Avanti Polar Lipids, catalog number: 610006-1EA)
EppendorfTM Concentrator Plus (Eppendorf, catalog number: 5305000568)
Zetasizer Nano ZS (Malvern Instruments)
Ultra-micro cuvette 10 mm pathlength, 100 μL (Hellma Analytics, catalog number: 105.201-QS)
Spectrofluorometer (JASCO, model: FP-8300)
Ti45 rotor (Beckman, 339160)
Software and datasets
Excel (Microsoft Office, https://www.office.com/)
GraphPad Prism 9 (GraphPad Software, https://www.graphpad.com/)
Procedure
ATG3 expression and purification from BL21 (DE3) E. coli
Transform pGEX6P1-GST-3C-ATG3 into BL21 (DE3) E. coli cells, spread the bacteria on an LB agar plate containing 100 μg/mL of ampicillin, and incubate the plate at 37 °C overnight.
Inoculate a single colony into 120 mL of LB medium containing 100 μg/mL ampicillin in a 250 mL glass flask and incubate at 37 °C shaking at 180 rpm overnight in a shaking incubator.
Dilute 25–30 mL of the culture to 1 L of the LB medium containing 100 μg/mL ampicillin in a 2 L glass flask (OD600 around 0.1). In total, prepare 4 L of culture. Grow the culture for 3–5 h at 37 °C shaking at 200 rpm in a shaking incubator, until OD600 reaches 0.8.
Induce protein expression by adding 500 μL of 1 M IPTG (final concentration 0.5 mM) and incubate at 18 °C shaking at 200 rpm in a shaking incubator overnight.
Pellet the bacteria at 4,000 rpm for 15 min and discard the medium.
Resuspend the bacteria pellet in 50 mL of lysis buffer 1 and directly place in a -20 °C freezer for at least one day to break the cells. This is an optional step; the aim is to break the bacteria through gradual ice crystal formation.
Thaw the bacteria pellets in ice-cold water and sonicate the pellet on ice at maximum amplitude of 20 μm for 40 s, interval 60 s. Repeat 5–10 times until the cell lysate becomes less viscous.
Pellet cell debris using the Ti 45 rotor at 25,000× g at 4 °C for 30 min.
Collect supernatant and add 4 mL of Glutathione Sepharose® 4B slurry (i.e., 2 mL of Glutathione Sepharose® 4B beads). Incubate rolling at 4 °C for 90 min.
Pellet the beads at 500× g for 3 min at 4 °C. Wash the beads with wash buffer five times.
Resuspend the beads in 10 mL of wash buffer, add 200 μL of GST-3C (5 mg/mL, obtained from Crick Structural Biology Platform), and incubate rolling at 4 °C overnight. GST-3C protease is also commercially available, and can be purchased from Merck (PreScission Protease, catalog number: GE27-0843-01).
Collect eluate and concentrate the protein with a Vivaspin 20 concentrator column (MWCO 10K) until the volume reaches 1–2 mL.
Further purify the protein by size exclusion chromatography (SEC) using the Superdex 200 16/60 GL column equilibrated in equilibration buffer.
Check the protein purity by SDS-PAGE, collect the peak fraction, and concentrate the protein using a Vivaspin 20 concentrator column (MWCO 10K). Final stock protein concentration is usually ~6 mg/mL.
Flash freeze in liquid N2 and store at -80 °C until use.
LC3B S3C expression and purification from BL21 (DE3) E. coli
Transform pAL-GST-3C-LC3B(1-120) S3C construct into BL21 (DE3) E. coli cells, spread the bacteria on an LB agar plate containing 50 μg/mL of kanamycin, and incubate the plate at 37 °C overnight.
Inoculate a single colony into 120 mL of LB medium containing 50 μg/mL kanamycin in a 250 mL glass flask and incubate at 37 °C shaking at 180 rpm overnight in a shaking incubator.
Dilute 25–30 mL of the culture to 1 L of the LB medium containing 50 μg/mL kanamycin in a 2 L glass flask (OD600 around 0.1). In total, prepare 4 L of culture. Grow the culture for 3–5 h at 37 °C shaking at 200 rpm in a shaking incubator, until OD600 reaches 0.8.
Induce protein expression by adding 500 μL of 1 M IPTG (final concentration 0.5 mM) and incubate at 18 °C shaking at 200 rpm in a shaking incubator overnight.
Pellet the bacteria at 4,000 rpm for 15 min and discard the medium.
Resuspend the bacteria pellet in 50 mL of lysis buffer 1 and directly place in a -20 °C freezer for at least one day to break the cells. This is an optional step; the aim is to break the bacteria through gradual ice crystal formation.
Purification steps are performed as described in section A (steps 7–14). Final stock concentration is usually ~7 mg/mL.
Flash freeze in liquid N2 and store at -80 °C until use.
Note: This purification protocol was also used to purify other ATG8 proteins.
ATG7 expression and purification from insect cells Sf9
Filter the transfer plasmid pBacPAK-His3-GST-ATG7 with centrifugal filters (0.2 μm) to sterilise before use.
Seed 0.5 × 106 Sf9 cells per well in a 6-well plate and incubate in a static incubator at 27 °C for at least 1 h.
Prepare the transfection mix: add 0.5 μg of the transfer plasmid and 50 ng of flashBAC GOLD DNA into 200 μL of Sf-900TM II SFM medium. Then, add 2 μL of Fugene HD transfection reagent. Incubate at room temperature for 20 min.
Add 800 μL of Sf-900TM II SFM medium to the transfection mix.
Aspirate the medium from the cells in the 6-well plate and add ~1 mL of the mixture on the top of cells. Incubate in a static incubator at 27 °C overnight.
Add another 1 mL of Sf-900TM II SFM medium containing gentamicin (10 μg/mL) and amphotericin B (0.25 μg/mL) on the top of the cells in the 6-well plate.
Incubate for another four days in a static incubator at 27 °C. Harvest and pool the cells and supernatant and add 1.5 mL of the cell mixture into 50 mL of S9 cells at a density of 1 × 106 cells/mL. Incubate at a shaking incubator at 27 °C and 140 rpm for three days.
Collect the supernatant, which contains baculovirus harbouring His3-GST-ATG7 construct.
Note: The successful production of baculovirus harbouring His3-GST-ATG7 construct was checked by immunoblotting ATG7 or the GST-tag in the cell pellet.
Culture 400 mL of Sf9 cells in Sf-900TM II SFM medium to a density of 1.5 × 106 cells/mL and add 1 mL of baculovirus harbouring His3-GST-3C-ATG7 construct. Incubate in a shaking incubator at 27 °C and 140 rpm.
After 60 h of infection, harvest the cells by centrifugation at 600× g for 10 min at 4 °C.
Freeze at -80 °C until ready for purification.
Thaw the cell pellet on ice, resuspend it with 30 mL of lysis buffer 2, and sonicate the mixture on ice at 50% amplitude for 10 s, interval 60 s. Repeat 3–5 times.
Pellet cell debris using the Ti 45 rotor at 30,000× g for 30 min at 4 °C.
Collect supernatant and add 2 mL of Glutathione Sepharose® 4B slurry (i.e., 1 mL of Glutathione Sepharose® 4B beads). Incubate rolling at 4 °C for 90 min.
Pellet the beads at 500× g for 3 min at 4 °C. Wash the beads with wash buffer five times.
Resuspend the beads in 5 mL of wash buffer, add 100 μL of GST-3C (5 mg/mL, obtained from Crick Structural Biology Platform), and incubate rolling at 4 °C overnight.
Collect eluate and concentrate the protein with a Vivaspin 6 concentrator column (MWCO 30K) until the volume reaches 300–500 μL.
Further purify the protein by SEC using the Superdex 200 10/300 GL column equilibrated in equilibration buffer.
Check the protein purity by SDS-PAGE, collect the peak fraction, and concentrate the protein using a Vivaspin 6 concentrator column (MWCO 30K). Final stock concentration is ~2 mg/mL.
Flash freeze in liquid N2 and store at -80 °C until use.
NBD-labelling of ATG8 proteins with single cysteine mutation (LC3B S3C)
Prepare 10 mM IANBD-amide stock dissolved in DMSO. For longer storage, keep the stock at -20 °C.
Equilibrate a PD Minitrap G-25 column with 8 mL of labelling buffer. Apply 250 μL of 5 mg/mL LC3B S3C to the column and collect the eluate.
Note: Alternative sources of desalting columns or other buffer exchange methods such as dialysis can be used as well.
After buffer exchange, re-measure the protein concentration, prepare 400 μL of ~100 μM LC3B S3C, and mix with 100 μL of 10 mM IANBD-amide. Incubate the reaction mix at room temperature for 1 h in the dark.
Add 20 μL of 100 mM L-cysteine to quench the reaction.
Equilibrate another PD Minitrap G-25 column with 8 mL of assay buffer. Apply ~500 μL of the reaction mix on the column and collect the eluate. This step removes the excess IANBD-amide.
Measure protein concentration by Bradford assay and run SDS-PAGE to check the NBD labelling and if there is no excess dye (unreacted dye migrates in dye front).
Mix 125 μL of glycerol with 500 μL of NBD-labelled LC3B S3C to a final concentration of 20%. Aliquot the protein, flash freeze in liquid N2, and store at -80 °C.
Liposome preparation
To prepare liposome stock (1 mL, 2 mM final concentration)
Mix POPC and DOPE lipids at a ratio of 50:50 (% mol).
Dry the lipids under nitrogen gas for 5 min and further dry in the EppendorfTM Concentrator for 2 h.
Add 1 mL of assay buffer onto the lipid film and vortex thoroughly until the lipid film is fully resuspended.
Conduct five freeze-thaw cycles in liquid nitrogen and 42 °C water bath until the lipid solution is fully thawed.
Prepare the unilamellar vesicles with the Mini Extruder. Extrude the liposome solution first by passing the solution 21 times through a 0.2 μm membrane. Then, extrude the solution by passing it at least 41 times through a 0.1 μm membrane.
Note: It is recommended to extrude the liposomes through a 0.2 μm membrane before a 0.1 μm membrane in order to make homogeneous liposomes.
Check the liposome size by Zetasizer Nano ZS [for example, Figure 1: average size 105.5 nm, PDI (polydispersity index) = 0.071]. If the size of liposomes is not homogeneous (PDI > 0.2), repeat the extrusion step through the 0.1 μm membrane.
Figure 1. Result of liposome size distribution measured by dynamic light scattering [polydispersity index (PDI) = 0.071]
Keep the liposomes at 4 °C and use within two days.
Real-time ATG8 lipidation assay
Purified ATG7, ATG3, LC3B S3C, and LC3B S3CNBD are resolved by SDS-PAGE (Figure 2).
Figure 2. SDS-PAGE result of purified ATG7, ATG3, LC3B S3C, and NBD-labelled LC3B S3C
Table 1 indicates the concentration and volume for each component in the real-time lipidation ATG8 assay.
Table 1. Components for the real-time ATG8 lipidation assay (total volume: 78.4 μL)
Component Final concentration Volume to add (stock concentration)
ATG7 0.2 μM 0.8 μL (20 μM)
ATG3 0.2 μM 0.8 μL (20 μM)
E3 complex
Liposomes
LC3B S3CNBD
0.05 μM
1 mM
1 μM
0.2 μL (20 μM)
40 μL (2 mM)
1.6 μL (50 μM)
Assay buffer n/a 35 μL
Note: To characterise the effect of each component on the real-time assay, add the amount of buffer instead of the component of interest. For example, for “no ATG7” condition, prepare the reaction mix without ATG7 and add buffer instead. The protein stock can be diluted to a lower concentration so that it will be easier to pipette a larger volume of protein.
Transfer the reaction mix into a 10 mm path length quartz cuvette (100 μL).
Record NBD fluorescence (ex/em 468 nm/535 nm) over time using a FP-8300 spectrofluorometer.
Set cuvette holder temperature to 37 °C.
Set excitation at 468 nm (bandwidth 5 nm) and emission at 535 nm (bandwidth 10 nm).
Set the total measurement time to 20 min with two measurement ranges:
Time interval 1: 0–80 s; record NBD fluorescence every 20 s (equilibration step before the reaction).
Time interval 2: 80–1,200 s; record NBD fluorescence every 10 s (measurement step after adding ATP).
Start the measurement. By the end of time interval 1 (between 60 and 80 s), add 1.6 μL of MgATP (50 mM) to the reaction (final concentration of MgATP: 1 mM) and mix properly.
Repeat these steps for each condition: no ATG7, no ATG3, no E3, no liposomes, no ATP, and All.
Note: With “no ATP” condition, add 1.6 μL of assay buffer instead between 60 and 80 s (the end of time interval 1).
Data analysis
The NBD fluorescence increase ΔEm535 nm at each time point in the reaction groups was calculated by subtracting the NBD fluorescence recorded from control group (i.e., no ATP condition), as shown in Figure 3A.
The relative fluorescence ΔF at each time point was normalised to the time point at 80, as shown in Figure 3B.
Analyse the data in Excel and plot data in Prism 9.0.
Figure 3. Example of the real-time lipidation assay. (A) Raw data of NBD spectra over time. After adding ATP between 60 and 80 s, signals were increased except for “no ATG7” and “no ATP” conditions. The NBD signal increase (ΔEm535 nm) was calculated by subtracting the NBD signal from the control group no ATP condition from the time point 80 s. (B) Relative fluorescent increase normalised to no ATP condition at time point 80 s. One repeat of three independent experiments is shown here.
Validation of protocol
This protocol was used in Zhang et al. (2023), DOI: 10.7554/eLife.89185
General notes and troubleshooting
The E3 complex used in this protocol was purified by Anne Schreiber (The Francis Crick Institute), as previously described in Zhang and Nishimura et al. (2023). The protein purification protocol for the in vitro ATG8 lipidation reaction has been described previously, for example Landajuela et al. (2016), Zheng et al. (2017), Lystad et al. (2019), and Fracchiolla et al. (2020).
This protocol employed a simple lipid model, which is ~100 nm liposomes containing 50% PC and 50% PE. To further assess the effects of specific lipids or membrane curvature on ATG8 lipidation reaction, the lipid composition and liposome size can be modified, respectively, according to the experimental purpose.
Acknowledgments
We thank Alicia Alonso for human ATG7 and ATG3 plasmids, Svend Kjaer for pBacPAK-His3-GST plasmid and the technical assistance with insect cell culture. We thank Simone Kunzelmann for the technical assistance and helpful suggestions on fluorescence spectroscopy. We thank Colin Davis and Anne Schreiber for providing the E3 complex. This work was funded by The Francis Crick Institute, which receives its core funding from Cancer Research UK (CC2134 to S.A.T.), the UK Medical Research Council (CC2134 to S.A.T.). This research was funded in whole, or in part, by the Wellcome Trust (CC2134 to S.A.T.). W.Z. and S.A.T received funding from the European Research Council under the European Union's Seventh Framework Programme (FP7/2007-2013)/ERC grant agreement n° 788708. For the purpose of Open Access, the author has applied a CC BY public copyright licence to any Author Accepted Manuscript version arising from this submission. This study was supported by PRESTO (JPMJPR20EC to T.N.) from Japan Science and Technology (JST), a Grant-in-Aid for Transformative Research Areas (B) (grant 21H05146 to T.N.) from the Japan Society for the Promotion of Science (JSPS), and a grant from the Japan Foundation for Applied Enzymology (to T.N.). This protocol is based on our previous publication Zhang et al. (2023).
Competing interests
The authors declare no competing interests.
References
Fracchiolla, D., Chang, C., Hurley, J. H. and Martens, S. (2020). A PI3K-WIPI2 positive feedback loop allosterically activates LC3 lipidation in autophagy. J Cell Biol 219(7): e201912098.
Huang, X., Yao, J., Liu, L., Luo, Y. and Yang, A. (2022). Atg8-PE protein-based in vitro biochemical approaches to autophagy studies. Autophagy 18(9): 2020–2035.
Landajuela, A., Hervas, J. H., Anton, Z., Montes, L. R., Gil, D., Valle, M., Rodriguez, J. F., Goni, F. M. and Alonso, A. (2016). Lipid Geometry and Bilayer Curvature Modulate LC3/GABARAP-Mediated Model Autophagosomal Elongation. Biophys J 110(2): 411–422.
Lystad, A. H., Carlsson, S. R., de la Ballina, L. R., Kauffman, K. J., Nag, S., Yoshimori, T., Melia, T. J. and Simonsen, A. (2019). Distinct functions of ATG16L1 isoforms in membrane binding and LC3B lipidation in autophagy-related processes. Nat Cell Biol 21(3): 372–383.
Mizushima, N. (2020). The ATG conjugation systems in autophagy. Curr Opin Cell Biol 63: 1–10.
Mizushima, N. and Komatsu, M. (2011). Autophagy: renovation of cells and tissues. Cell 147(4): 728–741.
Mizushima, N., Yoshimori, T. and Levine, B. (2010). Methods in mammalian autophagy research. Cell 140(3): 313–326.
Nath, S., Dancourt, J., Shteyn, V., Puente, G., Fong, W. M., Nag, S., Bewersdorf, J., Yamamoto, A., Antonny, B. and Melia, T. J. (2014). Lipidation of the LC3/GABARAP family of autophagy proteins relies on a membrane-curvature-sensing domain in Atg3. Nat Cell Biol 16(5): 415–424.
Nishimura, T. and Tooze, S. A. (2020). Emerging roles of ATG proteins and membrane lipids in autophagosome formation. Cell Discov 6(1): 32.
Noda, N. N., Satoo, K., Fujioka, Y., Kumeta, H., Ogura, K., Nakatogawa, H., Ohsumi, Y. and Inagaki, F. (2011). Structural basis of Atg8 activation by a homodimeric E1, Atg7. Mol Cell 44(3): 462–475.
Taherbhoy, A. M., Tait, S. W., Kaiser, S. E., Williams, A. H., Deng, A., Nourse, A., Hammel, M., Kurinov, I., Rock, C. O., Green, D. R. and Schulman, B. A. (2011). Atg8 transfer from Atg7 to Atg3: a distinctive E1-E2 architecture and mechanism in the autophagy pathway. Mol Cell 44(3): 451–461.
Zhang, W., Nishimura, T., Gahlot, D., Saito, C., Davis, C., Jefferies, H. B. J., Schreiber, A., Thukral, L. and Tooze, S. A. (2023). Autophagosome membrane expansion is mediated by the N-terminus and cis-membrane association of human ATG8s. eLife 12: e89185.
Zheng, Y., Qiu, Y., Gunderson, J. E. and Schulman, B. A. (2017). Production of Human ATG Proteins for Lipidation Assays. Methods Enzymol 587: 97–113.
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Live Imaging and Analysis of Meiotic Cytokinesis in Drosophila Testes
GK Govind Kunduri
JA Jairaj K. Acharya
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4918 Views: 1056
Reviewed by: Giansanti MariagraziaPradeep Kumar Bhaskar Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in PLOS Biology Sep 2022
Abstract
All living organisms require the division of a cell into daughter cells for their growth and maintenance. During cell division, both genetic and cytoplasmic contents are equally distributed between the two daughter cells. At the end of cell division, cytoplasmic contents and the plasma membrane are physically separated between the two daughter cells via a process known as cytokinesis. Hundreds of proteins and lipids involved in the cytokinetic process have been identified; however, much less is known about the mechanisms by which these molecules regulate cytokinesis, being therefore an intense area of current research. Male meiotic cytokinesis in Drosophila melanogaster testes has been shown to be an excellent model to study cytokinesis in vivo. Currently, several excellent protocols are available to study cytokinesis in Drosophila testes. However, improved methods are required to study cytokinesis under in vitro and ex vivo conditions. Here, we demonstrate a simple method to perform live imaging on individual spermatocyte cysts isolated from adult testes. We evaluate amenability of this in vitro method for treatment with pharmacological agents. We show that cytokinesis is strongly inhibited upon treatment with Dynasore, a dynamin inhibitor known to block clathrin-mediated endocytosis. In addition, we also demonstrate an ex vivo method to perform live imaging on whole mount adult testes on gas permeable membrane chambers. We believe the protocols described here are valuable tools to study cytokinetic mechanisms under various genetic and treatment conditions.
Key features
• In vitro method to study male meiotic cytokinesis in dissected spermatocyte cysts.
• In vitro method allows acute treatment with various pharmacological agents to study cytokinesis.
• Ex vivo method to image male meiosis cytokinesis in intact adult testes.
• Requires 15–60 min to set up and could be imaged up to 6–12 h.
Graphical overview
In vitro and ex vivo live imaging of male meiotic cytokinesis in adult Drosophila testes
Keywords: Live imaging Cytokinesis Male meiosis cytokinesis Drosophila melanogaster Spermatocyte cysts Spermatogenesis Testis Dynasore
Background
Cytokinesis is a final step of cell division wherein parent cell cytoplasmic contents and plasma membrane are physically divided between the two daughter cells. In animal cells, immediately after chromosomes are segregated to the poles, the spindle-mediated mechanisms determine the site of actomyosin ring assembly at the cell cortex. Subsequently, the actomyosin contractile ring that anchors the plasma membrane, and the central spindle begins to constrict, leading to the generation of cleavage furrow (Figure 1B). In mitotic cells, the plasma membrane between two daughter cells is pinched off in a process known as abscission, leading to the physical separation of daughter cells. However, unlike mitotic cells, in male germ cells abscission does not occur at the end of cytokinesis. As a result, all the daughter cells originating from the gonioblast are interconnected via their cytoplasmic bridges, known as ring canals. It is thought that the ring canals play an important role in intercellular communication and signaling [1]. All the cells in a cyst connected via cytoplasmic bridges undergo synchronous divisions [2, 3] (Video 1).
Video 1. In vitro live cell imaging of isolated adult Drosophila spermatocyte cyst undergoing meiosis I and meiosis II cytokinesis. The cysts are isolated from fly testes expressing GFP-tagged PH domain of Grp1/Step protein under the control of tubulin 84B promoter (tGPH) and mCherry tagged fascetto (feo) protein under the control of Ubi-p63E promoter (Feo-mCherry). The GFP-PH-Grp1/Step localizes to cytosol, plasma membrane, and cleavage furrow in cysts with intact cyst cell membrane. However, tGPH localizes only to the cytosol in cysts that have lost cyst cell membrane envelop. Feo-mCherry localizes to central spindle, independent of cyst cell membrane envelop. Live imaging was performed with the 63× water immersion objective and a pre-defined area of interest (AOI) size of 1024 × 1024 and at intervals of 2 min for 120 repeats. Video shown here represents 10 frames/second.
Drosophila melanogaster has been shown to be a valuable genetic model to study cytokinesis in vivo. Conventionally, Drosophila embryos undergoing cellularization [4, 5], proliferating larval neuroblasts [6, 7], and male meiotic spermatocytes [8, 9] have been used in cytokinesis studies. Meiotic spermatocytes in Drosophila testes offer several unique advantages over other systems to study cytokinesis. These include the relatively large size of meiotic spindle, weak spindle checkpoint, unambiguous identification of cytokinetic mutants due to characteristic morphological differences, and finally, live imaging of spermatocytes in isolated cysts or intact pupal testes. The relatively large size of male meiotic spindle is a favorable cytological feature in spermatocytes that allows precise localization of many cytokinetic proteins including microtubule and contractile ring–associated proteins. Male meiotic cells also show prominent central spindle (Figure 1B) (a bundle of antiparallel, interdigitating microtubules localizes to the center of segregating chromosomes where the assembly of contractile ring begins) [10, 11]. In neuroblasts, the presence of an abnormal spindle activates spindle checkpoint, which in turn arrests the cells in metaphase and precludes the observation of subsequent steps in cell division including cytokinesis [11–14]. However, due to weak spindle checkpoint, spermatocytes continue through the cell division stages despite the abnormal spindle. Thus, using meiotic cytokinesis, it is possible to ask whether a gene involved in spindle formation also has a role in later stages of cell division including cytokinesis. For instance, it was shown that mutations in the polo and abnormal spindle (asp) genes cause metaphase arrest in larval neuroblasts but produce frequent cytokinetic failures in spermatocytes [12–14]. Drosophila male meiosis is also an excellent model for cytokinesis due to the unique organization of mitochondria in round spermatids. During male meiotic cytokinesis, mitochondria is equally partitioned between the two daughter cells; immediately after cytokinesis, it aggregates to form a phase-dark structure known as Nebenkern. Failure in cytokinesis abrogates mitochondrial segregation; as a result, round spermatids show unusually large mitochondria associated with multiple nucleus (Figure S1) [15]. Thus, the presence of unusually large mitochondria with multiple nucleus is a characteristic feature of meiotic cytokinesis in spermatocytes (Figure S1) [3, 15].
Figure 1. Male meiotic cytokinesis in adult Drosophila testis. (A) Cartoon representing dissected adult testis showing various stages of germ cell differentiation. Drosophila spermatogenesis shares a significant amount of similarity with the mammalian system. In many species, including Drosophila melanogaster, male germ cell differentiation is a sequential process. It begins with the germline stem cell (GSC) attached to the hub cells at the tip of the testis undergoing asymmetric division to yield one committed progenitor called gonioblast (GB). Two cyst stem cells (CySCs) that surround the GSC also undergo asymmetric division to produce two cyst cells that encase the GB. Subsequently, the GB undergoes four rounds of mitotic divisions, resulting in the 16-cell stage known as spermatogonia (SG). With the premeiotic S phase and an impressive growth during G2 phase due to robust transcriptional activity, spermatogonia become spermatocytes (SC), which in turn synchronously undergo two rounds of meiotic divisions (meiosis I & meiosis II) resulting in the haploid, 64-cell stage known as round spermatids (RS). Due to a lack of abscission step at the end of cytokinesis in germ cells, all cell divisions originating from a single GB are connected to each other via cytoplasmic bridges known as ring canals (RCs). After completion of meiosis, all the mitochondria in individual spermatids aggregate around the basal body at one end of the nucleus to form a unique structure called nebenkern. Subsequently, round spermatids elongate via complex morphological changes including polarization of spermatid nucleus, dramatic nuclear size reduction due to histamine to protamine switch, elongation of nebenkern, axoneme growth, and formation of acrosome, a membrane-bound organelle required for fertilization. Finally, the individual sperms are physically separated by a complex process known as individualization, wherein actin-based cones migrate from head to tail fashion with the concomitant removal of cytoplasmic contents [cystic bulge (CB) and waste bag (WB)] and wrapping each spermatozoon with its own plasma membrane. (B) Representative live images showing localization of central spindle–associated protein feo (fascetto tagged with mCherry) (left panel) and a plasma membrane–associated protein tGPH (PH domain of Grp1fused to GFP) (middle panel), during cytokinesis. Right panel shows the merge of both channels and arrow indicates the site of cleavage furrow.
Several excellent protocols have been described in the literature for the isolation of Drosophila testes from embryo to adult developmental stages and ex vivo imaging of whole mount pupal testes and isolated spermatocyte cysts [16–22]. However, detailed protocols focused on live imaging of male meiotic cytokinesis in dissected germ line cysts have not been described. Further, live imaging of male meiotic cytokinesis in whole mount adult testes has not been demonstrated. The protocols described here build upon the methods developed by Gartner et al. [18] and Karabasheva, G. and Smyth, J.T. [19], extending their application to male meiotic cytokinesis in adult Drosophila testes. Gartner et al. have standardized a method for live imaging of post-meiotic histone to protamine switch in intact pupal testes and dissected pupal germ line cysts [18]. Karabasheva, G. and Smyth J.T. established a method for live imaging of male meiotic cytokinesis in intact pupal testes [19]. Here, we demonstrate both in vitro and ex vivo methods to image male meiotic cytokinesis in adult testes. The in vitro method involves isolation of male germline cysts and imaging in a cover glass bottom dish. This in vitro method is of particular interest when the experimental settings require direct treatment of spermatocytes with external agents such as proteins, lipids, and pharmacological agents. Using this method, we show that treatment of spermatocytes with Dynasore, a dynamin inhibitor known to block clathrin-mediated endocytosis [23], also blocks male meiotic cytokinesis by preventing normal assembly of central spindle. The ex vivo method presented here involves dissection of adult testes and imaging on gas permeable membrane dish chambers. Both in vitro and ex vivo methods demonstrated here are extremely valuable to study male meiotic cytokinesis under various treatment conditions and genetic backgrounds.
Materials and reagents
Biological materials
Refer to Giansanti, G.M et al. [24], for a list of tagged proteins that can be used to mark central spindle/spindle midzone and cleavage furrow. For analysis of cytokinesis, a few examples of fly stocks are listed below (Table 1).
Table 1.List of fly stocks to label plasma membrane, central spindle, and actomyosin contractile ring .
No Protein Localization Fly stock number/reference
1 PH domain of Steppke/Grp1, expressed under the control of Tub84B promoter (tGPH), binds to PIP3 Plasma membrane, enriched on early and late stage of cleavage furrow membranes BDSC#8163
2 Ubi-p63E-Feo-mCherry, mCherry-tagged feo protein is expressed under the control of ubiquitin regulatory sequence Spindle midzone/central spindle BDSC#59277
3 Sqh-GFP.RLC, Myosin II regulatory light chain tagged to GFP is expressed under its own promoter control Contractile ring BDSC#57145
4 UAS-mRFP-Scraps, PH domain containing Anillin protein tagged with mRFP expressed under UAS control Cleavage furrow/contractile ring BDSC#52220
5 UAS-PLCdels-PH-EGFP, PH domain of human PLCD1 tagged with EGFP expressed under UAS control Plasma membrane and cleavage furrow BDSC#39693
Reagents
Shields and Sang M3 insect media (Sigma-Aldrich, catalog number: S8398-1L)
Heat-inactivated FBS suitable for insect cell culture (Thermo Fisher Scientific, catalog number: 10082147)
Penicillin-Streptomycin (10,000 U/mL) (Thermo Fisher Scientific, catalog number: 15140122)
Distilled water, 1,000 mL (Thermo Fisher Scientific, catalog number: 15230147)
Poly-D-Lysine hydrobromide (Sigma-Aldrich, catalog number: P0899)
Potassium bicarbonate (KHCO3) (Sigma-Aldrich, catalog number: 237205)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D2650)
Dynasore hydrate (Sigma-Aldrich, catalog number: D7693)
Ethanol (70 %)
M3 media (see Recipes)
Solutions
200 μg/mL Poly-D-Lysine hydrobromide in distilled water
31 mM or 10 mg/mL Dynasore hydrate in DMSO
Recipes
M3 media
Prepare this recipe in a biological safety cabinet. Dispense one bottle of powdered media (Shields and Sang M3 insect media) into a 1 L beaker (caution: may cause irritation to eyes and skin) and add 0.5 g of KHCO3 and 800 mL of sterile distilled water. Dissolve the powder completely using a magnetic stirrer and a bar. Subsequently, add 100 mL of heat-inactivated FBS and 10 mL of penicillin/streptomycin solution and mix well on a magnetic stirrer. Make up the final volume to 1 L and filter sterilize by passing through a bottle-top filter unit with 0.2 μm pore sized membrane. Aliquot the filter-sterilized media into 50 mL Falcon tubes and store at -20 °C. On the day of experiment, thaw one aliquot of media in a 37 °C water bath and filter the media again into a fresh Falcon tube using a 0.2 μm syringe filter. When performing the experiment, media should be at room temperature. Excess media can be stored at 4 °C for up to 1–2 weeks.
Laboratory supplies
Glass-bottom cell culture dish, 15 mm (NEST, catalog number: 801002)
Glass well plate, 9 wells (Corning Life Sciences, catalog number: 7220-85)
Dumont#5 ceramic-coated forceps (Fine Science Tools, catalog number: 11252-50)
Dissection pins, tip diameter 0.0175 mm (Fine Science Tools, catalog number: 26002-15)
Pin holders (Fine Science Tools, catalog number: 26018-17)
Lumox® dish 50 (Sarstedt, catalog number: 94.6007.410)
SecureSeal imaging Spacers, 0.12 mm depth (Grace Bio-Labs, catalog number: 654008)
Disposable sterile filter units, 500 mL (Thermo Fisher Scientific, catalog number: 450-0020)
Syringe filters, sterile, 0.22 um (GenClone®, catalog number: 25-244)
BD 10 mL syringe, luer-lok tip (BD, catalog number: 302995)
Microtubes, clear 1.7 mL (Olympus plastics, catalog number: 24-281)
Falcon, 50 mL, sterile, polypropylene conical tube, (Corning, catalog number: 352098)
15 mL centrifuge tube, sterile polypropylene tube (Corning, catalog number: 430052)
Equipment
Zeiss SteREO Discovery microscope (Zeiss Group, model: V12)
Spinning disk confocal on Leica DMi8 microscope base (Andor)
Software and datasets
Andor Fusion (version 2.3.0.44) (https://andor.oxinst.com/downloads/view/fusion-release-2.3)
Imaris (version 10.0) (https://imaris.oxinst.com/)
Fiji/ImageJ (https://imagej.net/software/fiji/downloads)
Prism 9 (https://www.graphpad.com/features)
Procedure
In vitro live cell imaging of male meiotic cytokinesis in isolated spermatocyte cysts.
Dissection of adult Drosophila testes
Collect approximately 10–20 newly eclosed male flies on a CO2 pad.
Decapitate fly heads using razor blade and transfer flies to an Eppendorf.
Pipette 400 μL of M3 media into a glass well plate. We recommend using M3 media throughout the procedure. However, Schneiders’s media could be used as an alternate choice.
In a separate well, dispense decapitated flies and, using forceps, hold the fly thorax in such a way that the ventral side of the abdomen faces upward (Figure 2A–C and Video 2).
Immerse the fly in M3 cell culture media and, using another forceps, pinch the distal end of the ventral abdominal skin (Figure 2D and Video 2).
Using forceps, hold on to the male terminalia and slowly pull away, such that the whole male reproductive system is released into the media (Figure 2E, F and Video 2).
Using forceps, separate the testes from the rest of the reproductive system. Drosophila testes can be easily identified by their characteristic spiral shape (Figure 2 G, H and Video 2).
This procedure is repeated until 10–20 pairs of testes are dissected out in M3 cell culture media.
Figure 2. Dissection of testes and germ line cysts. (A) Decapitated male Drosophila flies. (B, C) Position the fly ventral side up and hold the thorax with forceps. (D) Pinch ventral abdominal skin with another forceps. (E) Hold male terminalia with forceps. (F) Slowly pull away male terminalia. (G) Separate the testis from the accessory glands and collecting ducts. (H) Wash the testes with M3 media. (I–M) Under high magnification, focus on individual testes and, using a pair of needles, tear open the testis muscle sheath. (N–O) All the torn open testes are gently agitated to release cysts into the media. (P) Cysts collected at the center of the dissection well.
Video 2. Detailed procedure for Drosophila testes dissection, isolation of individual germline cysts, and mounting on cover glass bottom dish
Isolation of male meiotic spermatocyte cysts
Rinse the testes by replacing old media with fresh M3 media (400 μL each wash, repeat three times).
Cut the pipette tip with scissors and prewet it with M3 media by aspirating in and dispensing out.
Using the precut and prewetted pipette tip, transfer washed testes to a fresh well containing 400 μL of M3 media.
Adjust the SteRio V12 microscope to high magnification (63×) while focusing on the individual testes.
Using a pair of needle holders that carry tungsten needles with a tip diameter of 0.0175 mm, tear open the testis muscle sheath to release all of the differentiating germ cell cysts, including meiotic spermatocyte stages, into the media (Figure 2I–M and Video 2). Repeat this procedure until the rest of the testes are torn open (Figure 2N). Note that this is the most difficult part of the procedure, which could be mastered with practice. We recommend holding needles in a crisscross fashion adjacent to the site of dissection to get a grip on the testis, followed by poking and holding the testis with one pin and tearing with another pin (Figure 2I–M and Video 2).
All the torn open testes are bundled and, by gently agitating with needles, all the cysts that are loosely attached to testes muscle walls will fall into the media. Subsequently, remove all the other testes debris including spermatid bundles and muscle sheath by pushing them out of the media with needles (Figure 2N–O and Video 2).
Gently flush the germ cell cysts settled at the bottom of the glass well plate to the center of the plate by pipetting with fresh media (Figure 2P and Video 2). Wash the cysts concentrated at the center of the well with fresh media by replacing old media with fresh media three times (3× 400 μL). If an experiment requires treatment with a pharmacological agent, replace the old media with fresh media containing the appropriate drug and incubate for the desired time with gentle rocking. As an example, we have treated spermatocyte cysts with Dynasore, a dynamin inhibitor that blocks clathrin-mediated endocytosis [23], at a concentration of 80 μM/mL in M3 media and incubated for 1 h at room temperature with gentle rocking. For the control, we have used a similar volume of the vehicle used to dissolve the drug, in this case DMSO (2.5 μL/mL). As shown in Video 3, compared to control, Dynasore-treated spermatocytes show defects in the assembly of central spindle during early stages of cytokinesis, implicating a crucial role of dynamin activity and perhaps endocytosis in male meiotic cytokinesis.
Video 3. In vitro live imaging of isolated adult Drosophila spermatocyte cysts treated with vehicle DMSO or dynamin inhibitor Dynasore (80 μM/mL).Cysts are isolated from fly testes expressing tGPH (PH domain of Grp1/Step) and Feo-mCherry ubiquitously. The tGPH is predominantly localized to cytosol due to loss of cyst cell membrane envelop. Live imaging was performed with the 63× water immersion objective and an AOI of 1024 × 1024 and at intervals of 2 min for 120 repeats. Video shown here represents 10 frames/second.
Mounting cysts onto a cover glass bottom dish
Prewet a 200 μL pipette tip with M3 media by aspirating in and dispensing out.
Using the prewetted 200 μL pipette tip, aspirate cysts concentrated at the center of the glass well plate and dispense as a drop at the center of the cover glass bottom dish (Figure 3A, B and Video 2). Note: Poly L-Lysine was shown to be detrimental to male germline cysts (Gartner et al. 2014); therefore, we do not recommend coating glass bottom dish with Poly L-Lysine/Poly D-Lysine. In our hands, coating with low concentrations of Poly D-Lysine (50 μg/mL) followed by thorough washing with water did not show much effect on cytokinesis [3]. However, cytokinetic defects are apparent, particularly in the meiosis II cytokinesis, in glass-bottom dishes coated with higher concentrations of poly D-Lysine (200 μg/mL), even after thorough washing (Video 4).
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Figure 3. Mounting male germ line cysts in a cover glass bottom dish and moving onto spinning disk confocal microscope stage. (A) Prewashed male germ line cysts were aspirated with a prewetted pipette tip. (B) Cysts were dispensed at the center of a cover glass-bottom dish (Ø 15 mm). (C) Gently, 200 μL of M3 media was added without disturbing the cysts at the center. (D) Leica DMi8 microscope base setup. (E) Glass bottom dish mounted on a slide/dish holder with a magnetic clamp. A drop of water dispensed on 63× water immersion objective (arrow). (F) Slide/dish holder gently inserted on the microscope stage.
Video 4. In vitro live imaging of dissected adult Drosophila spermatocyte cysts undergoing meiosis I and meiosis II cytokinesis in cover glass-bottom dish coated with poly-D-Lysine (200 μg/mL overnight at room temperature followed by 3× wash with distilled water). Cysts are isolated from fly testes expressing tGPH (PH domain of Grp1/Step) and Feo-mCherry ubiquitously. Live imaging was performed with the 63× water immersion objective and an AOI of 1024 × 1024 and at intervals of 2 min for 120 repeats. Video shown here represents 10 frames/second.
Slowly add 200 μL of the fresh M3 media to the observation area of the cover glass bottom dish without disturbing the cysts from the center (Figure 3C and Video 2). (Note: The volume of media added could differ depending on the observation area of the cover glass bottom dish used in the experiment). Close the dish with the lid and gently move it to the confocal imaging station (Figure 3D and F). Make sure cysts remain at the center of the dish; cysts tend to spread to the corners if the media level is too low or too high.
If desired, cysts can be individually separated by gently pushing each cyst away from the pool with a needle, followed by aspirating it with the prewetted pipette tip and dispensing into a cover glass bottom 94-well plate containing 100 μL of media in each well. Individual cysts settled in each well could be examined under inverted confocal microscope to identify the differentiation stage of interest. This approach allows one to arrest the cyst at the desired differentiation stage by simply adding paraformaldehyde (PFA) to the well (final concentration should be 4%) [3].
Time-lapse imaging on a spinning disc confocal microscope
We use Andor spinning disk confocal microscope with the Leica DMi8 base for imaging; however, the protocol described below could be adapted to any other imaging setup. This microscope has Yokogawa CSU-W1 confocal scanner 8 laser lines and also provides epifluorescence, brightfield, and differential interface (DIC) imaging modalities. It also has a motorized XY stage with piezo Z scanner (ASI), Andor Zyla4.2 + sCMOS camera, and a range of objectives including 5× air, 10× air, 20× air/oil, 63× oil, 25× water, and 63× water (1.2 NA, WD 0.30 mm).
Turn on the spinning disc confocal microscope and the computer.
On the computer, click on ASI console app → select MS2000 → click Connect → set x, y, and z speed to 5 and exit from ASI console.
Double-click on Andor Fusion app to open the software.
Place the glass bottom dish in the slide/Petri dish holder and lock the dish with magnetic clamps (Figure 3D, F).
Return to Fusion software. In the Navigation window, enter image name and save location by clicking on the three dots as shown in Figure 4A with first arrow.
In the Protocol window, if a previously saved protocol exists, select and reload it as shown by the second arrow in Figure 4A.
In the Microscope window (arrow 3), select objective of interest. We normally select 10× air objective first. Go to Active channels window (arrow 4) and select Bright field eyes (BF eyes). Return to the microscope and, using the eyepiece, locate and focus on the isolated cysts (Figure 5A). Use the joystick to move the stage x and y positions and the fine/course adjustment for moving in z axis as desired (Figure 3D).
Return to Fusion. In the Active channels window from the drop-down list (arrow 4), select brightfield camera and press live mode (Figure 4B, arrow 6). Adjust the exposure time to 100 ms (below arrow 4) and adjust the contrast by pressing auto contrast (Figure 4B, arrow 7). Take a snap of the image (Figure 5A) if needed (Figure 4B, arrow 10).
Switch the objective to 63× water immersion on the Microscope window (Figure 4A, arrow 3) and select bright field eyes from the active channels drop-down list (arrow 4). Go to the microscope, gently lift up the slide/Petri dish holder, place one or two drops of double-distilled water on the objective lens, and return the dish holder to its original position (Figure 3D, F). Note that water on the objective tends to evaporate during longer hours of imaging (>4 h). Using brightfield and eyepiece, search for cysts undergoing cytokinesis or spermatocytes that are likely to undergo meiotic cytokinesis within the next 10–15 min. Prophase/metaphase spermatocytes typically have a round nucleus in the brightfield (Figure 5B, C). Prophase spermatocytes have evenly distributed mCherry fascetto in the nucleus (Figure 5E, F) (Video 5). During metaphase, nuclear membrane is ruptured/perforated; as a result, mCherry fascetto translocate from nucleus to cytoplasm (Video 5). Anaphase cells typically appear as oval shaped, with the elongated nucleus surrounded by parafusorial membranes (Figure 5G, H).
Figure 4. Time lapse imaging on Andor spinning disk confocal microscope. Andor Fusion software (2.3.0.48) workstation showing acquisition control, channel manager windows (A), and protocol manager window (B).
Once identified, mark the cyst location of interest on the Multifield window in the Fusion software (Figure 4B, arrow 12 and 16). Find and add up to 2–4 cyst locations to the Multifield window.
Return to the Fusion software and switch to brightfield camera (BFC)/differential interphase mode (DIC) from the active channels menu (Figure 4A, arrow 4) and press live mode (Figure 4B, arrow 6). Locate and position the cysts of interest to the center of the field using the joystick and update the coordinates for each cyst of interest in the multi-position settings (Figure 4B, arrow 16, arrowhead).
Switch from brightfield camera mode to confocal preview mode (select from drop-down menu arrow 4 in Figure 4A) and select the appropriate laser channel. Adjust the laser power between 0.2% and 2% and the exposure time range from 50 to 300 ms to visualize fluorescence in the cysts (arrow 4). Adjust the contrast in Map channels window (Figure 4B, arrow 8 and 9) by setting up minimum (100) and maximum (150).
While in live mode, use fine adjustment on the microscope to scan through the Z-plane and identify the start z-position for each cyst of interest; set it as “0” in z-scan settings menu (Figure 4B, arrow 15) and update it on Multi position window (arrow 16, arrowhead). Add laser channels of interest to protocol channels (Figure 4B, arrow 13); Confocal 488 and Confocal 561 channels are selected in this example (Figure 4B, arrow 13). Select individual protocol channels from the active channels drop-down menu and adjust their laser power and exposure times. For instance, in our experiments, 488 laser power was set to 5% and the exposure time was set to 30 ms for tGPH. Similarly, 561 laser power was set to 10% and the exposure time was set to 30 ms for mCherry fascetto.
Go to Time repeat window (Figure 4B, arrow 14) and setup time-lapse imaging to capture Z-stack of all positions every 2 min for 120 times (or any number of times of your interest).
Go to Z-scan settings (Figure 4B, arrow 15) and set up Z-scan thickness to 41 μm and the optical section step size to 0.5 μm.
Finally, go to the Run protocols window and press the acquire button (Figure 4B, arrow 17) to start imaging. Live data could be viewed in 3D or individual channels in 2D from options in the Image window (Figure 4B, arrow 18 and 19).
Under the above-described conditions, we were able to image three independent cysts simultaneously for up to 4 h without causing major phototoxicity.
Figure 5. Identification of male germline cysts undergoing meiotic cytokinesis. (A) Low magnification brightfield image of isolated cysts (10×). (B, C) Brightfield image of spermatocytes likely to undergo cell division, at high magnification (63×). Note the spermatocytes with characteristic round nucleus in prophase (B) and spermatocytes with elongating nucleus in metaphase, shown with arrow (C). (D, F) Fluorescence image of prophase spermatocyte cyst showing GFP-tagged pleckstrin homology domain of Grp1that binds to phosphatidylinositol 3,4,5 triphosphate (PIP3) and localizes to plasma membrane expressed under the control of tubulin promoter (green) and central spindle binding protein fascetto tagged with mCherry expressed under the control of ubiquitin promoter (red). Note that mCherry-fascetto is uniformly distributed in the nucleus of prophase spermatocytes and, as the cells progress into metaphase, fascetto translocate to cytoplasm. As a result, net fluorescence reduces due to the diffusion of fascetto (arrow). (G-I) Brightfield images of spermatocytes undergoing cytokinesis. (G) Anaphase spermatocytes showing overall elongated cell shape and spindle envelope called parafusorial membranes around the separating chromosomes (arrow). (H) Spermatocytes in early telophase showing cleavage furrow (arrow) (I) Due to incomplete cytokinesis, spermatocytes in late telophase remain inter-connected with each other via cytoplasmic bridges (arrow).
Video 5. Ex vivo live imaging of meiosis I and meiosis II cytokinesis in adult Drosophila testes. GFP-tagged PH domain of Grp1/Step (tGPH) and mCherry-tagged fascetto (feo) are expressed ubiquitously. Live imaging was performed with the 63× water immersion objective with an AOI of 1024 × 1024 and at intervals of 3 min for 100 repeats. Video shown here represents 10 frames/second.
Ex vivo live cell imaging of meiotic cytokinesis in whole mount adult testes
The protocol described in this section focuses on ex vivo imaging of whole mount adult testes; however, we believe this method could be adaptable to testes from all developmental stages and other tissue types of choice such as larval brain.
Prepare breathable membrane dish for imaging
Take 50 mm Lumox® Dish and turn it upside down so that its flat membrane base faces upward (Figure 6C).
Place SecureSeal imaging spacer sticker on the flat membrane base (Figure 6D) and keep it aside.
Dissection of adult testes
Refer to section A1 and Figure 2A–H and Video 2 for instructions on how to dissect adult testes. Microscope setup for dissection is shown in Figure 6A and B.
Mounting adult testes on breathable membrane dish
Using precut and prewetted pipette tip, aspirate 2–3 pairs of dissected testes and dispense at the center of spacer well, as shown in Figure 6E.
Gently place cover glass over the testes on the spacer well without creating an air bubble (Figure 6F).
Using forceps, gently apply pressure evenly around the cover glass (Figure 6G).
Carefully remove excess media between the cover glass and dish membrane using Kimwipe (Figure 6H). Note that excess media around the cover glass will be problematic, especially when water immersion objectives are used, as water tends to enter the space between cover glass and dish membrane by capillary action.
Figure 6. Ex vivo live imaging of male meiotic cytokinesis in adult testes. (A) Zeiss SteREO Discovery V12 microscope for dissection purposes. (B) Tools required for dissection are displayed, which include one pair of forceps, one pair of needle holders with needles, and a 9-well Pyrex glass plate. (C) Upside down view of 50 mm Lumox dish showing gas permeable membrane surface up. (D) SecureSeal imaging Spacers sticker fixed on gas permeable membrane (well diameter 8–9 mm and depth of 0.12 mm). (E) Mount two pairs of testes in a drop of M3 media (20–30 mL). (F) Gently place a coverslip without creating an air bubble. (G) Gently apply pressure around the well on the coverslip so that excess media comes out of the well and the coverslip firmly attaches to the spacer surface. (H) Gently remove excess media around the coverslip with Kimwipe. (I) Andor Spinning Disk confocal with Leica DMi8 microscope base. (J) Microscope stage showing slide/dish holder and a drop of deionized water (50–100 mL) on 63× water immersion objective (arrow). (K) Place 50 mm Lumox dish on slide/dish holder with the coverslip facing the objective and lock with sliders. (L) Push the condenser arm back to its position and, using brightfield, locate and focus on the spermatocyte cysts undergoing meiosis.
Imaging adult testes on spinning disc confocal microscopy
Place a drop of water on the 63× water immersion objective on confocal microscope (Figure 6I–J).
Mount 50 mm Lumox dish in slide/Petri dish holder with the cover glass facing the objective and lock the dish (Figure 6I–L).
Using brightfield, focus on position 3 of the testis (Figure 1A) and identify the cysts likely to undergo cytokinesis as described in section A4i [e.g., in the brightfield, prophase spermatocyte nucleus appears round (Figure 5B and C)]. Once identified, set up image acquisition as described in section A4. Note that cysts tend to move significantly in whole mount adult testes (Video 5); however, increasing area of interest (AOI) to 2048 × 2048 in Zyla window (Figure 4A, arrow 5, arrowhead) or changing the objectives to lower magnification will reduce the chance of meiotic cysts going out of imaging frame (Video 6).
Lumox dish can be reused several times. Remove the cover glass and testes from the Lumox dish by immersing and rinsing the dish in deionized water, followed by wiping the surface with the 70% ethanol.
Video 6. Ex vivo live imaging of meiosis I and meiosis II cytokinesis in adult Drosophila testes. GFP-tagged PH domain of Grp1/Step (tGPH) and mCherry-tagged fascetto (feo) are expressed ubiquitously. Live imaging was performed with the 63× water immersion objective and a pre-defined area of interest size of 2048 × 2048 and at intervals of 3 min for 140 repeats. Video shown here represents 10 frames/second.
Data analysis
Image processing
Andor Fusion saves data in Imaris image file (.ims) in the specified folder. Double-click on the image to open the Imaris software (license required).
When a time series image opens on Imaris with 3D view, adjust bright/contrast by setting up minimum/maximum values for each channel (e.g., min 100, max 150) (Edit → Display Adjustments). To crop time series to remove any unacquired time points or to trim overall time series: Edit → crop time → in a pop-up window, enter start and end time points → OK → Save). Similarly, to trim Z-slices go to Edit → delete slice → in a pop-up window enter which slices to delete → OK → Save. To combine two-time series, open first image series and then go to Edit → add time points → in a pop-up browser window select second.ims file and enter. To crop 3D volume of a time series, go to Edit → crop 3D → in a pop-up window select area to be cropped → OK → Save.
To save time series image as video, press record button (red button left to the time slider in 3D view). A pop-up window will appear; select 10 frames/sec and H264 Video (*.mp4) format.
To further process time series in ImageJ/FIJI, press record button → in a pop-up window select 10 frames/sec and Raw Video (*.avi) file format and save.
Measure cleavage furrow diameter at different time points
Open Raw Video.avi file on ImageJ/FIJI (Figure 7A).
To set up scale bar, draw a straight line on top of image scale bar using straight/segmented or freehand lines or arrows tool (Figure 7A), and go to Analyze → Set Scale → in a pop-up window enter the known distance and unit of length (e.g., “um” for micrometer) → Ok.
Identify the meiotic spermatocyte whose cleavage furrow diameter is to be determined. Using the straight-line tool, draw a line at the equatorial region of the elongated spermatocyte where central spindle will assemble (Figure 7B, C) and go to Analyze → Measure (in a pop-up window, the length of the furrow will be displayed, Figure 7D). Repeat the length measurement for each timepoint until the cleavage furrow reaches its minimum. Measure furrow length from at least three meiotic spermatocytes from three independent cysts.
Plot the cleavage furrow length/diameter as a function of time using XY, column, and statistical analysis in Prism 9. We have taken Dynasore- and DMSO-treated spermatocyte cysts as an example and determined the cleavage furrow diameter as a function of time, shown in Figure 7E, F.
Figure 7. Measurement of cleavage furrow diameter at various time points . (A) Fiji/ImageJ software, showing straight line tool. (B, C) Example images with a straight line (yellow line) drawn at the equatorial region of spermatocyte undergoing cytokinesis (shown with arrow). (D) Example length measurement results shown for the first five-time frames of the video. (E) Comparison of cleavage furrow diameter as a function of time in spermatocytes treated with either vehicle alone (DMSO) or endocytosis inhibitor Dynasore (80 μM). Data represents length measurements from six spermatocytes from three independent cysts. (F) Comparison of furrow diameter at single timepoint (40 min). Each dot in the graph represents a single spermatocyte in a cyst undergoing cytokinesis. Spermatocytes from three independent cysts were used in analysis.
General notes and troubleshooting
General notes
The protocol for in vitro and ex vivo live imaging of the male meiotic cytokinesis described here can be applied to larval and pupal testes.
Water on the immersion objectives tends to dry out during imaging. When imaging for more than 4 h, pause the acquisition, add a drop of water to the objective, and resume the acquisition. However, special care should be taken to prevent the movement of cysts during this process.
The movement of male germ line cysts sets one of the limitations of this protocol, as they are not attached to the cover glass bottom dish and tend to go out of the imaging frame. Frequent monitoring of image acquisition may help prevent this possibility.
Dissected germ line cysts are sensitive to phototoxicity, and use of higher laser powers (>20% with longer exposure times >50 ms) may induce cytokinetic defects in control spermatocytes. Therefore, imaging samples with weaker fluorescence signals will be a challenge. Further, imaging cysts for longer than 6 h will be a problem, as the water in the media tends to evaporate and condense on the lid surface. As a result, the net concentration of media constituents will change, which in turn may affect cytokinesis.
Media conditions greatly affect the germ line cyst behavior during cytokinesis. Do not use the media if it is too old (>2 weeks from the time of thawing an aliquot). Measure the cytokinetic efficiency for the control cysts before going ahead with the treatment conditions.
Whole mount adult testes are relatively less sensitive to phototoxicity compared to dissected germ line cysts. Therefore, imaging whole mount adult testes will be of choice for the samples with weaker fluorescence signals; for the samples with strong fluorescence signals, it is possible to image for longer hours (we were able to image testes up to 7 h without causing any damage, see Video 6). Further, it will be of choice if the sample size is small; for instance, there are <5 male flies available to study cytokinesis. However, significant movement of cysts due to testis muscle contractions will be a limitation. This problem could be counteracted by increasing the image frame size (2024 × 2024) to cover a larger imaging area.
Ex vivo live imaging cannot be performed on wild-type testes containing yellow pigment cells. However, individual germ line cysts could be dissected from these testes and imaged in vitro.
Troubleshooting
Problem 1: Dissected testes look abnormal in the media.
Possible causes: Media pH or osmolarity may not be optimal, or media may be too old.
Solution: Make sure potassium bicarbonate is added during media preparation. Thaw a fresh aliquot of media and warm it to room temperature before using it for dissection.
Problem 2: Germ line cyst count is low.
Possible causes: Old flies were used for dissection; a smaller number of testes were used; or the testes’ muscle sheath was not fully torn open.
Solution(s): Use newly enclosed flies (<24 h) for testes dissection, increase testes number (15–20 pairs), and make sure the testes’ muscle sheath is fully torn to release all the cysts.
Problem 3: Testes or germ line cysts attached to pipette tip walls.
Possible causes: The pipette tip was not prewetted with M3 media containing 10% FBS.
Solution: Prewet the pipette tip with M3 media before aspirating testes or germ line cysts.
Problem 4: Cysts spread out in a cover glass bottom dish.
Possible causes: Media in the observation area of the cover glass bottom dish is not optimal, or the dish agitated excessively while shifting from the dissection area to the confocal imaging stage.
Solution: Make sure the media level in the observation area is flat (should not be convex or concave). In our experiments, a cover glass bottom dish with Ø15 mm area takes approximately 150–200 μL of media. Once cysts are mounted in the cover glass bottom dish, take extra care to prevent excessive movement. For instance, gently put the dish in a small tray and walk slowly while transporting it to the microscope for imaging. When all the cysts remain at the center, it will take much less time and increase the chances of finding the cysts undergoing cytokinesis.
Problem 5: Cysts move while imaging.
Possible causes: Excessive vibration on the microscope stage or media level is too high in the dish.
Solutions: Make sure the confocal microscope unit is on an anti-vibration table; avoid touching the microscope while imaging; make sure the media level in the observation area of the dish is flat (refer to Problem 4).
Problem 6: Difficulty in finding cysts undergoing cytokinesis.
Possible cause: Cysts were isolated from >2-day-old fly testes; or a low number of testes were used for dissection; or cyst count was low; or the cysts were spread out in the cover glass bottom dish.
Solutions: Use newly eclosed flies (<24 h) for dissecting testes; increase the number of testes (15–20 pairs); make sure the pipette tip is prewet when aspirating and dispensing cysts; make sure cysts remain at the center of the dish.
Problem 7: Cytokinetic defects seen in control spermatocytes.
Possible cause: Something went wrong with the media; the media is too old; a poly-L-lysine-coated cover glass bottom dish was used; higher laser powers were used for imaging.
Solutions: Make sure M3 media is prepared as described; discard old media and prepare fresh from frozen aliquots; make sure the cover glass bottom dish is not treated with high concentrations of poly L-Lysine; reduce laser power to image (preferably keep it to less than 10%).
Problem 8: Testes’ muscle sheath moves a lot while imaging ex vivo whole mount adult testes.
Possible cause: Testes are not properly mounted on a gas permeable dish; there is too much media around the cover glass and the spacer or water on the immersion objective entered the space between the cover glass and membrane.
Solutions: Make sure the cover glass is firmly sitting on the spacer and remove any excess media around the cover glass and membrane. Normally, testis muscle from newly eclosed flies tends to contract more than 2–3-day-old flies. If the problem persists, it may be a good idea to test testes from 2–3-day-old flies for imaging. However, in older flies, cysts undergoing cytokinesis are lower than the newly eclosed flies.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Kunduri et al. (2022). Delivery of ceramide phosphoethanolamine lipids to the cleavage furrow through the endocytic pathway is essential for male meiotic cytokinesis. PLOS BIOLOGY (Figure 4, panel C, D).
Acknowledgments
We thank Valentin Magidson (Staff, Optical Microscopy and Analysis Laboratory at NCI-Frederick) for technical assistance in imaging on Andor spinning disk confocal microscope. This study was funded by the intramural division of the National Cancer Institute, National Institutes of Health, Department of Health and Human Services. The content of this publication does not necessary reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. Government.
Competing interests
The authors declare no conflicts of interest.
References
Frappaolo, A., Piergentili, R. and Giansanti, M. G. (2022). Microtubule and Actin Cytoskeletal Dynamics in Male Meiotic Cells of Drosophila melanogaster. Cells 11(4): 695. doi: 10.3390/cells11040695
Kunduri, G., Acharya, U., Acharya, J. K. Lipid Polarization during Cytokinesis. (2022) Cells 11(24): 3977. doi: 10.3390/cells11243977
Kunduri, G., Le, S. H., Baena, V., Vijaykrishna, N., Harned, A., Nagashima, K., Blankenberg, D., Yoshihiro, I., Narayan, K., Bamba, T., et al. (2022). Delivery of ceramide phosphoethanolamine lipids to the cleavage furrow through the endocytic pathway is essential for male meiotic cytokinesis. PLoS Biol. 20(9): e3001599. doi: 10.1371/journal.pbio.3001599
Adams, R. R., Tavares, A. A., Salzberg, A., Bellen, H.J. and Glover, D. M. (1998). pavarotti encodes a kinesin-like protein required to organize the central spindle and contractile ring for cytokinesis. Genes Dev. 12(10): 1483–1494. doi: 10.1101/gad.12.10.1483
Prokopenko, S. N., Brumby, A., O’Keefe, L., Prior, L., He, Y., Saint, R. and Bellen, H. J. (1999). A putative exchange factor for Rho1 GTPase is required for initiation of cytokinesis in Drosophila. Genes Dev. 13(17): 2301–2314. doi: 10.1101/gad.13.17.2301
Gatti, M. and Baker, B. S. (1989). Genes controlling essential cell-cycle functions in Drosophila melanogaster. Genes Dev. 3(4): 438–453. doi: 10.1101/gad.3.4.438
Giansanti, M. G., Gatti, M. and Bonaccorsi, S. (2001). The role of centrosomes and astral microtubules during asymmetric division of Drosophila neuroblasts. Development 128(7): 1137–1145. doi: 10.1242/dev.128.7.1137
Giansanti, M. G., Bonaccorsi, S., Williams, B., Williams, E. V., Santolamazza, C., Goldberg, M. L. and Gatti, M. (1998). Cooperative interactions between the central spindle and the contractile ring during Drosophila cytokinesis. Genes Dev. 12(3): 396–410. doi: 10.1101/gad.12.3.396
Gunsalus, K. C., Bonaccorsi, S., Williams, E., Verni, F., Gatti, M. and Goldberg, M. L. (1995). Mutations in twinstar, a Drosophila gene encoding a cofilin/ADF homologue, result in defects in centrosome migration and cytokinesis. J Cell Biol. 131(5): 1243–1259. doi: 10.1083/jcb.131.5.1243
Cenci, G., Bonaccorsi, S., Pisano, C., Verni, F. and Gatti, M. (1994). Chromatin and microtubule organization during premeiotic, meiotic and early postmeiotic stages of Drosophila melanogaster spermatogenesis. J. Cell Sci. 107(12): 3521–3534. doi: 10.1242/jcs.107.12.3521
Giansanti, M. G., Bonaccorsi, S., Bucciarelli, E. and Gatti, M. (2001). Advances in Cytokinesis Research. Drosophila Male Meiosis as a Model System for the Study of Cytokinesis in Animal Cells. Cell Struct. Funct. 26(6): 609–617. doi: 10.1247/csf.26.609
Carmena, M., Riparbelli, M. G., Minestrini, G., Tavares, Ã. M., Adams, R., Callaini, G. and Glover, D. M. (1998). Drosophila Polo Kinase Is Required for Cytokinesis. J Cell Biol. 143(3): 659–671. doi: 10.1083/jcb.143.3.659
Herrmann, S., Amorim, I. and Sunkel, C. E. (1998). The POLO kinase is required at multiple stages during spermatogenesis in Drosophila melanogaster. Chromosoma 107: 440–451. doi: 10.1007/pl00013778
Wakefield, J. G., Bonaccorsi, S. and Gatti, M. (2001). The Drosophila Protein AspIs Involved in Microtubule Organization during Spindle Formation and Cytokinesis. J Cell Biol.153(4): 637–648. doi: 10.1083/jcb.153.4.637
Giansanti, M. G., Farkas, R. M., Bonaccorsi, S., Lindsley, D. L., Wakimoto, B. T., Fuller, M. T. and Gatti, M. (2004). Genetic Dissection of Meiotic Cytokinesis in Drosophila Males. Mol. Biol. Cell 15(5): 2509–2522. doi: 10.1091/mbc.e03-08-0603
Bonaccorsi, S., Giansanti, M. G., Cenci, G., Gatti, M., Sullivan, W., and Ashburner, M. (2000). Cytological analysis of spermatocyte growth and male meiosis in Drosophila melanogaster. Drosophila Protoc.: 87–109.
Cross, D. P. and R, D. L. S. (1979). The dynamics of Drosophila melanogaster spermatogenesis in in vitro cultures. Development 53(1): 345–351. doi: 10.1242/dev.53.1.345
Gärtner, S. M. K., Rathke, C., Renkawitz-Pohl, R. and Awe, S. (2014). Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment. J. Vis. Exp. (91): e51868. doi: 10.3791/51868
Karabasheva, D. and Smyth, J. T. (2020). Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue. J. Vis. Exp. (159): e60961. doi: 10.3791/60961
Nelson, K. A., Warder, B. N., DiNardo, S. and Anllo, L. (2020). Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad. J. Vis. Exp. (164): e61872. doi: 10.3791/61872
Sitaram, P., Hainline, S. G. and Lee, L. A. (2014). Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes. J. Vis. Exp. (83): e51058. doi: 10.3791/51058
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Giansanti, M. G., Sechi, S., Frappaolo, A., Belloni, G. and Piergentili, R. (2012). Cytokinesis in Drosophila male meiosis. Spermatogenesis 2(3): 185–196. doi: 10.4161/spmg.21711
Supplementary information
The following supporting information can be downloaded here:
Figure S1: Round spermatids from type and Ceramide phosphoethanolamine synthase (cpes) mutant testes
Article Information
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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4,919 | https://bio-protocol.org/en/bpdetail?id=4919&type=0 | # Bio-Protocol Content
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The Development of an Advanced Model for Multilayer Human Skin Reconstruction In Vivo
MP Maryna Pavlova
VB Velmurugan Balaiya
JF Jocelyn C. Flores
MF Michael Ferreyros
KB Katie Bush
AH Amy Hopkin
IK Igor Kogut
DR Dennis R. Roop
GB Ganna Bilousova
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4919 Views: 954
Reviewed by: Pilar Villacampa AlcubierreEVANGELOS THEODOROU Anonymous reviewer(s)
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Abstract
Human skin reconstruction on immune-deficient mice has become indispensable for in vivo studies performed in basic research and translational laboratories. Further advancements in making sustainable, prolonged skin equivalents to study new therapeutic interventions rely on reproducible models utilizing patient-derived cells and natural three-dimensional culture conditions mimicking the structure of living skin. Here, we present a novel step-by-step protocol for grafting human skin cells onto immunocompromised mice that requires low starting cell numbers, which is essential when primary patient cells are limited for modeling skin conditions. The core elements of our method are the sequential transplantation of fibroblasts followed by keratinocytes seeded into a fibrin-based hydrogel in a silicone chamber. We optimized the fibrin gel formulation, timing for gel polymerization in vivo, cell culture conditions, and seeding density to make a robust and efficient grafting protocol. Using this approach, we can successfully engraft as few as 1.0 × 106 fresh and 2.0 × 106 frozen-then-thawed keratinocytes per 1.4 cm2 of the wound area. Additionally, it was concluded that a successful layer-by-layer engraftment of skin cells in vivo could be obtained without labor-intensive and costly methodologies such as bioprinting or engineering complex skin equivalents.
Key features
• Expands upon the conventional skin chamber assay method (Wang et al., 2000) to generate high-quality skin grafts using a minimal number of cultured skin cells.
• The proposed approach allows the use of frozen-then-thawed keratinocytes and fibroblasts in surgical procedures.
• This system holds promise for evaluating the functionality of skin cells derived from induced pluripotent stem cells and replicating various skin phenotypes.
• The entire process, from thawing skin cells to establishing the graft, requires 54 days.
Graphical overview
Generation of a human skin equivalent on an immunodeficient mouse using a fibrin-based grafting system. A schematic of the protocol is shown. Cultured keratinocytes and fibroblasts resuspended in a fibrin-based gel are delivered as layers into a silicon chamber inserted underneath the skin of an immunocompromised mouse. First, a fibrin gel containing encapsulated fibroblasts (up to 2 × 106 per 1.4 cm2 wound) is delivered into the chamber and allowed to solidify for 15 minutes. Second, a fibrin gel containing 1.0–2.0 × 106 keratinocytes is applied on top of the fibroblast layer. On day 7 post-grafting, the chamber is removed, and the wound with the graft is allowed to heal for 4–5 weeks. During healing, a scab forms and eventually falls off. By day 54, the graft is fully established.
Keywords: Human skin equivalent Fibrin-based hydrogel Multilayered skin graft In vivo skin model Regenerative medicine
Background
Progress in mouse genetics has provided a powerful research tool for elucidating the mechanisms of epidermal stem cell commitment and skin homeostasis. However, given the dramatic difference in skin structure between humans and mice, one must be cautious in applying the insights gained in studying mouse skin to human skin, especially when investigating skin diseases. For example, the inbred genetic background of mice can influence the phenotype resulting from disease-associated mutations. Humans are of mixed genetic background, and this complexity results in phenotypical variations and a different penetrance of genetically defined diseases, such as epidermolysis bullosa, a group of rare inherited skin blistering diseases. For this reason, developing complex in vivo human xenograft models is critical to advance our understanding of human skin conditions and developing new therapeutic interventions.
Several human xenograft models have been previously published, including a skin flap assay (Qiao et al., 2008), human skin transplantation-like approaches for grafting dermal-epidermal equivalents onto opened wounds of recipient mice (Escamez et al., 2004; Martinez-Santamaria et al., 2012; Yanez et al., 2015; Jorgensen et al., 2020), and chamber grafting assay (Wang et al., 2000; Diette et al., 2020). While promising, all these assays suffer from several limitations. They include (1) the complexity of the procedure that requires the generation of 3D skin equivalents in cell culture conditions before the transplantation onto a mouse, and (2) the limited availability and proliferative capacity of primary human cells, especially cells derived from patients with rare inherited skin diseases. A chamber grafting assay (Wang et al., 2000; Diette et al., 2020) provides a simplified procedure for grafting primary human skin cells since the assay promotes the self-assembly of fibroblasts and keratinocytes in an in vivo environment without the need to produce skin equivalents in a dish. However, successful production of human skin equivalents in the grafting assay still requires at least 5 × 106 primary keratinocytes per 1.4 cm2 of the wound area, which is a significant number when the availability of primary patient cells is limited. In addition, since this assay relies on the ability of keratinocytes and fibroblasts to self-assemble and form the dermis and epidermis, this limits the ability to incorporate other cell types that normally reside in the skin into an appropriate skin layer. Therefore, we aimed to modify the grafting chamber assay by decreasing the number of primary skin cells necessary for transplantation and mimicking the natural three-dimensional structure of the human skin in vivo to make a more viable in vivo model to study human skin conditions.
To achieve this, we modified the delivery of human keratinocytes and fibroblasts into the grafting chamber by first suspending these cells in a fibrin-based hydrogel. We then performed the sequential transplantation of fibroblasts followed by keratinocytes to recapitulate two major layers of the skin, the dermis and the epidermis, in a controlled manner. Due to its simplicity, this method can be easily optimized by incorporating additional cell types of different origins and ensuring that they are targeted explicitly into the dermis or the epidermis. In addition to grafting primary keratinocytes and fibroblasts, this system can be appropriate for testing the functionality of skin cells derived from induced pluripotent stem cells. The system is also applicable for testing novel skin transplantation systems for potential clinical applications, including cell harvesting and application techniques for treating cutaneous skin defects and pigmentation disorders.
Materials and reagents
Biological materials
Nude mice as recipients for grafting: homozygous nude Foxn1nu, formerly Hfh11nu, females, 6–8 weeks (Jackson Laboratory, Nu/J, #002019; https://www.jax.org/strain/002019). Alternatively, the homozygous NOD-Prkdcscid mice can be recipients for grafting (Jackson Laboratory, NOD.Cg-Prkdcscid/J, # 001303). Both nude and NOD-SCID mouse strains show efficient engraftment of human cells. However, NOD-SCID mice require depilation on the back before surgery and often before graft harvest. Therefore, careful consideration should be given to selecting the appropriate mouse strain for immunological studies to ensure compatibility with the experimental requirements and to minimize any potential confounding factors related to hair removal (Waldron-Lynch et al., 2012; Cristobal et al., 2021).
Note: Before preparing cell cultures, ensure that all mice are healthy and fully acclimated to the facility for at least a week. Ensure that the weight of the mice is above 21 g.
Fibroblasts, primary, neonatal (ATCC, catalog number: PCS-201-010)
Human epidermal keratinocytes, primary, neonatal (HEKn) (ThermoFisher, catalog number: C0015)
Reagents
DMEM/F12 (1:1) (ThermoFisher, catalog number 11320-033)
Pooled human AB serum derived (Innovative Research, ISERAB)
MEM-NEAA (ThermoFisher, catalog number: 11140050
GlutaMax (ThermoFisher, catalog number: 35050061)
2-Mercaptoethanol (ThermoFisher, catalog number: 21985023)
L-Ascorbic acid (Sigma-Aldrich, catalog number: A4544)
Hydrocortisone (Sigma-Aldrich, catalog number: H0888)
Antibiotic/Antimycotic (ThermoFisher, catalog number: 15240062)
Human bFGF (ThermoFisher, catalog number: PHG0263)
Human EGF (ThermoFisher, catalog number: PHG0313)
EpiLife medium (ThermoFisher, catalog number:MEPI500CA)
EpiLife defined growth supplement (EDGS) (ThermoFisher, catalog number: S0125)
Bovine collagen solution, type I, 3 mg/mL (Advanced BioMatrix, catalog number 5005).
Fetal bovine serum, qualified, one shot, raw (ThermoFisher, catalog number: A31605-01)
Aprotinin (Sigma-Aldrich, catalog number: A6279-10ML)
Fibrinogen from human plasma (Sigma-Aldrich, catalog number: F3879-1G)
Thrombin from human plasma (Sigma-Aldrich, catalog number: T4393-100UN)
DPBS without calcium and magnesium (ThermoFisher, catalog number: 14190-144)
0.25% Trypsin-EDTA (ThermoFisher, catalog number: 25200-056)
Accutase (StemCell, catalog number: 7920)
CRYO defined, animal component free freezing medium, 2× (ZenBio, catalog number: CNT-CRYO-50)
CryoStor CS10, 1× (StemCell, catalog number: 100-1061)
Anti-mouse keratin (K)1 antibody, Rabbit, 1:500 dilution (BioLegend, catalog number: 905602)
Anti-mouse/human K14 antibody, Chicken, 1:2,000 dilution (BioLegend, catalog number: 906004)
Anti-human Loricrin antibody, Rabbit, 1:2,000 dilution (Abcam, catalog number: 176322)
Anti-human Vimentin antibody, Mouse, 1:500 dilution (Abcam, catalog number: 16700)
Anti-Rabbit-488 secondary antibody (Invitrogen, catalog number: A11008)
Anti-Chicken-594 secondary antibody (Invitrogen, catalog number: A11042)
Anti Mouse-594 secondary antibody (Invitrogen, catalog number: 11032)
Formalin solution, neutral buffered, 10% (Sigma-Aldrich, catalog number: HT501128)
Solutions
Stock solutions
Thrombin stock solution (25 U/mL): made from powder (see Recipes)
Fibrinogen stock solution (40 mg/mL): made from powder (see Recipes)
Bovine collagen solution, type I (3 mg/mL): ready to use
Working solutions
Complete FEM medium (fibroblasts culturing) (see Recipes)
Complete EpiLife medium (keratinocytes culturing) (see Recipes)
Collagen solution (coating for keratinocyte culture) (see Recipes)
Fibrin gel containing fibroblasts (see Recipes)
Fibrin gel containing keratinocytes (see Recipes)
Recipes
Recipes for stock solutions
Thrombin stock solution
Working under aseptic conditions in a biosafety cabinet, reconstitute thrombin in sterile DPBS (without Ca2+ and Mg2+) at a concentration of 25 U/mL. Make an aliquot and freeze at -20 °C. The day before the surgery, defrost the thrombin solution overnight at 2–8 °C. Keep thrombin on ice before applying it for hydrogel preparation in the grafting procedure. The solution should be used promptly; however, it can be refrigerated at 2–8 °C for up to three days.
Fibrinogen stock solution
Working under aseptic conditions in a biosafety cabinet, reconstitute fibrinogen in sterile DPBS (without Ca2+ and Mg2+) at a concentration of 40 mg/mL stepwise. First, lay a thin layer of fibrinogen powder on top of warm (37 °C) DPBS and let it soak for 5 min at 37 °C; repeat the soaking procedure with the rest of the fibrinogen. Second, put the tube with the fibrinogen-saline solution in a water bath at 37 °C for 30–60 min and agitate the tube gently every 10 min. Ensure that there are no clumps of dry fibrinogen on the tube walls. Third, leave the tube on the shaker at 2–8 °C overnight to complete the preparation of the fibrinogen stock solution. If a significant number of undissolved clumps are left, repeat the 30-min incubation at 37 °C. Do not exceed 1 h of incubation at 37 °C. The fibrinogen-saline solution must not be vortexed. Fibrinogen should be filter-sterilized using a 0.22 μm syringe filter. Do not use vacuum filtration since this will lead to the breakdown of the molecule during the filtration. Make 50–100 µL aliquots and freeze at -20 °C.
Note: Alternatively, bovine fibrinogen and thrombin can be used.
Recipes for working solutions
Complete FEM medium (Table 1)
Prepare FEM medium by supplementing DMEM/F12 with 5% human serum, 0.5× MEM-NEAA, 0.5× GlutaMAXTM supplement, 55 µM GibcoTM 2-mercaptoethanol, 1× HyClone Antibiotic/Antimycotic, 50 µg/mL ascorbic acid, and 1 µg/mL hydrocortisone. Perform 0.22 µm sterile filtration and store at 4 °C for up to one month. Before application, add human bFGF to a final concentration of 12 ng/mL and human EGF to a final concentration of 5 ng/mL.
Table 1. Composition of complete FEM medium
Reagent Stock concentration Final concentration Amount
DMEM/F12 (1:1) 92.763 mL
Human serum 100% 5% 5 mL
MEM-NEAA 100× 0.5× 0.5 mL
GlutaMax Supplement 100× 0.5× 0.5 mL
2-Mercaptoethanol 55 mM 55 µM 0.1 mL
Ascorbic acid 50 mg/mL 50 µg/mL 0.1 mL
Hydrocortisone 5 mg/mL 1 µg/mL 0.02 mL
Antibiotic/Antimycotic 100× 1× 1 mL
1. Sterilize using a 0.22 µm filter; 2. Add bFGF and EGF as shown below AFTER filtration
Reagent Stock concentration Final concentration Amount
bFGF 100 µg/mL 12 ng/mL 0.012 mL
EGF 100 µg/mL 5 ng/mL 0.005 mL
Total n/a n/a 0.005 mL
Complete EpiLife medium (Table 2)
Prepare complete EpiLife medium by adding EpiLife Defined Growth Supplement (EDGS) following the manufacturer’s instruction. Add antibiotic/antimycotic.
Table 2. Composition of complete EpiLife Medium
Reagent Stock concentration Final concentration Amount
EpiLife basal medium n/a n/a 500 mL
EDGS 100× 1× 5 mL
Antibiotic/Antimycotic 100× 1× 5 mL
Total n/a n/a 510 mL
Collagen solution
To prepare the collagen working solution for the coating of the tissue culture dishes, dilute the 3 mg/mL bovine collagen stock solution to a final working concentration of 30 µg/mL in 1× DPBS. Use fresh.
Fibrin gel containing fibroblasts (Table 3)
To prepare the fibrin gel containing fibroblasts, first make working Solution 1. To make working Solution 1, supplement DMEM/F12 1:1 with 1× Antibiotic/Antimycotic, 1% raw FBS, and aprotinin. Mix components in an Eppendorf tube and keep the tube on ice until ready to graft. Just before grafting, mix Solution 1 with fibroblasts and then add fibrinogen and thrombin. Mix components and use for grafting.
Critical: Do not add fibroblasts, fibrinogen, and thrombin into Solution 1 until ready to graft*.
Table 3. Composition of the gel containing fibroblasts
Reagent Stock concentration Final concentration Amount
Solution 1
DMEM/F12 (1:1) n/a n/a 291.7 µL
Antibiotic/Antimycotic 100× 1× 4 µL
Raw FBS 100% 1% 4.1 µL
Aprotinin (3-7 TIU/mg protein) n/a 23.5 µL
Just before grafting, add fibroblasts, fibrinogen, and thrombin as shown below.
Reagent Stock concentration Final concentration Amount
Fibroblasts* n/a 2 × 106 50 µL
Fibrinogen* 40 mg/mL 1.73 mg/mL 17.3 µL
Thrombin* 25 U/mL 0.59 U/mL 9.4 µL
Total n/a n/a 400 µL
Fibrin gel containing keratinocytes (Table 4)
To prepare the fibrin gel containing keratinocytes, first make working Solution 2. To make a working Solution 2, supplement EpiLife medium with 1× Antibiotic/Antimycotic and aprotinin. Mix components in an Eppendorf tube and keep the tube on ice until ready to graft. Just before grafting, mix Solution 2 with keratinocytes and then add fibrinogen and thrombin. Mix components and use for grafting.
Critical: Do not add keratinocytes, fibrinogen, and thrombin into Solution 2 until ready to graft**.
Table 4. Composition of the gel containing keratinocytes
Reagent Stock concentration Final concentration Amount
Solution 2
EpiLife n/a n/a 295.8 µL
Antibiotic/Antimycotic 100× 1× 4 µL
Aprotinin (3-7 TIU/mg protein) n/a 23.5 µL
Just before grafting, add keratinocytes, fibrinogen, and thrombin as shown below.
Reagent Stock concentration Final concentration Amount
Keratinocytes** n/a 2 × 106 50 µL
Fibrinogen** 40 mg/mL 1.73 mg/mL 17.3 µL
Thrombin** 25 U/mL 0.59 U/mL 9.4 µL
Total n/a n/a 400 µL
Note: For a large-scale experiment, the working hydrogel solutions for the dermal and epidermal layers can be prepared as a combined mix to facilitate the generation of multiple grafts simultaneously. To maximize efficiency, 2–4 grafts can be performed in a single experimental setup. Optional: recalculate the volumes for Solution 1 and Solution 2 based on the number of grafts.
Equipment
Silicone chambers [produced by Qure Medical, 1810 Renaissance Boulevard, Sturtevant, WI, 53177, (262) 417–1307, http://www.qure-med.com, according to the following dimensions: 12 mm inner diameter (ID), 20 mm outer diameter (OD), 10 mm tall (Part# 12–24, upper chamber)]. Chambers are made of Momentive LSR 2650, two-component liquid silicone rubber for injection molding processes (Figure 2A).
Note: The chambers are reusable and autoclavable. Manually punch three holes in the dome of the chamber using a 2 mm biopsy punch to allow for gas exchange during the time the chamber is in place under the skin.
Nikon Eclipse 90i + Photometrics Cool SNAP HQ2+ DS Fi1 (Nikon, model: Ti2-E)
Nikon eclipse Ti-U inverted microscope (Nikon, model: Ti-U)
CO2 incubator (ThermoFisher, model: HERA cell VIOS 160)
Straight forceps (VWR, catalog number: 82027-398)
Surgical scissors (VWR, catalog number: 82027-584)
Curved forceps (VWR, catalog number: 89259-946)
Pipettes P1000 (Gilson, catalog number: F144059M)
Pipettes P200 (Gilson, catalog number: F144058M)
Pipettes P20 (Gilson, catalog number: F144056M)
Sterile filtration system (Corning, catalog number: 431097)
-80 °C freezer (ThermoFisher, model: TSX Series with V-Drive)
Heating pad-T/Pump Professional (Gaymar, catalog number: TP700)
Heating pad-blanket (Cincinnati Sub-Zero, model: 274)
Liquid bandage New skin (Amazon, catalog number: 851409007011)
Software and datasets
NIS-elements Nikon (https://www.nikoninstruments.com/Products/Software)
Microsoft Excel (Microsoft, https://products.office.com/en-us/excel)
GraphPad Prism (GraphPad, https://www.graphpad.com/scientific-software/prism/)
Adobe Illustrator (https://www.adobe.com/products/illustrator/free-trial-download.html)
Procedure
The generation of complete grafts in this protocol spans 54 days and involves several distinct steps, as depicted in Graphical overview. In Step 1, lasting from Days 1 to 12, cryopreserved keratinocytes and fibroblasts are thawed, cultivated, and subcultured. Step 2, which occurs on Day 12, entails the surgical insertion of a silicon chamber and the establishment of cell-seeded fibrin hydrogels. In Step 3, which takes place on Day 19, the silicon chamber is removed to promote the adherence of the graft to the surrounding tissue. Finally, in Step 4, spanning Days 19–54, the graft establishes and undergoes maturation.
The described protocol outlines the process for conducting a single surgery.
Defrosting and plating keratinocytes
Coat two 10 cm tissue culture dishes with collagen.
Dilute bovine collagen stock solution (3 mg/mL) to a final working concentration of 30 µg/mL in 1× DPBS. For each 10 cm dish, resuspend 60 µL of the collagen solution in 6 mL of 1× DPBS in a 15 mL conical tube. Pipette to mix.
Add 6 mL of diluted collagen solution to each 10 cm (~60 cm2) tissue culture dish. Rock back and forth to ensure complete coverage.
Place into the 37 °C/5% CO2 incubator for 1 h.
Prepare complete EpiLife medium (see Recipes).
Prewarm a 25 mL aliquot of complete EpiLife medium to room temperature (RT).
Aspirate non-polymerized collagen from the 10 cm dishes prepared in step A1. Immediately add 9 mL of complete EpiLife medium and transfer the dishes into a 37 °C/5% CO2 tissue culture incubator to equilibrate for 30 min.
Pipette 5 mL of EpiLife into a 15 mL conical tube. Set aside in a biosafety cabinet.
Remove a cryovial that contains approximately 5 × 105 frozen keratinocytes from liquid nitrogen storage and immediately place it into a 37 °C water bath. Thaw quickly (approximately 1–2 min), avoiding contamination by keeping the cap above the water level.
Note: The thawing time of frozen cells depends significantly on the volume of the cells being thawed. After one minute of thawing, gently shake the vial and quickly observe the contents. If most of the cell suspension has melted, you can proceed with the next step.
Spray the entire vial with 70% ethanol, wipe, and proceed under strict aseptic conditions in a biosafety cabinet.
Transfer thawed keratinocytes into the conical tube prepared in step A5.
Centrifuge the cells at 200× g for 5 min at RT.
Gently resuspend the pellet in 1 mL of complete EpiLife medium and transfer 500 µL of the cell suspension with 2.5 × 105 keratinocytes into each 10 cm dish with the pre-equilibrated EpiLife medium prepared in step A4, to reach a proper/required plating density of 4 × 103 cells/cm2.
Gently and thoroughly disperse the cells by alternating between an up/down and then left/right motion. Repeat these motions two more times to ensure even distribution. Do not swirl the plate to mix. Once the cells are evenly dispersed, place the plate into a 37 °C/5% CO2 tissue culture incubator.
The next day, check cell attachment under a microscope.
Change the complete EpiLife medium every other day until cells reach approximately 60% confluency, then change daily until keratinocyte culture reaches 70%–80% confluency, at which point the cells must be passaged.
Note: If plated correctly at the density of 2.5–4 × 103/cm2, the culture should be ready for subculturing in 5–7 days.
Keratinocytes must be subcultured to perform better during grafting (see below). The rest of the cells can be frozen for future experiments.
Subculturing keratinocytes
Note: Two 10 cm dishes with keratinocytes at 80% confluency yield approximately 2.4 × 106 cells (see Figure 1A). Since 2.0 × 106 keratinocytes are needed per surgery, two 10 cm dishes are sufficient to obtain this number of cells.
Figure 1. Representative picture of keratinocyte and fibroblast cultures ready to harvest for grafting. Keratinocytes (A) and fibroblasts (B) are shown at the appropriate confluency for grafting. Scale bar, 50 µm.
Allow Accutase and complete EpiLife medium to equilibrate to RT.
Prepare two 10 cm tissue culture dishes and coat them with bovine collagen following the procedure described in step A1. Place into a 37 °C/5% CO2 tissue culture incubator for 1 h.
After 1 h of incubation, aspirate non-polymerized collagen from the 10 cm dishes prepared above and immediately add 9 mL of complete EpiLife medium. Transfer the dishes into a 37 °C/5% CO2 tissue culture incubator to equilibrate for 30 min.
Take the dish with keratinocytes that require passaging from the tissue culture incubator and aspirate spent media without disturbing the cell monolayer.
Add 6 mL of DPBS, gently rocking the dish to wash the cells. Aspirate DPBS.
Add 3 mL of Accutase and incubate at 37°C for 6–9 min.
Note: Assess the cells after 5–6 min of incubation. Many cells will start detaching or adopting a round shape morphology during this time. If more than 30% of the cells are still attached, gently tap the dish and return it to the incubator for an additional 2–3 min. After 9 min of incubation, nearly all cells should detach. It is crucial to avoid prolonged cell exposure to Accutase. Do not exceed 9 min.
Critical: If keratinocytes detach completely within only 3–4 min, this indicates that the keratinocytes have started differentiating. In this case, they should not be used for grafting; start again with earlier passage keratinocytes.
After incubation, tap the dish gently and add 3 mL of complete EpiLife medium to neutralize Accutase. Proceed to the next step immediately.
Observe under the microscope. Most cells should be floating in suspension. If not, use a wide cell scraper to collect all partially attached cells, especially from the edges of the dish.
Tilt the dish slightly and use a 10 mL serological pipette to gently pipette the keratinocytes one to two times, ensuring all cells are washed from the surface of the dish. Take care to avoid forming bubbles during this process.
Transfer the dissociated cells into a new, sterile 15 mL conical tube. Add 6 mL of DPBS to rinse away Accutase.
Centrifuge the cells at 200× g for 5 min at RT.
Aspirate supernatant. Resuspend the cell pellet in 1–5 mL of fresh EpiLife medium and count cells using a hemocytometer.
Transfer 2.5–4.0 × 103 keratinocytes per cm2 into each collagen-coated 10 cm dish with pre-equilibrated EpiLife medium prepared in step B3.
Gently and thoroughly disperse the cells by alternating between an up/down and then left/right motion. Repeat these motions two more times to ensure even distribution. Do not swirl the plate to mix. Once the cells are evenly dispersed, place the plates into a 37 °C/5% CO2 tissue culture incubator and leave the plates undisturbed for at least 24 h to allow the cells to attach to the surface of the dish.
The next day, examine cell attachment under a microscope.
Change complete EpiLife medium every other day until the keratinocyte culture reaches approximately 60% confluency. Once it reaches this density, change the medium daily until the culture reaches 70%–80% confluency. At this point, the cells are ready to be passaged.
Note: If keratinocytes are plated at a density of 2.5–4 × 103 cells/cm2, the culture should be ready for subculturing in 4–5 days (see Figure 1A).
Defrosting and plating fibroblasts
Prepare complete FEM medium as described above (see Recipes).
Prewarm a 25 mL aliquot of complete FEM medium to RT.
Add 9 mL of complete FEM medium into two 10 cm dishes and transfer them into a 37 °C/5% CO2 tissue culture incubator to pre-equilibrate for 30 min.
Pipette 5 mL of complete FEM medium into a 15 mL conical tube. Set aside in a biosafety cabinet.
Remove the cryovial containing approximately 5 × 105 frozen fibroblasts from liquid nitrogen storage and immediately place it into a 37 °C water bath. Thaw quickly (approximately 1–2 min), taking care to avoid contamination by keeping the cap above the water level.
Note: Thawing time depends on the volume of frozen cells. After one minute, shake the vial and quickly observe the contents. If most of the cell suspension has melted, you can proceed with the next step.
Spray the entire vial with 70% ethanol, wipe, and proceed under strict aseptic conditions in a biosafety cabinet.
Transfer thawed fibroblasts into a conical tube with 5 mL of complete FEM medium prepared in step C4.
Centrifuge the cells at 200× g for 5 min at RT.
Gently resuspend the pellet in 1 mL of complete FEM medium and transfer 500 µL of the cell suspension with 2.5 × 105 fibroblasts into each 10 cm dish with the pre-equilibrated FEM medium prepared in step C3, to reach a proper/required plating density of 4 × 103 cells/ cm2.
Gently and thoroughly disperse the cells by alternating between an up/down and then left/right motion. Repeat these motions two more times to ensure an even distribution. Do not swirl the plate to mix. Once the cells are evenly dispersed, place the plate into a tissue culture incubator and leave it undisturbed for at least 24 h to allow the cells to attach to the dish.
The next day, examine cell attachment under a microscope.
Change complete FEM medium every other day until the cells reach approximately 80% confluency. Once they reach this density, passage the cells.
Note: If fibroblasts are cultured in FBS-containing medium, they should be transitioned into FEM medium (see Recipes, Table 1). Culturing fibroblasts in complete FEM medium for four days is sufficient for a successful transition, which will result in changes in fibroblast phenotype and accelerated growth of these cells, as shown in Figure 1B.
Fibroblasts must be subcultured to perform better during grafting (see below). The rest of the cells can be frozen for future experiments.
Subculturing fibroblasts
Note: One 10 cm dish with fibroblasts at 85% confluency yields approximately 2.5 × 106 cells (see Figure 1B). Since 2.0 × 106 fibroblasts are needed per surgery, one 10 cm dish is sufficient to obtain this number of cells.
Prepare one 10 cm tissue culture dish and allow Trypsin-EDTA and complete FEM medium to equilibrate to RT.
Add 9 mL of complete FEM medium to the dish and transfer it into a 37 °C/5% CO2 tissue culture incubator to equilibrate for 30 min.
Take the dish with fibroblasts that require passaging from the tissue culture incubator and aspirate spent media without disturbing the cell monolayer.
Add 10 mL of DPBS, gently rocking the dish to wash the cells. Aspirate DPBS.
Add 3 mL of Trypsin-EDTA and incubate at 37 °C in a tissue culture incubator for 2–3 min.
After incubation, tap the dish gently and add 6 mL of complete FEM medium to neutralize Trypsin-EDTA.
Observe the cells under a microscope. Most cells should be floating in suspension. Alternatively, use a wide cell scraper to collect all partially attached cells, especially from the edges of the dish.
Tilt the dish and gently pipette with a 10 mL serological pipette one to two times to wash all fibroblasts from the surface of the dish. Avoid forming bubbles.
Transfer the dissociated cells into a new, sterile 15 mL conical tube. Add 3 mL of DPBS to dilute Trypsin-EDTA.
Centrifuge cells at 200× g for 5 min at RT.
Aspirate supernatant. Resuspend the cell pellet in 1–5 mL of fresh complete FEM medium and count cells using a hemocytometer.
Transfer 2.5 × 103 fibroblasts per cm2 to the dish with pre-equilibrated FEM medium prepared in step D2.
Gently and thoroughly disperse the cells by alternating between an up/down and then left/right motion. Repeat these motions two more times to ensure even distribution. Do not swirl the plate. Once the cells are evenly dispersed, transfer the plate into a 37 °C/5% CO2 tissue culture incubator and leave the plate undisturbed for at least 24 h to allow the cells to attach to the dish.
The next day, check cell attachment under a microscope.
Change the complete FEM medium every other day until cells reach approximately 70%–80% confluency (Figure 1B).
Note: If fibroblasts are plated at the density of 2.5 × 103/cm2, the culture should be ready for subculturing in 3 days.
Preparation of working solutions and cells for grafting (for one graft)
Defrost thrombin stock solution (25 U/mL) overnight on ice at 4 °C.
Defrost the fibrinogen aliquot in a fridge for approximately 1 h before use.
On the day of the surgery, prepare the working Solution 1 and Solution 2 according to Tables 3 and 4 (see Recipes for working solutions). Immediately place all components on ice while preparing cells and mice for surgery.
Collect expanded keratinocytes and fibroblasts from section B and section D, respectively.
Note: It is important that both keratinocytes and fibroblasts are of good quality and within 75% and 85% confluency, respectively, on the day of surgery (Figure 1A–1B). The use of differentiated or senescent cells will result in graft failure. Collect cells immediately before the surgery.
Collect 2.0 × 106 fibroblasts as described in steps D1–D10, count cells using a hemocytometer, and centrifuge the cells to obtain a pellet. Resuspend the pellet in 50 µL of complete FEM medium and put the tube with resuspended fibroblasts on ice. Promptly use for surgery to preserve cell viability.
Collect 2.0 × 106 keratinocytes as described in steps B1–B11, count cells using a hemocytometer, and centrifuge the cells to obtain a pellet. Resuspend the pellet in 50 µL of complete EpiLife medium and put the tube with resuspended keratinocytes on ice. Promptly use for surgery to preserve cell viability.
(Optional) If frozen-then-thawed keratinocytes and fibroblasts are grafted, follow the procedure below (described for one graft):
Thaw 2.0 × 106 keratinocytes and 2.0 × 106 fibroblasts as described in steps A6–A9 and C5–C8, respectively. Count the cells before centrifuging to ensure that the correct number of viable cells are used.
Carefully resuspend the pellet containing thawed fibroblasts in 50 µL of complete FEM medium and the pellet containing thawed keratinocytes in 50 µL of complete EpiLife medium. Keep the resuspended cells on ice and promptly use them for surgery to preserve cell viability.
Note: It is crucial to freeze and thaw cells under optimal conditions to ensure cell viability and functionality. See the supplementary information below.
Place the following on ice to transport to the animal facility for grafting. All volumes shown below are for one graft:
Solution 1 323.3 µL
Suspension of fibroblasts 50 µL
Solution 2 323.3 µL
Suspension of keratinocytes 50 µL
Fibrinogen (40 mg/mL stock) 50 µL
Thrombin (25 U/mL stock) 25 µL
Grafting procedure
Follow the protocol for the insertion of a silicone chamber into a nude mouse as outlined in Diette et al. (2020) or refer to Figure 2B–2D and Video 1. Briefly:
Video 1. Procedure for chamber insertion
Administer anesthesia to the mice via an intraperitoneal injection with prepared anesthetic solution in adherence to animal protocol.
Wipe the back skin of the anesthetized mouse with an alcohol swab and let the skin dry.
Use forceps to pinch and elevate the skin above the flat area of the back. Cut a piece of the skin with curved scissors to create a round incision, approximately 1 cm in size (Figure 2B).
Apply a 200 µL drop of sterile DPBS with antibiotics onto the opened muscle fascia to provide lubrication. Gently release a rim of the skin surrounding the incision (approximately 5 mm) by lifting the edges with forceps. Simultaneously, glide along the rim of the incision with a 1 mL tip, separating the skin from the underlying muscle fascia.
Using straight forceps, pull up the edge of the skin near the incision, creating an elongated slit-like opening between the skin and muscle fascia. Simultaneously, grasp a silicone chamber with curvy forceps and slightly squeeze the dome of the chamber.
Slide one side of the chamber’s rim beneath the skin on one side of the slit (Figure 2C).
With forceps, gently pull the skin over the chamber’s rim on the other side of the slit simultaneously releasing the curvy forceps. Release both forceps and secure the entire lower rim of the chamber beneath the skin by ensuring that the rim is flat under the skin (Figure 2D).
After the chamber has been successfully inserted, transfer the mouse to a clean cage placed on a heating pad without a wire feeder.
Proceed promptly with the cell transplantation procedure as described in section G.
Critical: Ensure that the wound bed inside the implanted chamber does not dry during the procedure.
Figure 2. Visual representation of the grafting procedure. (A) Dimensions of the silicone chamber are shown. (B) The mouse skin is cut to create a hole with a circumference of approximately 1 cm for chamber implantation. The chamber is then inserted into the wound (C–D), and a fibroblast-comprised fibrin hydrogel is delivered to form the dermal layer (E). Following the polymerization of the dermal layer, a keratinocyte-comprised fibrin hydrogel is delivered on top (F).
Delivery of the gel containing fibroblasts
Keep all tubes with working solutions, fibrinogen, thrombin, and cell suspensions on ice before use. Spray the vials with 70% ethanol, wipe them, and proceed under strict aseptic conditions in a biosafety cabinet or a clean procedure room in an animal facility.
Critical: To maintain the integrity and viability of the samples, it is imperative to perform all steps expeditiously, ensuring that reagents and materials are kept chilled on ice throughout the procedure.
Using a pipette, transfer the entire volume of cold working Solution 1 (323.3 µL) into the tube containing the suspended fibroblasts prepared in step E4a (or E5 if thawed cells are used). Gently mix the contents, taking care to avoid the formation of bubbles.
Immediately before transferring into an implanted chamber, add 9.4 µL of 25 U/mL thrombin and 17.3 µL of 40 mg/mL fibrinogen to the cell suspension from step G1. Resuspend gently by pipetting 2–3 times, avoiding bubbles (see Table 3).
Immediately transfer the final mix (400 µL) into the chamber using a narrow-tip 1 mL pipette.
Gently shake the mouse to facilitate the distribution of the hydrogel solution throughout the wound bed.
Note: With secure and proper chamber placement, the fibrin gel should be slightly visible in the lower portion of the silicone chamber (Figure 2E).
Place the mouse in the cage on a warming pad and allow it to remain there for a period of 15 min. This timeframe allows the fibrin hydrogel to undergo polymerization, resulting in the formation of a dermal layer. It is important to provide sufficient time for the hydrogel to solidify and establish the desired structural integrity.
Delivery of the gel containing keratinocytes
While the dermal layer solidifies, prepare the epidermal fibrin gel containing keratinocytes.
Using a pipette, transfer the entire volume of cold working Solution 2 (323.3 µL) into the tube containing the keratinocyte suspension prepared in steps E4b (or E5 if thawed cells are used). Gently mix the content, taking care to avoid the formation of bubbles.
Immediately before transplanting into an implanted chamber with the dermal layer, add 9.4 µL of 25 U/mL thrombin and 17.3 µL of 40 mg/mL fibrinogen to the cell suspension. Resuspend gently by pipetting 2–3 times, avoiding bubbles (Table 4).
Immediately transfer the final mixture (400 µL) into the chamber on top of the solidified fibrin gel containing fibroblasts.
Note: Avoid pulling or tilting the silicon chamber when transferring the mixture. It is important to handle the chamber carefully to prevent any disruption or displacement of the grafted material.
Gently shake the mouse to ensure even distribution of the hydrogel-containing keratinocytes over the solidified hydrogel containing fibroblasts previously transferred into the silicone chamber.
Note: With secure and proper chamber placement, the hydrogels should be visible and fill one-half to three-fourths of the silicone chamber (Figure 2F).
Place the mouse in the cage on a warming pad and allow it to remain there for a period of 15–40 min to let the fibrin gel polymerize to create the epidermal layer.
Critical: Ensure that the mouse remains calm for approximately 15 min to allow each layer to polymerize. Additionally, after applying both fibrin layers, keep the mouse undisturbed for at least 1 h. Consequently, the most optimal duration for anesthesia is 2 h.
Keep the chamber on mice for seven days.
Note: During the first 10 days post-surgery, check the mice regularly and provide them with moist chow daily. This helps to ensure the mice are hydrated and supports their post-operative recovery.
Chamber removal, wound healing, and skin graft maturation
On day 7 post-surgery (Day 19), remove the chamber from grafted mice as described in Diette et al. (2020) or refer to Video 2. Briefly:
Video 2. Procedure for chamber removalAdminister anesthesia to the mice via an intraperitoneal injection with prepared anesthetic solution in adherence to animal protocol.
Administer anesthesia to the mice via an intraperitoneal injection with prepared anesthetic solution in adherence to animal protocol.
Wipe the skin around the chamber and the dome of the chamber with an alcohol swab.
Carefully loosen the skin around the chamber with sterile forceps.
With a clean, gloved finger, gently squeeze the dome of the silicone chamber and carefully pull one side from under the skin. Carefully release one edge of the rim using forceps.
Ensure that the graft is not attached to the inner part of the chamber and the forming skin graft stays in the wound.
Notes:
If the edge of the graft and connective tissues around it remain attached to the silicone chamber, run another pair of forceps along the silicone lip of the chamber to free the graft.
The resulting graft should appear as an off-white plug with no yellow areas, which indicates minimal tissue decay (Figure 3A–3D). The wound surface can exhibit varying levels of dryness, which does not significantly affect graft formation.
Apply New skin liquid bandage onto the mouse skin around the open wound to prevent mouse skin contraction.
Place the mouse into a clean cage (without a wire feeder) that has been resting on a heating pad for at least 1 h. After the mouse recovers from anesthesia, remove the heating pad and monitor the mouse for the duration of the healing (approximately six weeks after surgery).
Note: The scab that forms after chamber removal will naturally detach within 2–2.5 weeks.
Harvest the graft for histological analysis or live in vivo studies six weeks after surgery (day 54 of the protocol).
Note: The size of the human skin grafts after chamber removal is constrained by the chamber dimensions and ranges from 1.0 to 1.5 cm in diameter (Figure 3A–3D). Following healing, the grafts typically measure between 0.4 and 1.2 cm in diameter (Figure 3E–3H). This size is suitable for various applications, such as assessing skin cell functionality, modeling the phenotype of skin disorders, investigating wound healing stages, studying aging, and evaluating skin barrier function. The graft can be analyzed earlier, at five weeks post-surgery, if necessary.
Figure 3. Representative images of grafts at different stages of the healing process. The fibrin-based grafting was performed with (A, E) 2.0 × 106 freshly cultured keratinocytes together with 2.0 × 106 of freshly cultured fibroblasts; (B, F) 1.0 × 106 freshly cultured keratinocytes together with 1.0 × 106 freshly cultured fibroblasts, and (C, G) 2 × 106 frozen-then-thawed keratinocytes together with 2 × 106 frozen-then- thawed fibroblasts. As a control, grafts generated using a classical grafting chamber assay with 5.0 × 106 freshly cultured keratinocytes and 5.0 × 106 freshly cultured fibroblasts are shown (D, H). Representative images of forming grafts on the day of chamber removal (day 7) are shown on top and corresponding fully established grafts at 5 weeks post-grafting are shown on the bottom. OW–opened wound after chamber removal, and CW–closed wound at the day of harvest of the skin graft.
Histological examination of human skin xenografts
Euthanize the mouse on the day of harvesting (Day 54) in accordance with the approved protocol by the Institutional Animal Care and Use Committee (IACUC).
Use a permanent marker to mark the borders of the graft and carefully cut the mouse skin around the graft, maintaining a 5 mm distance from the edge of the graft.
Gently peel off the skin and transfer it to Whatman paper to ensure that it lays flat.
Fix sections with skin grafts in a buffered solution of 10% Formalin for 24 h and then transfer them into 70% ethanol.
Fixed tissue can be paraffin-embedded and sectioned for further histological analysis. If histological analysis is performed, prepare 7 μm sections that include both the center and edge portions of the graft.
Confirm the presence of the human skin graft through a conventional histological analysis of tissue sections stained with hematoxylin and eosin (H&E) (Yanez et al., 2015). In this staining method, the epidermis shows up as a thin, compact multilayer structure on the top of the skin, exhibiting a dark pink color with a stratum corneum overlay (Figure 4A). The dermis appears as a light-pink, substantial layer beneath the epidermis, featuring nuclei stained in purple. Notably, a human graft typically displays a thicker stratum corneum and epidermis compared to adjacent mouse skin (Figure 4A, 4D, 4G, and 4J).
Conduct immunohistochemical analysis following the protocol outlined in Yanez et al. (2015). To label the epidermis, use the anti-K14 antibody (BioLegend 906004) at a 1:2000 dilution. Co-staining with the anti-mouse K1 antibody (BioLegend 905602) at a 1:500 dilution aids in delineating human graft borders (refer to Figure 4B, 4E, 4H, and 4K, where K14 is represented in red and K1 is in green). For visualizing the human dermis, employ a human specific anti-Vimentin antibody (ab16700) at a 1:500 dilution. When co-stained with the anti-human Loricrin antibody (ab176322) at a 1:2000 dilution, the human Vimentin staining provides insights into the proper localization of the human dermis beneath the human epidermis (see Figure 4C, 4F, 4I, and 4L, with Vimentin represented in red and Loricrin in green). Use corresponding secondary antibodies at a 1:300 dilution, as listed in the reagents section. Capture images using a Nikon Eclipse 90i microscope equipped with a Photometrics Cool SNAP HQ2+ DS Fi1 Nikon confocal system or another comparable microscope system.
Figure 4. Histological examination of human skin xenografts. Representative images of fibrin-based grafts generated with 2.0 × 106 freshly cultured keratinocytes together with 2.0 × 106 freshly cultured fibroblasts (A–C), 1.0 × 106 freshly cultured keratinocytes together with 1.0 × 106 freshly cultured fibroblasts (D–F), as well as with 2.0 × 106 frozen-then-thawed keratinocytes together with 2.0 × 106 frozen-then-thawed fibroblasts (G–I) are shown. As a control, grafts generated using a classical grafting chamber assay with 5.0 × 106 freshly cultured keratinocytes and 5.0 × 106 freshly cultured fibroblasts were also analyzed (J–L). H & E staining of the grafts (A, D, G, J). Immunofluorescence staining using mouse-specific Keratin (K)1 (moK1; green) and human/mouse-specific K14 (red). Note that the moK1 antibody stains only the mouse epidermis (B, E, H, K). Immunofluorescence staining using human-specific Loricrin (Lor; green) detects the human epidermis, while human-specific Vimentin (Vim, red) detects the human dermis (C, F, I, L). Nuclei stained with DAPI (blue). Scale bar, 100 μm.
Data analysis
To assess the quality of the grafts, several parameters can be assessed such as (1) the length of the developed human skin graft and (2) the thickness of the epidermis and dermis using histological analysis and immunofluorescence staining, as well as (3) the wound contraction. To compare the means of graft measurements between the grafted groups, conduct an ANOVA test (GraphPad Prism).
Graft measurements
Wound contraction
Using a digital camera and a sterile ruler calibrated in centimeters placed laterally underneath the graft, capture an image of the opened wound (OW) on Day 19 after chamber removal and the closed wound (CW) on Day 54 before graft harvesting (Figure 3D and 3H).
Utilizing the Measure tool in Adobe Illustrator, ascertain the number of pixels equivalent to 1 centimeter on the ruler within the photograph. Establish a pixel-to-centimeter conversion.
Determine the number of pixels representing the diameter of the OW and the CW.
Recompute the wound diameters into centimeters, using the pixel-to-centimeter conversion established earlier.
Divide the calculated wound diameter in two to find the radius of the wound (r).
Recalculate the wound area (cm2) following:
Define the percentage of wound contractions as follows:
Skin layers measurement
Use NIS-elements Nikon Software to merge colors and set the 100 µm scale bar (Figure 4).
Utilize the Measure tool in Adobe Illustrator to measure the size of the human graft and the thickness of the epidermis and dermis based on immunohistochemical staining of skin layers.
Validation of protocol
To establish this grafting method, a total of 28 grafting procedures were performed. Eight grafting procedures were performed with 2 × 106 freshly cultured keratinocytes together with 2 × 106 freshly cultured fibroblasts with a 100% success rate. When the number of cultured cells decreased, the success rate of grafting decreased to 80%. Specifically, five grafting procedures were performed with 1 × 106 freshly cultured keratinocytes together with 1 × 106 freshly cultured fibroblasts, and four grafts successfully formed. Importantly, a typical success rate of a classical chamber grafting assay that utilizes 5 × 106 of freshly cultured keratinocytes together with 5 × 106 of freshly cultured fibroblasts also fluctuates between 60% and 80% (Wang et al., 2000; Diette et al., 2020). Therefore, our grafting approach allows for more consistent human skin graft formation with fewer cell numbers compared to other methods of skin grafting.
Interestingly, when 2 × 106 frozen-then-thawed keratinocytes together with 2 × 106 frozen-then-thawed fibroblasts were used for grafting, our approach resulted in a 72% success rate (8 successful engraftments out of 11 grafts), providing feasibility data for using frozen-then-thawed skin cells in skin transplantation. Additionally, as a control, four classical skin grafting chamber assays were also performed.
The procedure described in this manuscript is currently being used in our laboratory in at least three different projects and is reproducible among multiple laboratory members.
The quality of the human graft, particularly the size and thickness of the epidermis and dermis, are important factors that influence graft persistence and usefulness in downstream applications. We plotted different measurements of the grafts generated under different conditions and found significant differences in the parameters of the resulting grafts (Figure 5A–5D). Four grafts from each group were analyzed in week 6 after surgery.
First, we analyzed the contraction of the wound area during graft healing. The extent of mouse skin contraction can significantly impact the size of resulting human xenografts once they have healed. Our experimental data revealed that grafts created with 2.0 × 106 freshly cultured keratinocytes together with 2.0 × 106 freshly cultured fibroblasts exhibited wound contraction of 52.8% ± 11.2%, while grafts generated with 1.0 × 106 cultured keratinocytes together with 1.0 × 106 freshly cultured fibroblasts demonstrated a contraction of 88.7% ± 3.7%. The utilization of 2 × 106 frozen-then-thawed keratinocytes together with 2 × 106 frozen-then-thawed fibroblasts resulted in the contraction of 62% ± 5.2%. In comparison, the control graft produced using the classical chamber assay performed with 5.0 × 106 cultured keratinocytes together with 5.0 × 106 cultured fibroblasts exhibited a contraction rate of 60.9% ± 4.7% (Figure 5A). The accuracy of graft size measurements was affirmed through concurrent histological analysis, which showed that the grafts generated with 2.0 × 106 freshly cultured keratinocytes together with 2.0 × 106 freshly cultured fibroblasts exhibited the largest surface area (Figure 4A–4C and Figure 5B).
The reported contraction rates of our grafts were in alignment with other reports that utilized other hydrogel-based methods (Del Rio et al., 2002; Llames et al., 2004; Kalyanaraman and Boyce, 2009; Martinez-Santamaria et al., 2012; Yanez et al., 2015; Jorgensen et al., 2020). For instance, Kalyanaraman and Boyce (2009) reported a contraction rate of approximately 64%–70% in the grafts generated with a 4 cm2 fibroblast-populated collagen-glycosaminoglycan sponge as the dermal component (Kalyanaraman and Boyce, 2009). Another study demonstrated a 50% contraction rate by week 3 when a 6.25 cm wound was covered with a three-layered fibrin-based bio-printed skin (Jorgensen et al., 2020). Similarly, 3D-printed collagen-based grafts exhibited a 62.5% contraction rate by week 6 when applied to a 2.5 cm wound (Yanez et al., 2015).
The thickness of the graft is crucial as it affects the degree of shrinkage within the wound bed over time and influences the diffusion of nutrients from the wound bed during the initial healing stages. We measured the thickness of the dermis and epidermis in our grafts based on histological analysis. The average dermal layer thickness within our grafts ranged from 314.2 to 621.4 µm, which was on the lower end of dermal thickness in the native human skin, with no significant differences between grafting variants (Figure 5C). The thickness of the epidermis ranged between 34.2 and 92.3 µm, which was also on the lower end of the epidermal thickness of native human skin (Lintzeri et al., 2022). The only deviation from this number was observed in grafts generated with 2 × 106 frozen-then-thawed keratinocytes together with 2 × 106 frozen-then-thawed fibroblasts. These grafts developed a thicker epidermis, which ranged from 128 to 140 µm (Figure 5D). The differences in the parameters of the grafts generated with frozen-then-thawed cells vs. those produced with freshly cultured cells probably result from functional alterations during the freezing and thawing processes.
The observed differences in graft characteristics highlight the importance of cell numbers and processing methods in achieving desired outcomes in skin grafting procedures.
In summary, grafts generated with 2.0 × 106 freshly cultured keratinocytes and 2.0 × 106 freshly cultured fibroblasts were reproducible and of superior quality. Therefore, this composition emerges as the most optimal variant within the grafting protocol delineated in this study. Grafts made with frozen-then-thawed keratinocytes and fibroblasts can also serve as an appropriate alternative if needed.
Figure 5. Digital planimetry. Individual plots for a wound contraction rate are shown in A. The length of grafts (B), the thickness of the dermis (C), and the thickness of the epidermis (D) were determined by analyzing immunofluorescence images of grafts developed at 6 weeks after surgery (****p < 0.0001; ***p < 0.002; **p < 0.0094; *p < 0.02).
General notes and troubleshooting
General notes
For the first time, our study reports the successful layer-by-layer engraftment of skin cells in vivo. It shows the feasibility of transplanting layered skin without the need to perform 3D skin bioprinting or generate complex 3D skin equivalents in a dish, both labor intensive and requiring specialized equipment (Cooper et al., 1993; Jorgensen et al., 2020). Therefore, with further modifications, the provided method may serve as a foundation for new clinical protocols for skin transplantation. Our protocol is expected to yield superior outcomes compared to grafting approaches involving keratinocyte sheets, semi-split, or fragile full-thickness grafts.
Our protocol is specifically optimized for the use of human neonatal keratinocytes and fibroblasts on passage two. However, it can be tailored to skin cells collected at later passages (up to passage four for keratinocytes and up to passage six for fibroblasts) and of different origins (data not shown).
Our protocol also allows for the successful engraftment of frozen-then-thawed keratinocytes and fibroblasts (Figures 3C and 3G). This is the first report that shows the successful generation of in vivo full-thickness skin grafts using frozen-then-thawed skin cells without prior cell culture expansion. Since the transportation and storage of frozen cells are more accessible than in vitro–produced skin equivalents, the feasibility of transplanting frozen skin cells may dramatically simplify skin transplantation in burn patients and patients with chronic wounds.
The human skin grafts generated with this protocol persist for at least three months after grafting onto immunocompromised mice.
Human fibrin-based hydrogels are commonly used as a matrix for tissue bioengineering and skin transplantation and are FDA-approved. To ensure the clinical applicability and success of our method, we use highly pure and cell culture–tested human plasma-derived fibrinogen and thrombin in this protocol.
In our modified chamber assay, the final concentration of fibrinogen is 1.73 mg/mL and the thrombin concentration is 0.59 U/mL. These concentrations are significantly lower than the recommended ranges for clinical and research applications, where fibrinogen concentrations typically range from 5 to 80 mg/mL and thrombin concentrations range from 1 to 125 U/mL. However, the concentrations used in our formulation produce soft gels that solidify within 15 min. The soft gel structure has relatively large pore sizes that facilitate cell migration within the gel. In addition, the presence of abundant surrounding medium promotes cell viability. The final stiffness of the gel does not affect the graft size, and the gel effectively covers the entire wound area. Therefore, these conditions are likely to enhance cell viability, support cell division and migration, and facilitate cellular rearrangement in a favorable environment.
A potential limitation of our proposed method is the time required to form hydrogel-encapsulated cells. Compared to the standard chamber assay, creating dermal and epidermal layers in our approach results in a 15-min delay in transferring cells from tissue culture dishes to the wound surface. However, this delay does not affect cell viability, and our method results in less contraction of the grafts (52.8%) compared to the standard chamber assay (60.1%)
Troubleshooting
Graft failure. The quality of keratinocytes is critical for successful engraftment. Preventing keratinocytes from reaching a confluency level higher than 75% is essential. Exceeding this threshold can lead to terminal differentiation of the cells, impeding their effective engraftment. Similarly, fibroblasts should not be allowed to reach 85% confluency before using them for surgery, as they should be mitotically active at the time of grafting. Make at least three biological repeats.
The number of fibroblasts is insufficient for surgery. The dermal component can be successfully generated with fewer than 1.0 × 106 fibroblasts without compromising the reproducibility of the method (data not shown).
The number of keratinocytes is insufficient for surgery. Our protocol can be performed with as low as 0.5 × 106 cultured keratinocytes and 0.5 × 106 cultured fibroblasts. However, the success rate of these grafts is lower, approximately 40% (data not shown).
Frozen cells do not engraft as efficiently as freshly cultured cells. It is crucial to freeze and thaw cells properly for their successful engraftment. Our protocol uses Cryo50 (ZenBio) for cryopreserving keratinocytes and CryoStor CS10 (StemCell Technologies) for fibroblasts. The freezing and thawing procedures are conducted according to the manufacturer’s recommendations, which are further described in Supplementary Materials.
The fibrin gel adheres to the silicon chamber during the grafting procedure. During the grafting procedure, one potential challenge is the adhesion of the fibrin gel to the silicon chamber's walls. To prevent this, ensure that the chamber is clean and free of debris or moisture before introducing the fibrin gel. Avoid overloading the chamber and follow the protocol’s recommended volumes of cell-loaded gels. Overfilling the chamber can lead to excess gel contacting the walls, increasing the likelihood of gel adhesion. Following the protocol diligently and using the specified volumes of gel solutions will contribute to the overall success of the procedure and minimize any issues related to the adherence of the gel to the chamber walls.
Acknowledgments
We are grateful for funding support from the following: the National Institutes of Health (R01AR059947, R01AR078551 and U01AR075932), the Department of Defense (DOD) (W81XWH-18-1-0706), the Epidermolysis Bullosa (EB) Research Partnership, the EB Medical Research Foundation, the Cure EB Charity, the Dystrophic Epidermolysis Bullosa Research Association (DEBRA) International, the Gates Frontiers Fund, and AVITA Medical. This protocol is partly based on the work by Diette, N., Kogut, I. and Bilousova, G. (2020). Generation of a Full-Thickness Human Skin Equivalent on an Immunodeficient Mouse. Methods Mol Biol 2109: 169–183.
Competing interests
DRR has an equity interest in AVITA Medical. KB is an employee of AVITA Medical, and AH was an employee at the time the research was conducted. AVITA Medical may potentially benefit from the research findings presented here. Other authors do not have any potential conflicts of interest to declare.
Ethical considerations
All animals were housed and handled in accordance with the Institutional Animal Care and Use Committee (IACUC) of The University of Colorado Anschutz Medical Campus. All studies were performed with the approval of the IACUC committee.
References
Cooper, M. L., Andree, C., Hansbrough, J. F., Zapata-Sirvent, R. L. and Spielvogel, R. L. (1993). Direct comparison of a cultured composite skin substitute containing human keratinocytes and fibroblasts to an epidermal sheet graft containing human keratinocytes on athymic mice. J Invest Dermatol 101(6): 811–819.
Cristobal, L., Asunsolo, A., Sanchez, J., Ortega, M. A., Alvarez-Mon, M., Garcia-Honduvilla, N., Bujan, J. and Maldonado, A. A. (2021). Mouse Models for Human Skin Transplantation: A Systematic Review. Cells Tissues Organs 210(4): 250–259.
Del Rio, M., Larcher, F., Serrano, F., Meana, A., Munoz, M., Garcia, M., Munoz, E., Martin, C., Bernad, A. and Jorcano, J. L. (2002). A preclinical model for the analysis of genetically modified human skin in vivo. Hum Gene Ther 13(8): 959–968.
Diette, N., Kogut, I. and Bilousova, G. (2020). Generation of a Full-Thickness Human Skin Equivalent on an Immunodeficient Mouse. Methods Mol Biol 2109: 169–183.
Escamez, M. J., Garcia, M., Larcher, F., Meana, A., Munoz, E., Jorcano, J. L. and Del Rio, M. (2004). An in vivo model of wound healing in genetically modified skin-humanized mice. J Invest Dermatol 123(6): 1182–1191.
Jorgensen, A. M., Varkey, M., Gorkun, A., Clouse, C., Xu, L., Chou, Z., Murphy, S. V., Molnar, J., Lee, S. J., Yoo, J. J., et al. (2020). Bioprinted Skin Recapitulates Normal Collagen Remodeling in Full-Thickness Wounds. Tissue Eng Part A 26(9-10): 512–526.
Kalyanaraman, B. and Boyce, S. T. (2009). Wound healing on athymic mice with engineered skin substitutes fabricated with keratinocytes harvested from an automated bioreactor. J Surg Res 152(2): 296–302.
Lintzeri, D. A., Karimian, N., Blume-Peytavi, U. and Kottner, J. (2022). Epidermal thickness in healthy humans: a systematic review and meta-analysis. J Eur Acad Dermatol Venereol 36(8): 1191–1200.
Llames, S. G., Del Rio, M., Larcher, F., Garcia, E., Garcia, M., Escamez, M. J., Jorcano, J. L., Holguin, P. and Meana, A. (2004). Human plasma as a dermal scaffold for the generation of a completely autologous bioengineered skin. Transplantation 77(3): 350–355.
Martinez-Santamaria, L., Guerrero-Aspizua, S. and Del Rio, M. (2012). [Skin bioengineering: preclinical and clinical applications]. Actas Dermosifiliogr 103(1): 5–11.
Qiao, J., Philips, E. and Teumer, J. (2008). A graft model for hair development. Exp Dermatol 17(6): 512–518.
Waldron-Lynch, F., Deng, S., Preston-Hurlburt, P., Henegariu, O. and Herold, K. C. (2012). Analysis of human biologics with a mouse skin transplant model in humanized mice. Am J Transplant 12(10): 2652–2662.
Wang, C. K., Nelson, C. F., Brinkman, A. M., Miller, A. C. and Hoeffler, W. K. (2000). Spontaneous cell sorting of fibroblasts and keratinocytes creates an organotypic human skin equivalent. J Invest Dermatol 114(4): 674–680.
Yanez, M., Rincon, J., Dones, A., De Maria, C., Gonzales, R. and Boland, T. (2015). In vivo assessment of printed microvasculature in a bilayer skin graft to treat full-thickness wounds. Tissue Eng Part A 21(1–2): 224–233.
Supplementary information
The following supporting information can be downloaded here:
Supplementary information
Article Information
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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Optical Modulation of the Blood-Brain Barrier for Glioblastoma Treatment
QC Qi Cai
HF Hanwen Fan
XL Xiaoqing Li
MG Monica Giannotta
RB Robert Bachoo
ZQ Zhenpeng Qin
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4920 Views: 795
Reviewed by: Mary Luz Uribe Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Communications Aug 2023
Abstract
The blood–brain barrier (BBB) is a major obstacle to the diagnostics and treatment of many central nervous system (CNS) diseases. A prime example of this challenge is seen in glioblastoma (GBM), the most aggressive and malignant primary brain tumor. The BBB in brain tumors, or the blood–brain–tumor barrier (BBTB), prevents the efficient delivery of most therapeutics to brain tumors. Current strategies to overcome the BBB for therapeutic delivery, such as using hyperosmotic agents (mannitol), have impeded progress in clinical translation limited by the lack of spatial resolution, high incidences of complications, and potential for toxicity. Focused ultrasound combined with intravenously administered microbubbles enables the transient disruption of the BBB and has progressed to early-phase clinical trials. However, the poor survival with currently approved treatments for GBM highlights the compelling need to develop and validate treatment strategies as well as the screening for more potent anticancer drugs. In this protocol, we introduce an optical method to open the BBTB (OptoBBTB) for therapeutic delivery via ultrashort pulse laser stimulation of vascular targeting plasmonic gold nanoparticles (AuNPs). Specifically, the protocol includes the synthesis and characterization of vascular-targeting AuNPs and a detailed procedure of optoBBTB. We also report the downstream characterization of the drug delivery and tumor treatment efficacy after BBB modulation. Compared with other barrier modulation methods, our optical approach has advantages in high spatial resolution and minimally invasive access to tissues. Overall, optoBBTB allows for the delivery of a variety of therapeutics into the brain and will accelerate drug delivery and screening for CNS disease treatment.
Key features
• Pulsed laser excitation of vascular-targeting gold nanoparticles non-invasively and reversibly modulates the blood–brain barrier permeability.
• OptoBBTB enhances drug delivery in clinically relevant glioblastoma models.
• OptoBBTB has the potential for drug screening and evaluation for superficial brain tumor treatment.
Graphical overview
Keywords: Glioblastoma Blood–brain barrier Blood–brain–tumor barrier Nanoparticles Pulsed laser Drug delivery Chemotherapy
Background
High-grade gliomas represent the most malignant primary brain tumors in adults. Among these, grade IV astrocytomas, also referred to as glioblastoma multiforme (GBM), constitute over 50% of all primary brain tumors (Straehla et al., 2022). Despite standard-of-care treatment that includes maximal safe resection, fractionated radiation to 60 Gy with concurrent daily temozolomide (TMZ) followed by up to six months of adjuvant TMZ, as well as the progress made in developing new treatment approaches, GBM continues to remain predominantly incurable (Stupp et al., 2005). In addition to the heterogeneous and immunosuppressive tumor microenvironment, a significant challenge for GBM therapy lies in the difficulty of delivering sufficient drug concentrations in the tumor due to the protective blood–brain barrier (BBB), which is also known as blood–brain–tumor barrier (BBTB) in brain tumors. The BBB governs the exchange of ions, molecules, and cells between the bloodstream and the brain, ensures optimal neuronal functioning, and shields brain tissue from harmful toxins and pathogens. Nevertheless, the BBB also prevents more than 98% of small-molecule drugs and all macromolecular therapeutics from entering the brain, resulting in a challenge in drug delivery for brain disease treatment. Although a variety of pathologic conditions, including brain tumors, can disrupt the integrity of the BBB (Sarkaria et al., 2018), the disruption is heterogeneous or inadequate to facilitate clinically meaningful drug penetration. Additionally, evidence suggests that GBM tumor cells infiltrate neighboring tissues without disrupting the BBB, leading to inevitable recurrence (Belykh et al., 2020). To date, numerous strategies have been developed to overcome the BBB for drug delivery (van Tellingen et al., 2015; Belykh et al., 2020; Rosenblum et al., 2018; Brunner and Borchard, 2021). Nonetheless, GBM is still considered incurable, and many promising results from preclinical studies do not translate successfully to clinical trials. Thus, it is of utmost importance to create and validate treatment strategies using clinically relevant GBM models to achieve a more efficient transition from preclinical testing to clinical implementation.
Here, we describe an optical method to open the BBTB (optoBBTB) to modulate the BBB for drug delivery in two clinically relevant genetically engineered mouse models (GEMMs) that recapitulate key GBM characteristics (e.g., PS5A1 GEMM for infiltrative tumor margin with an intact BBTB and 73C GEMM for angiogenic tumor core with a gradually disrupted BBTB) (Cai et al., 2023). We show that optoBBTB facilitates the delivery of paclitaxel (Taxol) to both GEMMs, which further reduces tumor growth and prolongs the mice’s survival. These results demonstrate that optoBBTB is effective for drug delivery and GBM treatment in these two clinically relevant GEMMs.
Materials and reagents
Biological materials
Immunodeficient nude mice Foxn1nu, 7 weeks old, female, 20–25 g (Jackson Laboratory, catalog number: Nu/J 002019)
PS5A1 cell line, generated from Dr. Robert Bachoo’s laboratory at the University of Texas Southwestern Medical Center (Singh et al., 2017; Gao et al., 2020)
73C cell line, generated from Dr. Robert Bachoo’s laboratory at the University of Texas Southwestern Medical Center (Singh et al., 2017; Gao et al., 2020)
Anti-JAM-A antibodies BV11, provided by Dr. Monica Giannotta at FIRC Institute of Molecular Oncology Foundation
Reagents
Reagents for glassware cleaning
Hydrochloric acid (HCl) 37% w/w (Fisher Scientific, catalog number: 7647-01-0)
Nitric acid (HNO3) 70% w/w (Fisher Scientific, catalog number: 7697-37-2)
Reagents for gold nanoparticle (AuNP) synthesis and antibody conjugation
Gold (III) chloride trihydrate (HAuCl4·3H2O) (Sigma-Aldrich, catalog number: 520918-1G)
Sodium citrate tribasic dihydrate (Sigma-Aldrich, catalog number: S4641-500G)
Hydroquinone (Sigma-Aldrich, catalog number: H9003-100G)
OPSS-PEG-SVA 3400 Da (Laysan Bio, Inc., catalog number: OPSS-SVA-3400)
mPEG-thiol 1,000 Da (Laysan Bio, Inc., catalog number: M-SH-1K)
Cytiva HyCloneTM phosphate-buffered saline (PBS) (Fisher Scientific, catalog number: SH3025801)
PierceTM borate buffer (20×) (Fisher Scientific, catalog number: PI28341)
Endotoxin-free ultrapure water (Sigma-Aldrich, catalog number: TMS-011-A)
Reagent for cell culture
GibcoTM DMEM high glucose GlutaMAXTM supplement (Fisher Scientific, catalog number: 10-566-016)
GibcoTM DMEM/F-12, no glutamine (Fisher Scientific, catalog number: 21-331-020)
GibcoTM Hanks' Balanced Salt Solution, no calcium, no magnesium, and no phenol red (HBSS) (Fisher Scientific, catalog number: 14-175-095)
Epidermal growth factor (EGF) (Sigma-Aldrich, catalog number: E4127)
Recombinant human fibroblast growth factor basic (FGF2) (Sigma-Aldrich, catalog number: F0291)
Progesterone (Sigma-Aldrich, catalog number: P6149)
GibcoTM B-27TM supplement (50×), serum-free (Fisher Scientific, catalog number: 17-504-044)
GibcoTM insulin-transferrin-selenium-ethanolamine 100× (Fisher Scientific, catalog number: 51-500-056)
GibcoTM fetal bovine serum (FBS) (Fisher Scientific, catalog number: A3160502)
GibcoTM penicillin-streptomycin (Fisher Scientific, catalog number: 15-140-122)
GibcoTM StemProTM AccutaseTM cell dissociation reagent (Fisher Scientific, catalog number: A1110501)
GibcoTM Trypsin-EDTA (0.25%), phenol red (Fisher Scientific, catalog number: 25-200-056)
Reagents for optoBBTB and drug delivery
0.9% saline solution, sterile (Fisher Scientific, catalog number: S5815)
Isoflurane USP (Covetrus, catalog number: NDC 66794-017-25)
Cremophor® EL (Millipore Sigma, catalog number: 61791-12-6)
Ethanol absolute (Fisher Scientific, catalog number: BP2818100)
Taxol Janelia Fluor® 646 (Tocris Bioscience, catalog number: 6266)
Paclitaxel (Sigma-Aldrich, catalog number: T7191)
Buprenorphine SR-LAB 5 mL (1 mg/mL) (ZooPharm, catalog number: BSRLAB1)
Reagents for tissue collection and ex vivo drug delivery analysis
Paraformaldehyde, 4% in PBS (Fisher Scientific, catalog number: AAJ61899AP)
Sucrose (Millipore Sigma, catalog number: 57-50-1)
Hoechst 33342 solution (Fisher Scientific, catalog number: PI62249)
Scigen Tissue-PlusTM O.C.T. compound (Fisher Scientific, catalog number: 23-730-571)
InvitrogenTM Fluoromount-GTM mounting medium (Fisher Scientific, catalog number: 50-187-88)
Solutions
HAuCl4 solution, 1 M (see Recipes)
HAuCl4 solution, 25 mM (see Recipes)
Sodium citrate solution, 112.2 mM (see Recipes)
Sodium citrate solution, 15 mM (see Recipes)
Hydroquinone solution, 25 mM (see Recipes)
2 mM borate buffer stock solution, pH = 8.5 (see Recipes)
10% sucrose stock solution (see Recipes)
20% sucrose stock solution (see Recipes)
30% sucrose stock solution (see Recipes)
Recipes
HAuCl4 solution, 1 M
*Note: Upon unsealing the bottle, rapidly transfer the HAuCl4·3H2O into a tube to measure the weight. Aliquot the stock solution into 0.5 mL Eppendorfs and snap-freeze the solution with liquid nitrogen for future utilization. The stocks can be stored at -80 °C.
Reagent Final concentration Quantity or Volume
HAuCl4·3H2O 1 M 1 g (see note*)
Ultrapure H2O n/a 2.539 mL
Total (optional) n/a n/a
HAuCl4 solution, 25 mM
Note: This solution should be prepared freshly.
Reagent Final concentration Quantity or Volume
HAuCl4 (1 M) 25 mM 0.05 mL
Ultrapure H2O n/a 1.95 mL
Total (optional) n/a 2 mL
Sodium citrate solution, 112.2 mM
Note: This solution should be prepared freshly.
Reagent Final concentration Quantity or Volume
Sodium citrate 112.2 mM 0.165 g
Ultrapure H2O n/a 5 mL
Total (optional) n/a n/a
Sodium citrate solution, 15 mM
Note: This solution should be prepared freshly.
Reagent Final concentration Quantity or Volume
Sodium citrate 15 mM 0.022 g
Ultrapure H2O n/a 5 mL
Total (optional) n/a n/a
Hydroquinone solution, 25 mM
Note: This solution should be prepared freshly.
Reagent Final concentration Quantity or Volume
Hydroquinone 25 mM 0.0138 g
Ultrapure H2O n/a 5 mL
Total (optional) n/a n/a
2 mM borate buffer stock solution, pH = 8.5
Reagent Final concentration Quantity or Volume
Borate buffer (20×) 2 mM 0.2 mL
Ultrapure H2O n/a 99.8 mL
Total (optional) n/a 100 mL
10% sucrose stock solution
*Note: PBS should be added in increments of 10 mL. Allow the solution to mix completely before adding any more PBS.
Reagent Final concentration Quantity or Volume
Sucrose 10% (w/v) 10 g
PBS (1×) n/a see note*
Total (optional) n/a 100 mL
20% sucrose stock solution
*Note: PBS should be added in increments of 10 mL. Allow the solution to mix completely before adding any more PBS.
Reagent Final concentration Quantity or Volume
Sucrose 20% (w/v) 20 g
PBS (1×) n/a see note*
Total (optional) n/a 100 mL
30% sucrose stock solution (100 mL)
*Note: PBS should be added in increments of 10 mL. Allow the solution to mix completely before adding any more PBS.
Reagent Final concentration Quantity or Volume
Sucrose 30% (w/v) 30 g
PBS (1×) n/a see note*
Total n/a 100 mL
Laboratory supplies
Supplies for AuNP synthesis and antibody conjugation
FisherbrandTM polygon stir bars (Fisher Scientific, catalog number: 14-512-127)
FisherbrandTM IsotempTM stirrer (Fisher Scientific, catalog number: S88850200)
PYREXTM short-neck heavy-wall round-bottom boiling flasks, standard taper joints (Fisher Scientific, catalog number: 10-068-1C)
PYREXTM narrow-mouth heavy-duty glass Erlenmeyer flask 250 mL (Fisher Scientific, catalog number: 10-040F)
DWK Life Sciences KimbleTM KIMAXTM west condensers (Fisher Scientific, catalog number: K452000-2430)
PYREXTM crystallizing dishes (Fisher Scientific, catalog number: 08-741-D)
FisherbrandTM sterile polystyrene disposable serological pipettes with magnifier stripe 10 mL (Fisher Scientific, catalog number: 13-678-11E)
FisherbrandTM sterile polystyrene disposable serological pipettes with magnifier stripe 50 mL (Fisher Scientific, catalog number: 13-678-11F)
FisherbrandTM premium microcentrifuge tubes 2.0 mL (Fisher Scientific, catalog number: 05-408-138)
FisherbrandTM premium microcentrifuge tubes 1.5 mL (Fisher Scientific, catalog number: 05-408-129)
Thermo ScientificTM low protein binding microcentrifuge tubes 1.5 mL (Fisher Scientific, catalog number: PI90410)
Thermo ScientificTM NuncTM 15 mL conical sterile polypropylene centrifuge tubes (Fisher Scientific, catalog number: 12-565-269)
Thermo ScientificTM NuncTM 50 mL conical sterile polypropylene centrifuge tubes (Fisher Scientific, catalog number: 12-565-271)
0.22 μm syringe filters (VWR, catalog number: 76479-030)
SpectrumTM trial size kits for biotech-grade CE dialysis membrane tubing (1 m), MWCO: 20 kd (Fisher Scientific, catalog number: 08-801-251)
Supplies for AuNP characterization
FisherbrandTM 384-well polystyrene plates (Fisher Scientific, catalog number: 12-566-625)
FisherbrandTM disposable cuvettes (Fisher Scientific, catalog number: 14-955-127)
Electron microscopy sciences carbon support film 5–6 nm thick on square 200 mesh copper grid (Fisher Scientific, catalog number: NC9044609)
Supplies for cell culture
Thermo ScientificTM NuncTM EasYFlaskTM cell culture flasks 75 cm2 (Fisher Scientific, catalog number: 12-565-349)
Thermo ScientificTM NunclonTM SpheraTM cell culture flasks 75 cm2 (Fisher Scientific, catalog number: 12-566-440)
Supplies for tumor cell transplantation in mouse and optoBBTB
Micropipette puller (Sutter Instrument, catalog number: Co. P-1000)
Instech PE-20 tubing (Fisher Scientific, catalog number: 50-890-048)
BD Micro-FineTM IV insulin syringes (Fisher Scientific, catalog number: 14-829-1D)
Med Vet International Exel butterfly infusion set, with 27 G × ¾ in. needle, 12 in. tubing (Fisher Scientific, catalog number: 50-209-2293)
PuritanTM cotton-tipped applicators (Fisher Scientific, catalog number: 22-029-553)
FisherbrandTM sterile alcohol prep pads (Fisher Scientific, catalog number: 22-363-750)
World Precision Instrument mouse dissecting kit (Fisher Scientific, catalog number: 50-822-920)
Henkel Loctite 4014 medical device instant adhesive clear (Henkel Adhesives, catalog number: 202152)
Body double standard set (Reynolds Advanced Materials)
Supplies for tissue processing and ex vivo drug delivery analysis
BD VacutainerTM eccentric tip syringe (Fisher Scientific, catalog number: B300613)
FisherbrandTM SuperfrostTM plus microscope slides (Fisher Scientific, catalog number: 12-550-15)
FisherbrandTM cover glasses: rectangles (Fisher Scientific, catalog number: 12-544-18P)
Equipment
New Era Pump Systems Inc single channel syringe pump (Fisher Scientific, catalog number: NC1072839)
FisherbrandTM accuSpinTM Micro 17 microcentrifuge (Fisher Scientific, catalog number: 13-100-675)
BioTek Synergy 2 plate reader multi-mode (The LabWorld Group, catalog number: 18531)
Dynamic light scattering (DLS) zetasizer nano zs (Malvern Panalytical)
Electron microscope (JEOL Ltd., model: JEM-1400 plus)
Diode-pumped high energy picosecond Nd: YAG lasers (EKSPLA, catalog number: PL2230)
Coherent® LabMax touch power and energy meter (Coherent®, catalog number: 2256258)
Coherent® EnergyMax laser energy sensors (Coherent®, catalog number: 1110746)
Isoflurane vaporizer (Kent Scientific, catalog number: VetFlo-1231)
DC temperature controller system (FHC, catalog number: 40-90-8D)
Rectal thermistor probe (FHC, catalog number: 40-90-5D-02)
Heating pad (FHC, catalog number: 40-90-2-07)
Syringe pump controller (World Precision Instruments, catalog number: micro-2T)
Brushless micromotor system (Osada, catalog number: EXL-M40)
Dual small animal stereotaxic instrument (KOPF, catalog number: 902)
Zeiss Stemi 305 microscope (Zeiss, catalog number: STEMI305-T-B)
HM525 NX cryostat (Epredia, catalog number: HM525 NX)
Virtual slide microscope (Olympus, model: VS120)
Spinning disk super-resolution microscope (Olympus, model: SD-OSR)
IVIS® Lumina Series III (PerkinElmer, catalog number: CLS136334)
Software and datasets
ImageJ (version 1.53c, 6/27/2020)
GraphPad Prism (version 9.5.0, 12/6/2022)
Procedure
General preparation for nanoparticle synthesis and conjugation
Prepare fresh aqua regia in the chemical fume hood, by gently mixing 3:1 (v/v) HCl: HNO3.
Soak the glassware, magnetic stir bar, and condenser in aqua regia for at least 30 min.
CAUTION: Mishandling of aqua regia can result in significant hazards, including the risk of explosions, skin burns, eye damage upon contact, harm to mucous membranes and the upper respiratory tract if inhaled, as well as potential harm to internal organs if ingested. It is crucial to never seal aqua regia in a closed container, as the gases produced can lead to pressure buildup and potentially over-pressurize the vessel. Proper personal protective equipment (PPE) is essential when working with aqua regia.
Rinse the glassware and magnetic stir bars thoroughly with Millipore-filtered water in the chemical fume hood followed by rinsing with endotoxin-free ultrapure water in the Class 2, type A2 biosafety cabinet.
To reduce the possibility of nanoparticle contamination for further in vivo experiments, the nanoparticle syntheses are performed in the biosafety cabinet by following all strict precautions normally adopted during cell culture.
CRITICAL STEP: To avoid nanoparticle contamination, all plasticware used in the synthesis should be endotoxin-free certified. All solvents and other reagents used for nanoparticle preparation in this work were strictly opened inside the biosafety cabinet. All the containers (i.e., conical flasks and dialysis containers) should be rinsed with 70% ethanol and Millipore-filtered water and covered properly with a piece of disinfected parafilm. All reagent solutions are filtered through 0.2 μm Millipore syringe filters in the biosafety cabinet before use unless specified.
Synthesis of 15 nm gold seeds
Add 98 mL of endotoxin-free ultrapure water to the 250 mL glass round-bottom flask.
Add 1 mL of HAuCl4 solution (25 mM) to the round-bottle flask.
Boil the solution under reflux and thorough stirring.
CRITICAL STEP: The thorough and effective mixing is one of the key steps to achieving homogenous nucleation and growth to obtain nanoparticles with high monodispersity.
Add 1 mL of sodium citrate solution (112.2 mM) while stirring.
CRITICAL STEP: The reducing agent should be added quickly to obtain a homogenous nanoparticle growth and high monodispersity.
Stir and boil the solution for 10 min.
Turn off the heat and allow the system to cool to room temperature (23–25 °C) while stirring. The resulting AuNPs should exhibit a ruby-red color.
Characterize the gold seeds.
Measure nanoparticle size and size distribution using dynamic light scattering (DLS) (zetasizer nano zs). The measured hydrodynamic diameter of the nanoparticles is approximately 17 nm. The polydispersity index (PDI) is approximately 0.05.
CRITICAL STEP: The hydrodynamic diameter of the seeds should be 17–18 nm with a relatively low PDI (<0.1) to grow larger nanoparticles with the desired size and quality.
TROUBLESHOOTING: If larger or polydisperse nanoparticle sizes are observed, ensure the solution is boiled before adding the reducing agent. Ensure thorough mixing.
Record UV-Vis-NIR absorption spectra using BioTek Synergy 2 plate reader for concentration measurements (Haiss et al., 2007).
Measure the nanoparticle size and morphology using transmission electron microscopy (JEM-1400 plus electron microscope). The nanoparticle shape should be semi-spherical, and the diameter should be approximately 15 nm.
PAUSE POINT: The seeds can be stored at 4 °C for two weeks. Characterization is required prior to reusing the seeds.
Seed-mediated synthesis of 50 nm AuNPs
Add 95 mL of endotoxin-free ultrapure water to a 250 mL Erlenmeyer flask.
Add 963 μL of HAuCl4 solution (25 mM).
While stirring at room temperature, add 3.71 mL of the as-prepared gold seeds (2.23 nM).
CRITICAL STEP: For gold seeds at different concentrations, use the molar calculation to determine the equivalent volume to be added.
Under thorough mixing, quickly add 963 μL of sodium citrate solution (15 mM) and 963 μL of hydroquinone solution (25 mM).
CRITICAL STEP: These two reducing solutions should be added in quick succession to obtain nanoparticles with high monodispersity.
Stir overnight. Cover the flask with a piece of disinfected parafilm to protect the solution from contamination.
Characterize the nanoparticles.
Measure nanoparticle size and size distribution using DLS (zetasizer nano zs). The measured hydrodynamic diameter of such prepared nanoparticles is approximately 47 nm with a PDI < 0.1.
Record UV-Vis-NIR absorption spectra using BioTek Synergy 2 plate reader.
Measure the nanoparticle size and morphology using transmission electron microscopy (JEM-1400 plus electron microscope). The nanoparticle shape should be spherical, and the diameter should be approximately 50 nm.
TROUBLESHOOTING: If polydisperse nanoparticle sizes are observed, ensure the reducing agents are freshly prepared. Ensure thorough mixing.
Concentrate the nanoparticles.
Allocate the nanoparticles in 2 mL endotoxin-free Eppendorf and centrifuge the nanoparticles at 1,300× g for 30 min at room temperature.
Discard the supernatant and collect the nanoparticle pellet in a 2 mL endotoxin-free Eppendorf.
Characterize the nanoparticles after the concentration process.
Dilute the nanoparticles 100-fold using endotoxin-free pure water and perform DLS for size measurement. The measured hydrodynamic diameter of such nanoparticles is approximately 50 nm with a PDI < 0.1.
TROUBLESHOOTING: Larger nanoparticle sizes after centrifugation indicate aggregation during the concentration step. Ensure proper centrifugation time and speed.
Use the diluted nanoparticles to record UV-Vis-NIR absorption spectra for concentration calculation.
PAUSE POINT: The concentrated nanoparticles can be stored at 4 °C for two weeks. Characterization is required before reusing the nanoparticles.
Preparation of polyethylene glycol–antibody conjugates
Thaw the BV11 (anti-JAM-A antibody) on ice, dilute to 0.5 mg/mL in PBS, then dilute in 2 mM borate buffer at pH 8.5 to 0.05 mg/mL.
CAUTION: The borate buffer and PBS should be cooled on ice before starting the experiment. The antibody solution should be kept on ice. Do not filter the antibody solution.
Dissolve OPSS-PEG-SVA in 2 mM borate buffer and quickly add to the antibody solution at a 125:1 molar ratio.
CAUTION: The OPSS-PEG-SVA solution should be kept on ice.
Transfer the solution to a 15 mL Falcon tube, vortex briefly, and shake on ice for 3 h.
Remove the unreacted OPSS-PEG-SVA by dialysis.
Disinfect a plastic container with 70% ethanol, followed by rinsing with Millipore-filtered water and then rinsing with endotoxin-free ultrapure water. Fill the container with 3 L of borate buffer (2 mM, pH 8.5). Cover the container with disinfected parafilm and aluminum film. Cool the buffer to 4 °C.
Immerse a 20 kDa MWCO membrane in the borate buffer for at least 30 min to activate the membrane.
Fill the dialysis bag with the antibody-PEG mixture, followed by dialysis at 4 °C overnight to remove free OPSS-PEG-SVA under mild stirring.
CRITICAL STEP: To ensure the thorough removal of the free OPSS-PEG-SVA, the borate buffer can be replaced after dialysis for 3 h.
Functionalization of AuNPs with polyethylene glycol–antibody conjugates
Collect the thiolated antibodies in a 15 mL Falcon tube and mix the solution with concentrated AuNPs at a 200:1 molar ratio. Shake the nanoparticle–antibody solution for 1 h on ice.
Add mPEG-thiol solution (1,000 Da) at 6 PEG/nm2 to backfill the empty space of AuNPs. Shake the solution for 1 h on ice.
Purify the resulting AuNP-BV11 conjugates by centrifugation (1,300× g, 30 min, 4 °C) with ice-cold borate buffer (2 mM, pH 8.5). Repeat the process three times to remove excess antibodies.
Dilute the nanoparticles 500-fold using borate buffer (2 mM) and characterize the nanoparticles by DLS and UV-Vis-NIR absorption spectra. The measured diameter of these nanoparticles should be approximately 70 nm and the nanoparticle concentration approximately 20 nM. Representative characterization of AuNP and AuNP-BV11 is shown in Figure 1.
PAUSE POINT: AuNP-BV11 can be stored at 4 °C for up to two weeks.
Figure 1. Characterization of the gold nanoparticles (AuNPs). (A) The morphology and size of the AuNP core are characterized by transmission electron microscopy. The size of the nanoparticles (50 ± 4 nm) was measured with ImageJ by manually counting 100 particles. (B) The localized surface plasmon resonance peaks of AuNP and AuNP-BV11 are characterized by UV-Vis-NIR spectroscopy. (C) The hydrodynamic diameter distribution by the relative intensity of AuNP and AuNP-BV11 is characterized by dynamic light scattering. The Z-average for AuNP and AuNP-BV11 was 49 nm and 69 nm, respectively. The data was plotted using GraphPad Prism software. The data was first published in Cai et al. (2023) and presented with modifications.
Glioma cell culture
Culture PS5A1 cells as free-floating neurospheres in DMEM/F12 medium containing B-27 2%, 20 ng/mL EGF, 20 ng/mL FGF2, 20 ng/mL progesterone, and 1% insulin-transferrin-selenium-ethanolamine. Subculture the glioma cells two days before transplantation to reach 70%–80% confluence to use.
Culture 73C glioma cells in DMEM high glucose medium containing 10% FBS and 1% penicillin- streptomycin. Subculture the glioma cells 2 days before transplantation to reach 70%–80% confluence to use.
Before cell transplantation, dissociate the PS5A1 and 73C glioma cells with cell dissociation reagent and Trypsin-EDTA, respectively, and resuspended in HBSS.
Prepare cell suspensions using HBSS solution to achieve a density of 2 × 105 cells/μL.
General preoperative procedure
At the start of surgical procedures and between surgeries, sterilize surgical tools (World Precision Instrument mouse dissecting kit) with a bead sterilizer for 15 min.
Clean the operative table with 70% ethanol and cover it with a sterile drape to create a sterile environment for placing sterile tools.
CRITICAL STEP: Maintain sterility throughout the entire surgical procedure.
For all procedures, anesthetize the mouse with 2%–3% isoflurane (in the air) and keep them on a heating pad at 37 °C.
CAUTION: Start the experiment once the mouse has become unresponsive to painful stimuli, such as a tail pinch. Improperly managed isoflurane can be harmful; prolonged exposure may result in adverse health effects for the operator, including symptoms like dizziness, headaches, vomiting, and nausea. The use of an exhaust system is imperative to effectively remove excess isoflurane during the surgery.
Cover the mice’s eyes with ophthalmic ointment.
Glioma cells transplantation
Subcutaneously inject Buprenorphine (1 mg/kg) into the mice before surgery.
Prepare glass micropipettes with 50 μm tips with a micropipette puller (Figure 2A). Connect the glass micropipette with a nanoinjector and set the injection speed to 30 nL/min.
CRITICAL STEP: Inject slowly to avoid an acute increase of intracranial pressure and facilitate diffusion of the fluid.
Glioma cells injection:
Stabilize the mouse on the small-animal Stereotaxic instrument with ear bars. Check the stability under the dissection scope by lightly probing with a pair of forceps.
CRITICAL STEP: There should be no movement of the skull relative to the ear bars.
Disinfect the scalp with an alcohol prep pad.
Make a midline incision of the scalp using small scissors.
Remove the membranes on top of the skull using a sterile cotton swab.
Place a thin layer of glue (Henkel Loctite 4014 medical device instant adhesive clear) on the skull and the surrounding skin. Wait 5–10 min for the glue to be bonded to the skin and skull.
Using a hand-held drill, make a single burr hole in the skull at the injection site (Figure 2B).
CRITICAL STEP: Use only light motions and avoid direct downward pressure. Stop drilling every 20–30 s to remove bone dust using compressed air.
Fill the glass micropipette with tumor cell suspension. Align the micropipette with the burr hole (Figure 2C).
Lower the micropipette until the tip touches the cortical surface and use this point as zero. Lower the micropipette to the desired depth (0.5 mm below the cortical surface).
Carefully inject 368 nL of PS5A1 glioma cell suspension or 92 nL of 73C glioma cell suspension into the mouse cortex.
After injection, allow 2–5 additional minutes of rest time before starting to withdraw the micropipette.
Seal the burr hole with box wax and apply a layer of body double to the skull for protection (Figure 2D).
House the mouse for three days (73C) to two weeks (PS5A1) to allow tumor growth before starting the experiments.
Figure 2. Glioblastoma cell transplantation. (A) Glass micropipette. The scale bar represents 50 μm. (B) Mouse with a burr hole in the skull. The arrow indicates the position of the burr hole. (C) Setup for the glioma cell injection. The zoom-in image illustrates the precise alignment of the glass micropipette with the burr hole. (D) Mouse with body double covered on its head after injection.
Minimal skin opening and laser stimulation of the brain for optoBBTB
Intravenously inject (tail vein) AuNP-BV11 into the GEMMs (18.5 and 37 μg/g in saline for PS5A1 and 73C GEMM, respectively) using PE-20 tubing and butterfly infusion set with 27G × ¾ in needle.
Procedures for optoBBTB:
One hour after the nanoparticle injection, peel the body double to expose the skull. Keep the skull moisturized using saline.
Align the laser beam to the tumor area using the minimum laser power (1%).
Apply one picosecond laser pulse (532 nm, 40 mJ/cm2, 6 mm beam size) through the skull above the tumor area (Figure 3).
Figure 3. Setup for optoBBTB. (A) Schematic diagram of a picosecond laser for optoBBTB. (B) Photograph capturing a mouse undergoing laser excitation as part of the optoBBTB procedure. The laser was configured in continuous mode to align its position accurately.
Evaluation of the optoBBTB efficiency
Fluorescent imaging of the delivery of Taxol Janelia Fluor® 646 (Taxol646) after optoBBTB (Figure 4A and 4B, left):
Dissolve the Taxol646 in a mixer of Cremophor EL:absolute ethanol (1:1 v/v, as the vehicle) to 6 mg/mL and then dilute to 2 mg/mL with saline.
Immediately after optoBBTB, intravenously inject fluorescent Taxol646 (12.5 mg/kg).
Thirty minutes after the injection, perfuse the mouse with ice-cold PBS (20 mL) followed by 4% PFA (20 mL).
Extract the brain and post-fix the brain with 4% PFA (15 mL) at 4 °C overnight.
Dehydrate the brain with 10 mL of 10%, 20%, and 30% sucrose until it sinks to the bottom of the tube.
CRITICAL STEP: The brain should be well-fixed with PFA before cryopreservation with sucrose because sucrose solutions above 10% are hypertonic and will cause water to flow out of cells and tissue shrinkage if tissues are not fully fixed.
Snap-freeze the brain on dry ice.
PAUSE POINT: The brains can be wrapped in aluminum foil and stored at -20 °C.
Put two drops of O.C.T. into a plastic cryomold. Place the brain in the center and pour O.C.T. to embed the brain. Place the mold at -20 °C to freeze.
Slice the brain into 20 μm thick slices using a cryostat.
PAUSE POINT: The slides can be stored in a slide storage box at -20 °C.
Stain the brain slices with Hoechst staining (1:2,000 in PBS) at room temperature for 10 min, followed by rinsing three times with PBS (5 min for each wash).
Blot excess PBS from the non-sample surface of the glass slide.
Slowly add 150 μL of the mounting medium on the glass slide and cover the glass slide with a cover slip.
CRITICAL STEP: Avoid creating bubbles when adding the mounting medium and lower the coverslip to its place.
Cure the slides at room temperature for at least 1 h before imaging.
Evaluation of the treatment efficiency of optoBBTB and Paclitaxel (Taxol) delivery in PS5A1 GEMM (Figure 4A, middle, right):
At 14 dpi, randomly divide the mice into four groups: (1) vehicle control; (2) free Taxol control (12.5 mg/kg); (3) optoBBTB+vehicle; (4) optoBBTB+Taxol (12.5 mg/kg). Prepare five mice for each group.
Perform optoBBTB as described above.
Immediately after optoBBTB, intravenously inject vehicle or Taxol into the mice.
Repeat the treatment every four days for three times. Sacrifice the mice at 42 dpi.
Perfuse the mice with PBS, extract the brains, and image the tumor size by fluorescent imaging.
To analyze the survival, use the same treatment groups with seven mice in each group. The tumor-bearing mice should be euthanized if they develop weight loss (>20%), loss of grooming, seizures, or focal motor deficits according to the approved animal protocol.
Evaluation of the treatment efficiency of optoBBTB and Paclitaxel (Taxol) delivery in 73C GEMM (Figure 4B, middle, right):
At 4 dpi, randomly divide the mice into four groups: (1) vehicle control; (2) free Taxol control (12.5 mg/kg); (3) optoBBTB+vehicle; (4) optoBBTB+Taxol (12.5 mg/kg). Prepare five mice for each group.
Perform optoBBTB as described.
Immediately after optoBBTB, intravenously inject vehicle or Taxol into the mice.
Repeat the treatment every four days for three times. Sacrifice the mice at 15 dpi.
Perfuse the mice with PBS, extract the brains, and image the tumor size by fluorescent imaging.
To analyze the survival, use the same treatment groups with seven mice in each group. The tumor-bearing mice should be euthanized if they develop weight loss (>20%), loss of grooming, seizures, or focal motor deficits according to the approved animal protocol.
Figure 4. Evaluation of Taxol delivery after optoBBTB. (A) Left: OptoBBTB facilitates the delivery of Taxol646 in PS5A1 GEMM. The tumor is indicated by GFP fluorescent (arrows), and BBB opening is characterized by Taxol646 leakage (asterisks). The scale bar represents 1 mm. Middle: Tumor size imaging by GFP fluorescent. Right: Kaplan-Meier survival analysis. N = 7 mice in each group. (B) Left: OptoBBTB facilitates the delivery of Taxol646 in 73C GEMM. The tumor is indicated by high density Hoechst staining (arrows), and BBB opening is characterized by Taxol646 leakage (asterisks). The scale bar represents 1 mm. Middle: Tumor size imaging by GFP fluorescent. Right: Kaplan-Meier survival analysis. N = 7 mice in each group. Statistical analyses were performed using GraphPad Prism software. The data was first published in Cai et al. (2023).
Validation of protocol
This protocol was validated in Cai et al., Nature Communications (2023), DOI: https://doi.org/10.1038/s41467-023-40579-1.
Acknowledgments
This research was partially supported by Cancer Prevention and Research Institute of Texas (CPRIT) grants RP190278 and RP210236, Department of Defense grant W81XWH-21-1-0219, and American Heart Association grant 19CSLOI34770004, National Institute of Health grant RF1NS110499, National Science Foundation grant 2123971, and funds from a Eugene McDermott Professorship to Z.Q. Illustration figures were created with Biorender.com. Figure 1 and Figure 4 were first published in Cai et al. (2023) and modified for this publication.
Competing interests
All authors declare no competing interests.
Ethical considerations
The animal ethics guidelines of the Institutional Animal Care and Use Committee (IACUC) at the University of Texas at Dallas were strictly followed, ensuring that the mouse was euthanized upon reaching either when its brain tumor surpassed the predetermined maximum volume of 1 cm3 or when its weight loss exceeded 20%.
References
Belykh, E., Shaffer, K. V., Lin, C., Byvaltsev, V. A., Preul, M. C. and Chen, L. (2020). Blood-Brain Barrier, Blood-Brain Tumor Barrier, and Fluorescence-Guided Neurosurgical Oncology: Delivering Optical Labels to Brain Tumors. Front. Oncol. 10: e00739. doi: 10.3389/fonc.2020.00739
Brunner, J., Ragupathy, S. and Borchard, G. (2021). Target specific tight junction modulators. Adv. Drug Delivery Rev. 171: 266–288. doi: 10.1016/j.addr.2021.02.008
Cai, Q., Li, X., Xiong, H., Fan, H., Gao, X., Vemireddy, V., Margolis, R., Li, J., Ge, X., Giannotta, M., et al. (2023). Optical blood-brain-tumor barrier modulation expands therapeutic options for glioblastoma treatment. Nat. Commun. 14: 4934. doi: 10.1038/s41467-023-40579-1
Gao, X., Zhang, Z., Mashimo, T., Shen, B., Nyagilo, J., Wang, H., Wang, Y., Liu, Z., Mulgaonkar, A., Hu, X. L., et al. (2020). Gliomas Interact with Non-glioma Brain Cells via Extracellular Vesicles. Cell Rep. 30(8): 2489–2500.e2485. doi: 10.1016/j.celrep.2020.01.089
Haiss, W., Thanh, N. T. K., Aveyard, J. and Fernig, D. G. (2007). Determination of Size and Concentration of Gold Nanoparticles from UV−Vis Spectra. Anal. Chem. 79(11): 4215–4221. doi: 10.1021/ac0702084
Rosenblum, D., Joshi, N., Tao, W., Karp, J. M. and Peer, D. (2018). Progress and challenges towards targeted delivery of cancer therapeutics. Nat. Commun. 9(1): e1038/s41467–018–03705–y. doi: 10.1038/s41467-018-03705-y
Sarkaria, J. N., Hu, L. S., Parney, I. F., Pafundi, D. H., Brinkmann, D. H., Laack, N. N., Giannini, C., Burns, T. C., Kizilbash, S. H., Laramy, J. K., et al. (2018). Is the blood–brain barrier really disrupted in all glioblastomas? A critical assessment of existing clinical data. Neuro-Oncology 20(2): 184–191. doi: 10.1093/neuonc/nox175
Singh, D. K., Kollipara, R. K., Vemireddy, V., Yang, X. L., Sun, Y., Regmi, N., Klingler, S., Hatanpaa, K. J., Raisanen, J., Cho, S. K., et al. (2017). Oncogenes Activate an Autonomous Transcriptional Regulatory Circuit That Drives Glioblastoma. Cell Rep. 18(4): 961–976. doi: 10.1016/j.celrep.2016.12.064
Straehla, J. P., Hajal, C., Safford, H. C., Offeddu, G. S., Boehnke, N., Dacoba, T. G., Wyckoff, J., Kamm, R. D. and Hammond, P. T. (2022). A predictive microfluidic model of human glioblastoma to assess trafficking of blood–brain barrier-penetrant nanoparticles. Proc. Natl. Acad. Sci. U.S.A. 119(23): e2118697119. doi: 10.1073/pnas.2118697119
Stupp, R., Mason, W. P., van den Bent, M. J., Weller, M., Fisher, B., Taphoorn, M. J. B., Belanger, K., Brandes, A. A., Marosi, C., Bogdahn, U., et al. (2005). Radiotherapy plus Concomitant and Adjuvant Temozolomide for Glioblastoma. N Engl J Med 352(10): 987–996. doi: 10.1056/NEJMoa043330
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4,921 | https://bio-protocol.org/en/bpdetail?id=4921&type=0 | # Bio-Protocol Content
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Isolation and Enrichment of Major Primary Neuroglial Cells from Neonatal Mouse Brain
SS Santosh Kumar Samal *
MS Madhav Sharma *
JS Jayasri Das Sarma
(*contributed equally to this work)
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4921 Views: 1435
Reviewed by: Xi FengAchira RoyNafisa M. Jadavji
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Original Research Article:
The authors used this protocol in Viruses Jan 2023
Abstract
The central nervous system (CNS) relies on the complex interaction of neuroglial cells to carry out vital physiological functions. To comprehensively understand the structural and functional interplay between these neuroglial cells, it is essential to establish an appropriate in vitro system that can be utilized for thorough investigation. Traditional protocols for establishing primary neuronal and mixed glial cultures from prenatal mice or neural stem cells require sacrificing pregnant mice and have the drawback of yielding only specific types of cells. Our current protocol overcomes these drawbacks by utilizing the brain from day-0 pups to isolate CNS resident neuroglial cells including astrocytes, microglia, oligodendrocytes [oligodendrocyte precursor cells (OPCs) and differentiated oligodendrocytes], and meningeal fibroblasts, as well as hippocampal neurons, avoiding sacrificing pregnant mice, which makes this procedure efficient and cost effective. Furthermore, through this protocol, we aim to provide step-by-step instructions for isolating and establishing different primary neuroglial cells and their characterization using cell-specific markers. This study presents an opportunity to isolate, culture, and establish all major CNS resident cells individually. These cells can be utilized in various cell-based and biochemical assays to comprehensively investigate the cell-specific roles and behaviors of brain resident cells in a reductionist approach.
Key features
• Efficient isolation of major neuroglial cells like meningeal fibroblasts, neurons, astrocytes, oligodendrocytes, and microglia from a single day-0 neonatal mouse pup’s brain.
• Circumvents the sacrifice of pregnant female mice.
• Acts as a bridging experimental method between secondary cell lines and in vivo systems.
• Isolated cells can be used for performing various cell-based and biochemical assays.
Graphical overview
Steps for isolation of meningeal fibroblast and neuroglial cells from day 0 pups of mice (Created using BioRender.com)
Keywords: Neurons Astrocytes Oligodendrocytes Microglia Meningeal fibroblasts Primary-cells Cell culture In vitro
Background
Intricate neuro-glial interactions, as an intertwined system of brain circuitry, play a vital role in various central nervous system (CNS) functions and in the maintenance of homeostasis [1, 2]. Contemporary research indicates that the role of glial cells is not limited to mere supportive cells for neurons; instead, glial cells carry out complex physiological functions such as recycling neurotransmitters, regulation of the blood–brain barrier, myelin sheath formation, immunological functions, and many more. Impairment or dysfunction of glial cell function frequently disrupts CNS cell communication, potentially leading to diverse pathological conditions [3–5]. To understand this complex cellular and molecular crosstalk in various pathological conditions, diverse experimental animal models have been developed [6, 7]. For instance, transgenic mice expressing mutated forms of amyloid protein precursor and presenilin-1 serve as a disease model for Alzheimer's [8], while transgenic mice overexpressing α-synuclein are widely employed to study Parkinson's disease [9]. Additionally, experimental autoimmune encephalitis and virus-induced demyelination models [mouse hepatitis virus (MHV) and Theiler's murine encephalomyelitis virus] have been extensively used to elucidate the etiology of multiple sclerosis [10, 11].
While in vivo models provide unique insights into neuropathological studies, investigating complex cellular interactions in an in vivo setup to emphasize the cell's individualistic role is an intricate process [12]. However, relying on secondary cell lines may be insufficient to understand the diverse functions of neuroglial cells, as they are transformed cells, and continuous cell divisions accumulate mutations [13, 14]. That is why a suitable in vitro primary culture system is necessary to elucidate the complex array of neuroglial crosstalk and, at the same time, overcome the limitations mentioned above. Existing primary neuroglial cell isolation procedures routinely involve using a certain developmental stage of pups to isolate a specific cell (8–12 pups), which can often be exhausting and time-consuming. Cells like astrocytes and meningeal fibroblasts are usually isolated from day-0 pups (P0) [15–17] whereas oligodendrocytes precursor cells’ (OPCs) isolation is done from 5–7-day-old pups [18]. However, neurons are terminally divided cells, and their isolation requires embryonic pups (E14–E15) [19], where the mice mother must be sacrificed.
Our current protocol overcomes these limitations by using day-0 pups to separate a variety of neuroglial cells. This procedure involves the collection of meninges and hippocampal tissue to isolate meningeal fibroblasts [16] and neurons [20], respectively. The remaining brain tissues (left after isolating meninges and hippocampus) can either be used to establish a mixed glial culture with subsequent isolation of enriched astrocytes [21] and microglia [22] or be used for OPCs culture, using serum-free oligodendrocyte-specific medium. OPCs can later be differentiated into mature oligodendrocytes using chemically defined differentiation medium [17]. In addition, this protocol avoids the sacrifice of female mice, as day-0 pups are used to harvest the cells. Astrocytes and fibroblasts isolated using this protocol can be passaged, whereas oligodendrocytes, neurons, and microglia should be seeded as per the experimental requirement. Therefore, if RNA or proteins from these cells are required, more pups are needed to obtain a larger number of these cells. Microglial cells isolated through this protocol can only be used for immunofluorescence experiments. For gene-level expression studies, it is advisable to use the CD11b microbeads–based magnetic cell sorting procedure to get a high yield of microglial cells [22].
The precision and handling can influence the yield of cells during the isolation process, which is discussed in detail in the procedure section. To mitigate such variability, it is essential that all steps be executed consistently and with the utmost care. This method allows for isolating all major CNS resident neuroglial cells without using fluorescence-activated cell sorting. These cells can be used to understand individual neuroglial cell–specific roles in CNS biology and pathology. This approach also increases the experimental opportunities for the scientific community more efficiently and cost effectively.
Materials and reagents
Biological materials
Animals
Mice: Day-0 mice pups, C57BL/6 (Strain #000664, common name B6) obtained from The Jackson Laboratory
Viruses: RSA59, an isogenic recombinant strain of murine beta corona virus (MHVA59) [23]
Reagents
Hank’s balanced salt solution (HBSS) (10×) (Gibco, catalog number: 14185-052)
Phosphate buffered saline (PBS) pH 7.4 (10×) (Gibco, catalog number: 70011-044)
Sodium bicarbonate (NaHCO3) (7.5%) (Gibco, catalog number: 25080-094)
CAUTION: NaHCO3 is skin and eye irritant. Wear suitable gloves, goggles, and protective clothing.
Dulbecco’s modified Eagle medium (DMEM) (powder) (Gibco, catalog number: 12100-046)
Trypsin 0.25% (1×) (Gibco, catalog number: 15050-057)
L-Glutamine 200 mM (100×) (Gibco, catalog number: 25030-081)
0.5% Trypsin-EDTA (10×) (Gibco, catalog number: 15400-054)
CAUTION: Components containing Trypsin may produce an allergic reaction. EDTA may cause irritation to the eyes and skin. Use respiratory protection as well as protection for skin and eyes.
Heat inactivated horse serum (origin: New Zealand) (Gibco, catalog number: 26050-070)
Penicillin-streptomycin (Pen-Strep) (Gibco, catalog number: 15140-122)
Distilled water (Gibco, catalog number: 15230-147)
Sterile deionized H2O
F12 nutrient mixture (Ham) medium (Gibco, catalog number: 11765-054)
Heat inactivated fetal bovine serum (FBS) (Gibco, catalog number: 10082-147)
D-(+)-Glucose, Hybri-Max, ≥99.5% (Sigma-Aldrich, catalog number: G5146)
Trypsin from bovine pancreas (Sigma-Aldrich, catalog number: T9201-5G)
Laminin from Engelberth-holm swarm murine sarcoma basement membrane (Sigma-Aldrich, catalog number: L2020-1MG)
MEM non-essential amino acids solution (Sigma-Aldrich, catalog number: M7145)
HEPES buffer (1 M) (Gibco, catalog number: 15630-080)
CAUTION: It may cause irritation to the skin, eyes, and upper respiratory tract. Wear protective gloves, goggles, and mask.
Insulin from bovine pancreas (Sigma-Aldrich, catalog number: I6634-50MG)
Poly-D-Lysine hydrobromide (Sigma-Aldrich, catalog number: P7886-100MG)
Neurobasal medium (NB) (Gibco, catalog number: 21103-049)
Deoxyribonuclease I (DNase I) from bovine pancreas (Sigma-Aldrich, catalog number: D5025-15KU)
CAUTION: May cause allergy or asthma symptoms or breathing difficulties if inhaled. Use protective clothing, gloves, and masks while using. Do not vortex, as DNase I is prone to physical denaturation.
B27 supplement 50× (Gibco, catalog number: 17504-044)
Magnesium chloride (MgCl2·6H2O) (SISCO Research Laboratories PVT LTD, catalog number: 1349130)
Calcium chloride (CaCl2·2H2O) (SRL, catalog number: 10035-04-8)
L-Thyroxine sodium salt pentahydrate (T4 thyroxine) (Sigma-Aldrich, catalog number: T0397-1MG)
Apo-Transferrin human (transferrin) (Sigma-Aldrich, catalog number: T-4382)
Putrescine dihydrochloride (Sigma-Aldrich, catalog number: P5780-5G)
Progesterone (Sigma-Aldrich, catalog number: P8783-5G)
Selenium dioxide (Sigma-Aldrich, catalog number: 325473)
CAUTION: Toxic if swallowed or inhaled. May cause damage to organs through prolonged or repeated exposure. Very toxic to aquatic life with long-lasting effects. Avoid contact with skin, eyes, and clothing. Use a face shield, safety glasses, gloves, and respiratory protectant.
Biotin (Sigma-Aldrich, catalog number: B-4501)
0.15 M Borate (Sigma-Aldrich, catalog number: B6768-1KG)
99.9% Ethanol, absolute, analytical CSS reagent (Changshu song sheng fine chemicals, catalog number: GB678-90)
CAUTION: Highly flammable liquid may cause drowsiness and nausea.
Sodium chloride (NaCl) (SRL, catalog number: 7647-14-5)
Potassium chloride (KCl) (MERCK, catalog number: 7447-40-7)
di-Sodium hydrogen phosphate (Na2HPO4) (MERCK, catalog number: 7558-79-4)
Potassium dihydrogen phosphate (KH2PO4) (MERCK, catalog number: 7778-77-0)
Trypan Blue (0.4%) (Gibco, catalog number: 15250061)
Growth factors
Nerve growth factor (NGF) (Invitrogen, catalog number: 13257-019)
Fibroblast growth factor (FGF) (R&D System, catalog number: 133-FB/CF)
Platelet-derived growth factor (PDGF) (R&D System, catalog number: 221-AA)
Neurotrophin-3 (NT3) (PeproTech, catalog number: 450-03-10UG)
Reagents for immunofluorescence
Triton X-100 (Sigma-Aldrich, catalog number: T8787-100ml)
CAUTION: Harmful if swallowed. Causes skin irritation and serious eye damage. Use skin and eye protectants.
Bovine serum albumin, for molecular biology (BSA) (HIMEDIA, catalog number: MB083-25G)
CAUTION: Causes skin and eye irritation; may cause an allergic response. Avoid contact with skin and eyes and wear respiratory protection, gloves, and goggles.
Antibodies
Primary antibodies against various cell markers are diluted with 0.1% BSA (prepared from 1% BSA) as per the given ratio (Table 1).
Table 1. Primary antibodies
Antibody name Dilution Cellular markers for Recommended storage condition
Mouse anti-GFAP (Sigma, catalog number: G3893) 1:100 Astrocytes Stored at -20 °C freezer
Rabbit anti-Vimentin (CST, catalog number: D21H3) 1:200 Meningeal fibroblasts
Rabbit anti-Iba1 (SIGMA Wako, catalog number: 019-19741) 1:600 Microglia
Mouse anti-NG2 (MERCK, catalog number: MAB5384-I) 1:50 OPCs
Rabbit anti-MAP 2 (SIGMA, catalog number: M3696) 1:200 Neurons
Mouse anti-NFM (SIGMA, catalog number: N5389) 1:100 Neurons
Mouse anti-Gal C (H8H9) (Homemade) 1:20 Mature oligodendrocytes
Mouse anti-A2B5 (Homemade) 1:20 OPCs
Rabbit anti-Cx43 (SIGMA, catalog number: C6219) 1:1,000 a gap junction protein
Fluorescently tagged secondary antibodies raised against mouse or rabbit are diluted in 0.1% BSA (prepared from 1% BSA) as per the given ratio (Table 2).
Table 2. Secondary antibodies
Antibody name Dilution Recommended storage condition
Donkey anti-Rabbit Alexa Fluor 568 (Invitrogen, catalog number: A10042) 1:1,000 Stored at 4 °C
Goat anti-Mouse Alexa Fluor 488 (Invitrogen, catalog number: A11001) 1:1,000
Goat anti-Mouse Alexa Fluor 546 (Invitrogen, catalog number: A11003) 1:1,000
Solutions
DMEM solution (see Recipes)
Astrocyte-specific medium (see Recipes)
Hanks for prep solution (see Recipes)
Serum for prep solution (see Recipes)
1× PBS (see Recipes)
HBSS solution (see Recipes)
NB/B27 medium (see Recipes)
Oligodendrocytes-specific media (see Recipes)
Neuron-specific media (see Recipes)
DM supplement (see Recipes)
T4 thyroxine (see Recipes)
30% glucose (1.66 M) (see Recipes)
Insulin (see Recipes)
Biotin (see Recipes)
Oligodendrocyte differentiation media (see Recipes)
DMEM Minus (-) (see Recipes)
DMEM Plus (+) (see Recipes)
HBSS (+) or 1% glucose HBSS solution (see Recipes)
DNase I solution (see Recipes)
Trypsin (see Recipes)
Poly-D-Lysine (PDL) coating (see Recipes)
Wash buffer (see Recipes)
Blocking serum (1% BSA) (see Recipes)
Recipes
DMEM solution
Prepare by dissolving DMEM powder in 1 L of double-autoclaved Milli Q water; then, add NaHCO3. Follow the table below for preparation. Mix properly (so that no clumps remain) and then adjust the pH to 7.2–7.4. Sterilize the solution by passing through a 0.22 µm filter paper. The solution can be stored at 4 °C for up to one month.
Components Quantity Final concentration Recommended storage condition
DMEM powder 1 pouch 13.4 g/L 4 °C
NaHCO3 3.7 g/L 44.04 mM Room temperature (20–25 °C)
Double-autoclaved Milli Q H2O 1 L NA Room temperature
Final volume 1 L 4 °C
Astrocyte-specific medium
Sterilize the solution by passing through a 0.22 µm filter paper. Always prepare fresh; the solution can be stored for approximately one month at 4 °C.
Components Quantity Final concentration Recommended storage condition
FBS 10 mL 10% (v/v) -20 °C
Pen-Strep 1 mL 1% (v/v) -20 °C
L-Glutamine 0.1 mL 0.2 mM -20 °C
DMEM solution 89.9 mL NA 4 °C
Final volume 100 mL 4 °C
Hanks for prep solution
Sterilize the solution by passing through a 0.22 µm filter paper before use. The solution can be stored at 4 °C for up to 1–2 weeks.
Components Quantity Final concentration Recommended storage condition
HBSS (10×) 10 mL 10% (v/v) 4 °C
1 M HEPES buffer 2.5 mL 25 mM 4 °C
7.5% NaHCO3 1 mL 1% (v/v) 4 °C
Pen-Strep 1 mL 1% (v/v) -20 °C
Double-autoclaved Milli Q H2O 85.5 mL NA Room temperature
Final volume 100 mL 4 °C
Serum for prep solution
Sterilize the solution by passing through a 0.22 µm filter paper before use. The solution can be stored at 4 °C for up to one week.
Components Quantity Final concentration Recommended storage condition
Heat-inactivated FBS 10 mL 10% (v/v) -20 °C
Non-essential amino acid solution 1 mL 1% (v/v) 4 °C
L-Glutamine 1 mL 2 mM -20 °C
Pen-Strep 1 mL 1% (v/v) -20 °C
DMEM 87 mL NA 4 °C
Final volume 100 mL 4 °C
1× PBS
Dissolve the following reagents as per the given quantities in 800 mL of distilled water and adjust the pH to 7.2–7.4; then, make up the volume up to 1 L and filter sterilize the solution by passing through a 0.22 µm filter paper.
Components Quantity Final concentration Recommended storage condition
NaCl 8 g 137 mM Room temperature
KCl 0.2 g 2.7 mM Room temperature
Na2HPO4 0.24 g 10 mM Room temperature
KH2PO4 1.44 g 1.8 mM Room temperature
Distilled water 1 L NA Room temperature
Final volume 1 L 4 °C
HBSS solution
100× Ca2+ (90 mM)
Dissolve 1.324 g of CaCl2·2H2O in 100 mL of distilled water.
100× Mg2+ (104.8 mM)
Dissolve 2.13 g of MgCl2·6H2O in 100 mL of distilled water.
HBSS with Ca2+ and Mg2+
Sterilize the solution by passing through a 0.22 µm filter paper. This solution can be stored for up to 1–2 months at 4 °C.
Components Quantity Final concentration Recommended storage condition
HBSS (10×) 10 mL 10% (v/v) 4 °C
Ca2+ (100×) 1 mL 0.9 mM 4 °C
Mg2+ (100×) 1 mL 1.05 mM 4 °C
Double-autoclaved Milli Q H2O 88 mL NA Room temperature
Final volume 100 mL 1% (v/v) 4 °C
NB/B27 medium
Sterilize the solution by passing through a 0.22 µm filter paper. Always prepare fresh and do not store the solution for more than two weeks.
Components Quantity Final concentration Recommended storage condition
B27 Supplement 50× 2 mL 2% (v/v) 4 °C
Pen-Strep 2.5 mL 2.5% (v/v) -20 °C
L-glutamine 2.5 mL 5 mM -20 °C
NB 93 mL NA 4 °C
Final volume 100 mL 4 °C
Oligodendrocytes-specific media
Always prepare fresh and use it according to your experimental planning.
Critical step: Do not filter the growth factors. Always add the growth factors instantly in the NB/B27 incomplete media just before use for your experiments and put growth factors back in the -20 °C freezer without any delay.
Components Quantity Final concentration Recommended storage condition
PDGF (1 µg/mL) 100 µL 2 ng/mL -20 °C
FGF (10 µg/mL) 50 µL 10 ng/mL -20 °C
NT3 (1 µg/mL) 50 µL 1 ng/mL -20 °C
NB/B27 medium 50 µL NA 4 °C
Final volume 50 µL 4 °C
Neuron-specific media
Always prepare the solution fresh just before the experiment.
Critical step: Do not filter the growth factors. Caution should be followed as aforesaid for oligodendrocytes-specific medium.
Components Quantity Final concentration Recommended storage condition
NGF (100 µg/mL) 7 µL 50 ng/mL -20 °C
NB/B27 medium 15 mL NA 4 °C
Final volume mL 4 °C
DM supplement
Filter sterilize the mixture after combining all components using a 0.2 µm filter. Store 1 mL aliquots at -20 °C for approximately two months.
Components Quantity Final concentration Recommended storage condition
Transferrin 5 mL 250 mg/5 mL DMEM -20 °C
Putrescine 5 mL 26.85 mg/5 mL DMEM (33.3 mM) Room temperature
Progesterone 10 mL 3 mg/5 mL [of 95% EtOH (1.908 mM)] (dilute 25 µL of this in 10 mL DMEM) Room temperature
Selenium dioxide 5 mL 3.3 mg/10 mL DMEM (2.974 mM) (dilute 40 µL of this in 5 mL DMEM) Room temperature
Final volume 25 mL -20 °C
T4 thyroxine
Aliquot and store at 4 °C.
Components Quantity Final concentration Recommended storage condition
T4 thyroxine 1 mg 22.5 µM -20 °C
DMEM 50 mL NA 4 °C
Final volume 50 mL 4 °C
30% glucose (1.66 M)
Dissolve 30 g of glucose in 100 mL of sterile ddH2O. Sterilize the solution by passing through a 0.22 µm filter paper and store at 4 °C.
Insulin
Mix the provided quantity of insulin in DMEM and then add 1 N HCl dropwise until the cloudy solution clears. Filter sterilize the solution and store 1 mL aliquots at 4 °C.
Components Quantity Final concentration Recommended storage condition
Insulin 25 mg 4.36 µM -20 °C
DMEM 10 mL NA 4 °C
1 N HCl NA NA Room temperature
Final volume 10 mL 4 °C
Biotin
Dissolve biotin in doubled-distilled water (ddH2O) (5 mg in 100 mL of ddH2O) to make a stock of 204.65 µM. Then, prepare a working stock of 40.93 µM as per the instructions below and sterilize the solution by passing through a 0.2 µm filter paper; store at 4 °C.
Components Quantity Final concentration Recommended storage condition
Biotin 20 mL 40.93 µM 4 °C
DD H2O 80 mL NA Room temperature
Final volume 100 mL 4 °C
Oligodendrocyte differentiation media
Mix all the given components and store the solution at 4 °C.
Components Quantity Final concentration Recommended storage condition
T4 thyroxine 4 mL 0.9 µM -20 °C
30% glucose 2 mL 33.4 mM 4 °C
L-Glutamine 2 mL 4 mM -20 °C
Pen-Strep 2 mL 2% v/v -20 °C
Insulin 1 mL 4.36 µM 4 °C
DM supplement 1 mL 1% v/v -20 °C
Biotin 200 µL 81.86 µM 4 °C
DMEM 94 mL NA 4 °C
F12 medium 94 mL NA 4 °C
Final volume 200 mL 4 °C
DMEM Minus (-)
Sterilize the solution by passing through a 0.22 µm filter paper.
Critical: Always prepare fresh and store at 4 °C for no more than 1–2 weeks.
Components Quantity Final concentration Recommended storage condition
1 M HEPES buffer 1 mL 10 mM 4 °C
Pen-Strep (100×) 1 mL 1% (v/v) -20 °C
Non-essential amino acid solution 1 mL 1% (v/v) 4 °C
DMEM 97 mL NA 4 °C
Final volume 100 mL 4 °C
DMEM Plus (+)
Sterilize the solution by passing through a 0.22 µm filter paper.
Critical: Prepare fresh and store at 4 °C refrigerator for not more than 1–2 weeks. FBS and HS should be filtered first to prevent the clogging of filter paper.
Components Quantity Final concentration Recommended storage condition
1 M HEPES buffer 1 mL 10 mM 4 °C
Pen-Strep (100×) 1 mL 1% (v/v) -20 °C
Non-essential amino acid solution 1 mL 1% (v/v) 4 °C
Heat inactivated FBS 5 mL 5% (v/v) -20 °C
Heat inactivated horse serum 5 mL 5% (v/v) -20 °C
DMEM 87 mL NA 4 °C
Final volume 100 mL 4 °C
HBSS (+) or 1% glucose HBSS solution
Sterilize the solution by passing through a 0.22 µm filter paper. The solution can be stored for up to one month at 4 °C.
Components Quantity Final concentration Recommended storage condition
HBSS (1×) 100 mL NA 4 °C
Glucose 1 g (w/v) 55.5 mM Room temperature
Final volume 100 mL 4 °C
DNase I solution
DNase I enzyme is commercially available as a powder. Sterilize the solution by passing through a 0.2 µm filter paper before use. Aliquot the solution to 2 mL tubes and store at -20 °C.
Components Quantity Final concentration Recommended storage condition
DNase I 3 mg 0.1 mg/mL -20 °C
1× PBS 30 mL NA 4 °C
Final volume 30 mL -20 °C
Trypsin
Sterilize the solution by passing through a 0.2 µm filter before use. Aliquot the solution in 2 mL tubes and store at -20 °C.
Components Quantity Final concentration Recommended storage condition
Trypsin from bovine pancreas 300 mg 10 mg/mL -20 °C
1× PBS 30 mL NA 4 °C
Final volume 30 mL -20 °C
Poly-D-Lysine (PDL) coating
Dissolve 5 mg of PDL in 100 mL of distilled water, then add 400 mL of distilled water to make up the volume.
Sterilize the solution by passing through a 0.2 µm filter paper. Store at 4 °C.
To coat:
Add 1 mL of 0.15 M borate (in 0.15 M NaOH, pH 8.4) per 100 mL of PDL solution and filter coat flasks with the PDL-borate for at least 2 h at room temperature.
Suction the liquid off the plastic and dry at 37 °C overnight.
Reagent setup for immunofluorescence
Wash buffer
Critical: Always prepare fresh just before using and store at 4 °C.
Components Quantity Final concentration Recommended storage condition
1× PBS 49 mL NA 4 °C
Ca2+ (100×) 0.5 mL 0.9 mM 4 °C
Mg2+ (100×) 0.5 mL 1.04 mM 4 °C
Final volume 50 mL 4 °C
Blocking serum (1% BSA)
Mix properly using the vortex and keep the solution for some time to settle the frothing.
Critical: Do not vortex for a long period, otherwise it will cause too much frothing in the solution.
Instead of BSA, goat serum can also be used for the preparation of blocking serum and antibody diluents.
Components Quantity Final concentration Recommended storage condition
BSA 1 g 1% (w/v) 4 °C
Wash buffer 100 mL NA 4 °C
Final volume 100 mL 4 °C
Laboratory supplies
Cell strainer (70 and 100 μm) (Falcon, catalog numbers: 352350 and 352360)
15 mL centrifuge tubes (Tarsons, catalog number: 546021)
50 mL centrifuge tubes (Tarsons, catalog number: 546041)
Serological pipette (5 mL and 10 mL) (Thermo scientific Nunc, catalog numbers: 170355N and 7128)
6-well plate (Nunc Thermo, catalog number: 140675)
35 mm disc (Nunc Thermo Scientific, catalog number: 150460)
60 mm disc (Nunc Thermo Scientific, catalog number: 150288)
100 mm disc (Nunc Thermo Scientific, catalog number: 150350)
Bacterial Petri dish (Tarson, catalog number: 460095)
Pipette controller (Gilson, catalog number: 101300)
Syringes (Dispo Van, Hindustan Syringes and Medical devices, catalog number: 840054SM1)
Pipette tips [Tarson, catalog numbers: 521020 (200–1,000 µL), 521010 (2–200 µL), 521000 (0.2–10 µL)]
0.22 µm filter (Merck, Life Sciences, catalog number: B20345)
4 chambers (mounted on permanox) slides (Nunc, catalog number: 120075LE 1007)
Equipment
Biosafety hood (Thermo Scientific, Heraguard, catalog number: 41237696)
Centrifuge (Eppendorf, model: 5702R, catalog number: 5703YH207703)
CO2 incubator (Thermo Scientific, model: Heracell 150i, catalog number: G-3168)
Dissection microscope (Nikon SMZ 745 Model C-LEDS, catalog number: 226763)
Microscope (Nikon, model: Eclipse Ts2, catalog number: 137714)
Analytical balance (Sartorius PRACTUM213-10IN, catalog number: S/N 003670729)
Shaker incubator (ZHICHENG, catalog number: G-1377 ZHWY-103B)
Water bath with shaker (Jio Tech, catalog number: G-3179)
Forceps (Fine Science Tools, catalog number: 00649-11)
Fine forceps (Fine Science Tools, catalog numbers: 11251-33, 11252-40, 11252-30)
Software and datasets
For image processing
ImageJ (NIH, USA)
Zen 2010 software (Carl Zeiss, Germany)
Procedure
Experimental design
Isolation of meningeal fibroblasts, neurons, and various glial cells such as astrocytes, oligodendrocytes, and microglia from day-0 mouse pups’ brain (and meninges) indicates the potential of the current protocol for mechanistic and translational studies [16, 17, 21, 22]. This protocol relies on the precise separation of meninges (rich in fibroblast cells) and hippocampus (rich in pyramidal neurons) from the brain of any mouse and even rat strain.
The protocol requires one litter (>6–8 pups) to harvest adequate CNS resident cells. In the institute animal facility, one 7–9-weeks-old male and one 7–9-weeks-old female mouse are bred. After visual confirmation of the pregnancy, separate the female mice from the male and keep them under observation for pups. Collect the neonatal pups at day 0 to isolate neuroglial and fibroblast cells. Carefully decapitate the pups and immediately place the head in ice-cold Hanks for prep solution. Under the dissection microscope, remove the cranium and collect the intact brain tissue in another ice-cold Hanks for prep solution. Remove the meninges as much as possible from brain tissue using fine forceps. The tissue should then be mechanically minced using a 5 mL serological pipette (refer to Graphical overview Panel C) and through enzymatic digestion using DNase I and trypsin for 30 min.
To calculate the number of cells, thoroughly mix the cell suspension, take out 20 µL of cell suspension in an Eppendorf tube, and mix it with trypan blue dye. Then, count the cells using a cell counter. After counting the cells, seed them as per the experimental requirement.
Astrocytes, OPCs (P0), and fibroblasts can be plated directly on the cell culture flask, while plates for oligodendrocytes (P1) and neurons need to be coated beforehand with Poly-D-Lysine and laminin for experiments (see Recipes). Plate microglial cells directly on precoated chambered slides for experiments.
Isolated neuroglial cells, except neurons, should be kept in serum for prep solution (refer to steps 45–47 of the Procedure) for 24 h at 37 °C (in 5% CO2), followed by adding the chemically defined medium as per the cell types. Neurons will be plated with DMEM+ medium for 24 h at 37 °C (in 5% CO2) before changing into a chemically defined medium (steps 31–33). Fibroblasts can be plated directly in an astrocyte-specific medium (steps 15–16) (see Recipes) (Figure 1).
Figure 1. Schematic representation showing the experimental design for the isolation procedure
Preparation of reagents and equipment (Timing ~15 min)
Keep the stock solutions ready before proceeding with experiments (refer to the Recipes section). All the necessary reagents should be kept on ice or at 4 °C (e.g., DNase I and Trypsin for thawing).
Turn on the shaker water bath and set it to 37 °C for pre-heating.
Turn on the laminar hood and do the UV sterilization for 30 min before starting the procedure. Then, after putting all items required for the procedure (centrifuge tubes, plates, pipette tips, 100 mm plates, cell strainer etc.) inside, repeat the UV sterilization for 30 min.
CRITICAL STEP: Wipe all the dissecting tools required for the procedure with 70% (v/v) ethanol and then do the UV sterilization on the laminar hood.
Aliquot 10–15 mL of Hanks for prep solution in 50 mL centrifuge tubes (for meninges, collect in a 15 mL centrifuge tube) and keep on ice. For neurons, aliquot HBSS+ glucose or 1% glucose HBSS solutions in a 15 mL centrifuge tube.
Tissue collection (Timing ~30 min)
Collect the day-0 pups and keep them in a Whitman filter paper before proceeding with experiments.
Initial procedures for all the glial cells, meningeal fibroblasts, and neurons are considerably the same.
Carefully decapitate the pups and place their head immediately in ice-cold Hanks for prep solution to efficiently isolate the neuroglial cells.
Caution: Animal sacrifice must be performed as per the institute's approved protocols.
Remove the skin and cranium carefully with minimum damage to the brain using a pair of serrated fine forceps while observing under the microscope.
Once the cranium is removed, scoop out the brain (Figure 2A) and transfer it to a fresh 100 mm Petri dish containing Hanks for prep solution.
Figure 2. Detailed step-by-step visualization of meningeal fibroblast cells isolation from meninges. A. Representative image of brain tissues dissected from a day-0 mice pup. Brain with intact meninges. B. The meninges are isolated from the brain and kept separately for fibroblast culture. Post meninges collection, the meninges-free brain is taken for neuroglial cell and neuron isolation (refer to Figure 3). C. Meninges are subjected to mechanical mincing and subsequently centrifuged at 200× g for 10 min. After re-suspending with HBSS, collected cell pellets are subjected to filtration through a 70 µm cell strainer and centrifuged again at 200× g for 10 min. The final re-suspended pellet is collected for cell counting and then plated according to individual experimental requirements.
Meninges removal (Timing ~10 min/brain)
Carefully remove the meninges from the brain while observing under a dissection microscope, using extra fine forceps. Meninges can be differentiated from brain tissue because of blood vessels that look reddish, whereas the brain is milky white.
Caution: Meninges should be removed carefully without disturbing the brain tissue mass, otherwise isolating the hippocampus for neuronal isolation will be tough.
CRITICAL STEP: This is a crucial step because inadequate removal of meninges can cause cross-contamination of fibroblast cells in astrocyte culture (as the growth medium for both the cells is the same).
Collect the meninges in a 15 mL centrifuge tube containing 13 mL of HBSS.
Pause Point: The collected meninges can be instantly used for fibroblast isolation or can be kept on ice for a few minutes (~30–45 min).
Isolation of meningeal fibroblast cells (Timing ~30 min)
Homogenize the collected meninges using a 5 mL serological pipette; pass it through a 70 μm cell strainer. (Use a syringe plunger to pass cells through.)
Collect the filtrate in a 15 mL centrifuge tube, rinse the strainer with HBSS, and centrifuge at 200× g for 10 min. The pellet is meningeal cells.
Add HBSS again to wash the cells, re-suspend the pellet, strain through a 70 µm strainer, and again centrifuge at for 200× g 10 min. (Repeat this step two times.)
After the washing is done, re-suspend the pellet in astrocyte-specific medium.
Plate the cells and allow to grow for 72 h. We prefer to plate in T75 flasks, but you can also plate in T25 flasks and other cell culture plates based on experimental requirements.
After 72 h, observe the attached cells under a microscope.
Gently wash with 1× HBSS (you can also use 1× PBS) to remove the non-adherent cells and debris and add fresh medium.
Change the culture medium every 2–3 days until confluency.
Isolation of hippocampus (Timing ~20 min/brain)
Observe the meninges-free brain under the microscope and remove the olfactory bulbs and cerebellum. Keep these tissues in a 50 mL centrifuge tube containing Hanks for prep solution and finally separate the cerebral hemisphere.
Cut the cerebral hemisphere into two halves and cut the hippocampus away from the medial surface of the cortex (refer to Figure 3A). Place in a 15 mL centrifuge tube containing 13 mL of HBSS+ on ice.
Caution: The hippocampus is a densely neuron-rich region. Inefficient isolation of the hippocampus can lead to contamination of other glial cells.
TROUBLESHOOTING: The hippocampus looks translucent under the light microscope so, for better visualization, use a black surface beneath the Petri dish or use DMEM- to keep the cerebral hemisphere as it will make the surroundings reddish.
After isolation of the hippocampus, collect the remaining brain tissue mass in the same 50 mL centrifuge tube where you have earlier collected cerebellum and olfactory bulb tissues.
These remaining tissues will be used for the glial cell (astrocyte, oligodendrocytes, and microglia) isolation.
Figure 3. Detailed step-by-step visualization of neuroglial cells isolation procedure. A. Brain with intact meninges (previous steps are described in Figure 2A and 2B); the meninges are removed (for fibroblast culture), and meninge-free brains are taken for major glial cells and neuron culture. The hippocampus region is collected for neuron isolation, while the remaining tissue mass (after isolating meninges and hippocampus) is taken for major glial cell isolation. B. Tissues kept for glial cell isolation are subjected to mechanical mincing (hippocampus will not be minced). C. Minced tissues for glial cells and hippocampus tissues are incubated with DNase I + Trypsin at 180 rpm for 30 min at 37 °C in a shaking water bath. The enzymatically digested tissues are subjected to neutralization using FBS and subsequent centrifugation at 600× g for 10 min at RT. D. The collected cell pellet, after re-suspending with Hanks for prep solution, is subjected to filtration through a 70 µm cell strainer. Final yields are counted and plated according to individual experimental requirements. Step numbers are not mentioned in Figure 3 for enzymatic digestion, neutralization, centrifugation, filtration, and cell plating procedure as they will vary for specific cell types (refer to “Procedure” section).
Isolation of neurons (Timing ~30 min)
Take the 15 mL centrifuge tube containing HBSS+ along with previously collected hippocampus tissues and centrifuge at 100× g for 5 min at 4 °C.
Discard the supernatant and add 1 mL of trypsin and 300 µL of DNase I to the tube. Mix properly and incubate for 15 min at room temperature.
Add 3 mL of DMEM+ media to stop the reaction (neutralization of enzymes).
CRITICAL STEP: Enzymatic over-digestion could damage the culture so do not keep it for a longer time than required.
Centrifuge at 400× g for 5 min at 4 °C.
Discard the supernatant and add DMEM+; gently pipette up and down until it makes a cell suspension.
Then, pass the cell suspension through a 70 µm cell strainer on a 50 mL centrifuge tube and rinse the strainer with an equal volume of media (e.g., 5 mL suspension/5 mL media).
Count the cell number using a cell counter and dilute it to 106 cells/mL with DMEM+.
Seed in PDL/Laminin-coated (see Recipes) tissue culture plates with an appropriate number of cells.
Incubate plates at 37 °C and 5% CO2.
After 24 h, change the medium to neuron-specific media (see Recipes) and observe the neurons for axon outgrowth.
Every 2–3 days, change the medium. After ~5 days, neurons will be ready for experiment.
Isolation of brain resident glial cells (astrocytes, oligodendrocytes, microglia) (Timing: vary according to cell type)
Take the 50 mL centrifuge tube consisting of Hanks for prep solution (5 mL for multiple brains) along with the remaining brain tissue mass (after isolation of meninges and hippocampus).
CRITICAL PERIOD: Isolate meninges (refer to step 10) from brain parenchyma and keep them separate to avoid cross-contamination of fibroblast cells in astrocyte culture.
Pipette up and down 10 times using a 5 mL serological pipette to break up tissue (tissue mincing).
Add 0.25 mL of trypsin and 0.25 mL of DNase I per brain.
Incubate in a shaking water bath at 37 °C for 30 min (refer to the enzymatic digestion step).
After the digestion, add 0.25 mL of FBS per brain and 5 mL of Hanks for prep solution.
Mix properly using a pipette and centrifuge at 600× g for 10 min.
Take the supernatant with a gentle vacuum.
Caution: Do not disturb the pellet while discarding the supernatant, otherwise it will affect the cell amount.
Add 5–10 mL of Hanks for prep solution and centrifuge again at 600× g for 10 min.
Again, take the supernatant with a gentle vacuum.
Add 5–10 mL of Hanks for prep solution, re-suspend the pellet, and put through a 70 µm cell strainer. Centrifuge again at 600× g for 10 min (repeat the step one more time).
Re-suspend the pellet in serum for prep solution.
Count the cells in a cell counter and plate the cells in non-coated tissue culture flasks (usually three brains per one T-75 flask).
Incubate the seeded cells in serum for prep solution for 24 h in a 5% CO2 incubator at 37 °C.
Cell-specific media change
After 24 h, wash cells with HBSS with Ca2+ and Mg2+ and put astrocyte-specific medium (see Recipes) in the flask where astrocytes are to be grown and oligodendrocyte-specific medium in the flask where OPCs are to be grown (go to step 55 for OPCs).
Enrichment of astrocytes and isolation of microglia from mixed glial cells
Once the mixed glial cells form a confluent monolayer, stop the addition of fresh medium for 10 days (starvation) to allow differential adhesion of astrocytes and microglia. Starvation leads to significant microglial growth. In these cultures, it peaked at 12–14 days.
After the starvation is complete, thoroughly agitate the culture flask in an orbital incubator shaker at 180 rpm for 45 min at 37 °C to remove the less adherent microglial cells from tightly adhered astrocytes.
Quickly remove the media with non-adherent cells and collect it in 15 mL tubes (go to step 54).
Microglial isolation (Timing ~10–12 days)
Take the cells suspended in the culture medium in a 15 mL tube and centrifuge at 300× g for 5 min at 4 °C.
Re-suspend the centrifuged pellet and dilute it with fresh astrocyte-specific medium, bringing the cells to a final concentration of 8 × 105 cells/mL; add 0.5 mL to each well of a 4-well chamber slide or 2 mL/well of a 6-well plate.
Astrocyte enrichment (Timing ~10–12 days)
After microglia has been shaken, maintain the flask in astrocyte-specific medium following a wash with sterile PBS. For astrocyte isolation, trypsinize the remaining adherent monolayers of astrocytes and use them as enriched astrocyte cultures for further experimentation.
OPCs isolation (Timing ~2 h)
After 24 h (step 47), remove non-adherent cells by washing with HBBS with Ca2+ and Mg2+.
Add oligodendrocyte-specific medium (see Recipes) to the cells 24 h after removal of serum for prep media.
Maintain these mixed glial cultures in oligodendrocyte-specific medium until confluent (approximately one week).
OPCs, which mainly grow on top of the mixed glia culture (mainly astrocyte layer) and are comparatively less adherent than astrocytes, should be dislodged by washdown method.
Plate the dislodged oligodendrocytes onto Poly-D-Lysine-coated coverslips (see Recipes) and maintain in oligodendrocyte-specific medium until the culture reaches 80% confluence.
Differentiation of OPCs (Timing ~7–10 days)
Remove the media once the OPCs are fully confluent; then, wash the OPCs with 1× HBSS with Ca2+ Mg2+ and add oligodendrocyte differentiation medium (see Recipes) for 7–10 days.
After day 7 onward, these can be used for experimentation.
Data analysis
Perform immunofluorescence experiments using the previously described protocol [24]. Fix 70%–80% confluent culture using 4% PFA). Then, permeabilize the cells with PBS containing 0.5% Triton X-100, followed by blocking with 1% BSA. Incubate the cells with primary antibodies in 0.1% BSA solution for 1 h, followed by washing using PBS with Ca2+ and Mg2+ and then labeling with secondary antibodies in 0.1% BSA for 1 h (see Recipes). Wash the cells and mount them with DAPI (VectaShield, Vector Laboratories) and visualize using the epifluorescence microscope [NIKON eclipse Ti2 microscope and DS Qi2 device camera (TYO)] or confocal microscope (Zeiss confocal microscope LSM710). Images are processed using ImageJ and Zen 2010 software (Carl Zeiss, Germany).
Meningeal fibroblast cells are the largest isolated cells, appearing transparent when viewed under a brightfield microscope. Growing microglial cells have short processes and a fusiform shape, while astrocyte cells form a monolayer in culture with a dense spread-out cell body. The morphological architecture of differentiated oligodendrocytes and OPCs differ significantly; OPCs are typically small cells with bright cytoplasm and short processes, whereas differentiated oligodendrocytes are multi-branched structures. Throughout their growth phase, neural cells exhibit a variety of morphologies; during their early stage, they are less branched and small in size, but with time, they develop networks with other neurons.
Validation of protocol
Experimental validation
Isolated and enriched primary neuroglial cells from day-0 mice pups’ brain show characteristic morphology throughout their growth period. These cells can be further characterized through immunolabeling using their cell-specific markers to verify their distinctiveness (Figures 4 and 5). Meningeal fibroblasts require astrocyte-specific medium as a growth medium. These cells show their elongated structure (largest among all other cells) around day 5–7 post-plating. These cells look transparent under the brightfield microscope (Figure 4A) and express type III intermediate filament protein vimentin (Figure 5A) [16, 25].
Figure 4. Morphological characterization of isolated primary neuroglial and meningeal fibroblast cells. Brightfield images show the specific neuroglial and fibroblast cells observed during the procedural steps of the cultures. A. Meningeal fibroblast cells (after five days of plating). B. Mixed glial cells before starvation. C. After starvation. D, E. Enriched astrocytes and microglia. F, G. Oligodendrocyte precursor cells before and after wash-down, respectively. H. Differentiated oligodendrocytes. I. Neurons at different stages of development (Day 2–Day 10). Scale bars are 50 µm for all brightfield images and 20 µm for Figure 3E).
Figure 5. Immunofluorescence characterization of isolated primary neuroglial and meningeal fibroblast cells. A. Meningeal fibroblast cells. B. Astrocytes are labeled with vimentin (in red) and GFAP (in green). Astrocyte cells are dual positive for both GFAP and vimentin, while meningeal fibroblast cells are only positive for vimentin and negative for GFAP. C. Microglial cells are marked with Iba1 (in red). D. Neuronal cells show double-positive for NFM (in green) and MAP2 (in red). E, F. OPCs and differentiated oligodendrocytes are characterized using NG2 (in green) and Gal C+ A2B5- (Gal C in green, A2B5 in red). For double-immunostaining, Alexa Fluor 488, 546, and 568 secondary antibodies were used (see Recipes), and nuclei were counter-stained with DAPI (in blue). Scale bars are 20 µm in A and B, 50 µm in D and E, and 100 µm in C and F.
After starvation, plate astrocytes as per the experimental requirement in astrocyte-specific medium. Astrocytes are star-shaped cells characterized by dense spread-out cell bodies (darker in color) forming a monolayer around day 10–12 post-plating (Figure 4B and 4D). When the mixed glial cells are fully confluent and subjected to starvation (starvation facilitates microglial growth) [22] for 10–12 days, small bright microglial cells are visible over the monolayer (Figure 4C). After isolation, microglial cells should be plated on a chambered slide. These cells show a characteristic spindle (fusiform) shape with short processes (Figure 4E) and are positive for ionized calcium-binding adaptor molecule 1 (Iba1) protein (Figure 5C) [26, 27]. Enriched astrocytes are characterized by double-immunolabeling with anti-GFAP (glial fibrillary acidic protein) and anti-vimentin antibodies. Astrocyte cells are positive cells for both markers [21, 28] (Figure 5B).
As previously described, both astrocytes and oligodendrocytes are isolated from the same CNS tissues only by growing them in separate cell-specific chemically defined media. OPCs are characterized by a small structure with bright cytoplasm and two processes emerging from the central cytoplasm (Figure 4F and 4G). Once OPCs are differentiated into mature oligodendrocytes, they attain myelinating behavior and show a more branched structure; these branches radiate from the central bright cytoplasm (Figure 4H). The OPCs and differentiated oligodendrocytes positively express proteoglycan nerve/glial antigen 2 (NG2) [29, 30] and galactocerebroside (GalC) [31] respectively (Figure 5E and 5F).
Neurons may take 1–2 weeks post-plating for growth. During the initial days, these cells look small with bright cytoplasm and few processes, but gradually they develop a more branched structure with increased network connections (Figure 4I). These neurons can be characterized by the presence of neuronal-specific marker neurofilament (NFM) and microtubule-associated protein 2 (MAP 2) (Figure 5D) [32, 33].
These isolation procedures enable us to carry out a myriad of downstream experiments, e.g., protein localization studies such as Connexin 43 localization on fibroblast cells (Figure 6A), protein trafficking, or gene expression (RNA and protein) studies. There are numerous prior studies where these cells have been used to understand neuroglial tropism of a hepato-neurotrophic β coronavirus (RSA59 virus: an isogenic recombinant strain of MHVA59) [10] and also virus-induced alteration of protein expression in primary cells [16, 21]. Our findings imply that at MOI 2, the RSA59 virus infects all neuroglial cells (Figure 6B–6E) as well as meningeal fibroblasts (Figure 6A). These infected neuroglial cells can be harvested for downstream RNA and proteins, which then can be examined in accordance with the needs of each individual investigation.
Validation from literature
Astrocytes, being the most abundant glial cell in the CNS, help in regulating pH and K+ buffering and communicate with other glial cells for the maintenance of CNS homeostasis. Failure of neuroglial cells to sustain this homeostasis is fatal for CNS [1, 34]. Astrocytes are involved in several neurodegenerative and neuroinflammatory diseases like Alzheimer’s, Parkinson’s, amyotrophic lateral sclerosis, and multiple sclerosis. Using the current protocol, studies indicate that murine coronavirus (MHVA59) infection of primary astrocytes alters the expression of the astrocytic gap junction protein Cx43 by obstructing the protein’s microtubular trafficking [refer to Figure 2 and Figure 7 (Basu et al., 2017)] [35]. It was also found that the downregulation of Cx43 in vivo corresponds to the depletion of the oligodendrocytic gap junction protein Cx47 [refer to Figure 10 and Figure 11 (Basu et al., 2017)] [21, 35]. &Bgr;-amyloid (Aβ) peptides impede functional gap junction’s activity in primary astrocytes [refer Figure 1 (Maulik et al., 2020)] [36]. Similarly, MHV-A59 infection also reduces Cx43 expression in meningeal fibroblasts, resulting in altered gap junction communication [refer Figure 5 (Bose et al., 2018)] [16], which could affect the integrity of the blood–brain barrier [16, 37]. Therefore, this procedure offers a platform to study blood–brain barrier integrity and protein localization, as well as protein trafficking in neuroglial and fibroblast cells. It also helps in the development of targeted therapies for Alzheimer’s disease, emphasizing Cx43 channels [16, 36]. MHV-A59 infection in primary astrocytes increases Endoplasmic reticulum (ER) stress along with downregulation of the ER-resident chaperone protein, Erp29. Additionally, it has been demonstrated that 4-PBA treatment reduces ER stress by enhancing the expression of the chaperone Erp29 [refer to Figures 1, 2, and 3 (Bose et al., 2023)] [38]. This suggests that the current isolation procedure can also be used to comprehend neuro-glial-cells-specific stress responses [38].
Compromised oligodendroglial lineages are found to be involved in various neuropathologies, including demyelinating diseases and metabolic diseases [39, 40]. In primary culture, the mouse corona viruses RSA59 (a demyelinating strain) and RSMHV2 (a non-demyelinating strain) infect OPCs and mature oligodendrocytes differently. RSA59 infects both lineages (OPCs along with mature oligodendrocytes); on the contrary, RSMHV2 only infects OPCs [refer to Figure 3 and 4 (Kenyon et al., 2015)] [17]. Oligodendrocyte lineages assist in the myelination of neurons and enhance neuronal functionality [41]. Recent investigations have also demonstrated that OPCs are important cytotoxic targets during inflammatory demyelination, participate in antigen presentation, and have immunological attributes [42]. Therefore, this procedure will also help to dissect the cellular mechanism behind virus-induced demyelination through in vitro studies using primary oligodendrocytes.
Figure 6. Downstream applications of isolated neuroglial and meningeal fibroblast cells. All neuroglial cells are getting infected with a recombinant murine coronavirus (RSA59), which shows EGFP expression (green) post-infection of cells (Panel A–E). Infected cells were fixed with 4% PFA and proceeded for an immunofluorescence experiment. A. Cx43, a surface gap junction forming protein (in red), got internalized post RSA59 (in green) infection in meningeal fibroblast cells. B–D. RSA59 virus (in green) shows tropism for astrocytes, OPCs, and microglial cells, confirmed using cell-specific markers GFAP, A2B5, and Iba1 (in red), respectively. E. Confocal image shows the RSA59 virus (in green) also has neuronal tropism marked by neuronal specific marker NFM (in red). For double-immunostaining, Alexa Fluor 488, 546, and 568 secondary antibodies were used as previously described (see Recipes); DAPI marks the cell nuclei (in blue). Scale bars are 20 µm in A, B, D, and E, and 100 µm in C.
Microglia are the brain-resident myeloid cells that perform a variety of dynamic functions such as antigen presentation, phagocytosis, and cytokine generation [43]. Primary astrocytes and microglia treated with TNF α result in differential functional activation of both cells [refer Figure 5 (Marek et al., 2008)] and demonstrate that specific isolation of primary cells is suitable for differential functional activity analysis [22]. In addition to their distinctive function in the brain, several reports also suggest that interacting nexus of neurons with glial cells, oligodendrocytes, and astrocytes enhances myelination in vitro [44]. According to earlier reports, embryonic prenatal rat pups’ hippocampus [19, 45] and adult rats’ hippocampus [46] are required for neuronal culture; however, with this isolation procedure, from a single day-0 post-natal pup brain, neurons can also be isolated along with a wide variety of CNS glial cells and meningeal fibroblasts, bypassing the sacrifice of female mice. The primary neurons also have been used to study differential infectivity of four isogenic recombinant strains of murine beta coronaviruses (RSA59PP, RSA59P, RSMHV2PP, and RSMHV2 P) [refer to Figure 1 (Safiriyu et al., 2023)] [20].
The presented protocol has been extensively employed for several published research works (references given below) in order to perform various cell-based and biochemical experiments including immunofluorescence and gene expression (RNA and protein) studies through RT-qPCR, western blot, and ELISA.
Isolation of meningeal fibroblast was done in Bose et al. (2018) [16]; isolation of astrocytes was done in Basu et al. (2016 and 2017) [21, 35]; isolation of OPCs and differentiated oligodendrocytes was done in Kenyon et al. (2015) [17]; isolation of microglia was done in Marek et al. (2008) [22]; and isolation of primary neurons was done in Safiriyu et al. (2023) [20].
General notes and troubleshooting
Troubleshooting (Table 3)
Table 3. Troubleshooting suggestions are listed here
Step Problem Possible reason Possible solution
11 Low yield of meningeal fibroblast Improper removal of meninges, contamination with brain tissues Properly isolate meninges using a pair of sharp fine forceps, kept separately to reduce astrocyte contamination
35 Fibroblast contamination in astrocyte culture Inadequate removal of meninges Carefully remove meninges as much as possible
36 The brain tissues are too big Poor mincing, diminished scissor functioning Mince for a longer duration, use sharp scissors
41 Low yield of neuroglial cells Removal of cell pellets along with the supernatant (contain viscous myelin debris) Carefully discard the supernatant by observing the pellets; in case of doubt, refilter the supernatant and again centrifuge
44 Clogging of mesh Improper mincing results in larger tissue fragments Refilter using a new 70 µm cell strainer
Acknowledgments
Author Contributions: Manuscript writing S.K.S., M.S. and J.D.S. Figure Preparation S.K.S., M.S. and J.D.S. All authors contributed to the article and read and approved the submitted version.
We would like to thank CSIR for providing fellowships to S.K.S. and M.S. We also thank the central imaging facility and Animal Facility of IISER Kolkata for all the necessary support for our experiments. We also want to thank SERB-POWER grant: SPG/20220/000454 (Promoting opportunities for Women in Exploratory Research) program for providing research funds to JDS for a structured effort toward enhanced diversity in research to ensure equal access and weighed opportunities for Indian women scientists engaged in research and development activities. The funders had no role in study design, data collection, and analysis, the decision to publish, or manuscript preparation.
This procedure has been extensively used in several previous research works: isolation of meningeal fibroblast was done in Bose et al. (2018) [16], isolation of astrocytes was done in Basu et al. (2016 and 2017) [21, 35], isolation of OPCs and differentiated oligodendrocytes was done in Kenyon et al. (2015), isolation of microglia was done in Marek et al. (2008) [22], and isolation of primary neurons was done in Safiriyu et al. (2023) [20].
Competing interests
The authors declare no competing interests.
Ethical considerations
All animal experiments were approved by the ethical committee of the Indian Institute of Science Education and Research (IISER) Kolkata. Animal experiments are performed in strict accordance with the Institute Animal Ethical Committee guidelines. IAEC Protocol number: IISERK/IAEC/AP/2019/29.01 SL NO-43.
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High-throughput Analysis of Capillary Density in Skeletal Muscle Cross Sections
TA Tooba Abbassi-Daloii
SM Sander D. Mallon
SA Salma el Abdellaoui
LV Lenard M. Voortman
VR Vered Raz
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4922 Views: 449
Reviewed by: Chiara AmbrogioWendy Leanne HempstockErin L Davies
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Original Research Article:
The authors used this protocol in eLIFE Feb 2023
Abstract
Capillary density in skeletal muscles is key to estimate exercise capacity in healthy individuals, athletes, and those with muscle-related pathologies. Here, we present a step-by-step, high-throughput semi-automated method for quantifying capillary density from whole human skeletal muscle cross-sections, in areas of the muscle occupied by myofibers. We provide a detailed protocol for immunofluorescence staining, image acquisition, processing, and quantification. Image processing is performed in ImageJ, and data analysis is conducted in R. The provided protocol allows high-throughput quantification of capillary density.
Key features
• This protocol builds upon the method and results described in Abbassi-Daloii et al. (2023b).
• It includes step-by-step details on image acquisition and image processing of the entire muscle section.
• It enables high-throughput and semi-automated image quantification of capillary density.
• It provides a robust analysis for determining capillary density over the entire muscle cross section.
Graphical overview
Keywords: Capillary density Image quantification Skeletal muscle Immunofluorescence CD31 Endoglin
Background
Capillaries in skeletal muscles play a vital role in the delivery of oxygen and nutrients essential for muscle metabolism and contraction, both at rest and during exercise. Capillary density, which refers to the number of capillaries in a given myofiber area, is critical for estimating oxygen consumption and determining exercise capacity in athletes, the elderly, and patients with muscle-related pathologies. Low capillary density in skeletal muscles is an indicator of reduced oxidative metabolism (Duscha et al., 2020). Conversely, a higher capillary density shortens the distance for oxygen diffusion, leading to improved muscle performance (Gliemann, 2016). Importantly, capillary density can adapt to different conditions and stimuli. For example, endurance training increases muscle capillary density, whereas physical or medical conditions associated with muscle disuse can negatively affect capillary density (Lemieux and Birot, 2021). Thus, determining capillary density is a key measure for assessing changes in skeletal muscle physiology and evaluating the exercise potential of skeletal muscles.
Capillary density is defined as the number of capillaries per unit of muscle cross-sectional area in a muscle biopsy (McGuire and Secomb, 2003; Abbassi-Daloii et al., 2023b). This only takes into account the myofibers within the muscle tissue, and therefore excludes fibrotic regions. Determination of capillary density is essential for estimating oxygen consumption and blood flow in skeletal muscles. It involves immunohistochemistry in muscle cross sections using antibodies specific for proteins expressed in endothelial cells, such as CD31 and/or CD105 (endoglin) (Pestronk et al., 2010; Duscha et al., 2020). In some protocols, capillaries are stained with Ulex europaeus agglutinin (Hendrickse et al., 2022), which stains lectins (N-glycans) and is used as a marker for endothelial cells (Holthöfer et al., 1982). Most protocols for measuring capillary density rely on manual, eye-based evaluation of fluorescence-stained muscle tissue, as shown in examples such as Andersen (1975), Duscha et al. (2020), and Baum et al. (2023). However, eye-based image scoring has limitations, due to its susceptibility to bias, time-consuming nature, and low throughput, resulting in reduced reproducibility and less robust results. Alternative procedures use image quantification, but these are also low throughput and cover only a small part of the muscle cross-section, leading to a spatial bias (Hendrickse et al., 2022). High-throughput semi-automated imaging and image quantification of the entire muscle cross section overcomes these limitations. We recently reported on a large study of human skeletal muscles that required high-throughput imaging and image analysis of immunohistochemistry in skeletal muscles (Abbassi-Daloii et al., 2023b). While we have previously presented a high-throughput protocol for myofiber typing (Abbassi-Daloii et al., 2023a), here, we present a high-throughput protocol for assessing capillary density in skeletal muscles. A flowchart summarizing the steps implemented in this protocol is shown in Figure 1.
Figure 1. Flowchart of the main steps in the protocol. In the immunostaining step, an anti-laminin antibody marks the cell membrane, while anti-CD31 and anti-CD105 antibodies mark epithelial cells.
Materials and reagents
Biological materials
Snap-frozen human skeletal muscle biopsy
Reagents
Antibodies
Anti-human CD31-Alexa Fluor® 594-conjugated; dilution 1:400 (BioLegend, catalog number: 303126)
Rabbit anti-laminin; dilution 1:2,000 (Sigma-Aldrich, catalog number: L9393)
Goat anti-rabbit Alexa Fluor® 750-conjugated; dilution 1:1,000 (Thermo Fisher Scientific, catalog number: A21039)
Optional:
Anti-human CD105 (endoglin, ENG) biotin-conjugated; dilution 1:100 (BioLegend, catalog number: 323214)
Streptavidin-Alexa Fluor® 647-conjugated; dilution 1:500 (Life Technologies, catalog number: S21374)
Chemicals
OCT Embedding matrix for frozen sections (Tissue-Tek) (VWR, part of Avantor, catalog number: 361603E)
NaCl (Sigma-Aldrich, catalog number: 7647-14-5)
Na2HPO4·2H2O (Sigma-Aldrich, catalog number: 10028-24-7)
KCl (Sigma-Aldrich, catalog number: 7447-40-70)
KH2PO4 (Sigma-Aldrich, catalog number: 7778-77-0)
Tween 20 (Sigma-Aldrich, catalog number: 9005-64-5)
Skim milk powder (FrieslandCampina)
DAPI (Sigma-Aldrich, catalog number: 28718-90-3)
ProLongTM Gold antifade mountant (ThermoFisher Scientific, catalog number: P10144)
Nail polish
Solutions
Phosphate-buffered saline (PBS) (see Recipes)
Phosphate-buffered saline containing 0.05% tween (PBST) (see Recipes)
PBST + 5% milk (see Recipes)
Recipes
PBS 10×
Reagent Final concentration Amount
NaCl N/A 80 g
Na2HPO4·2H2O N/A 15 g
KCl N/A 2 g
KH2PO4 N/A 1.2 g
Distilled water N/A up to 1 L
Total 10× 1 L
PBST
Reagent Final concentration Amount
Tween 20 0.05% 0.5 mL
PBS 1× 999.5 mL
Total N/A 1 L
PBST + 5% milk
Reagent Final concentration Amount
Milk powder 5% 2.5 mg
PBST 1× 50 mL
Equipment
Coverslip (Menzel-Glaser, catalog number: 631-1365)
ZEISS Axio Scan.Z1 (Carl Zeiss Microscopy GmbH, model: Axioscan 7)
EprediaTM SuperFrostTM microscope slides, ground 90° (Thermo Fisher Scientific, catalog number: 12372098)
A-PAP pen liquid-blocker (immunopen) (Cosmo Bio, model: DAI-APAP-R)
Straight tweezers
Cryostat (Leica Biosystems, model: CM3050 S)
Glass insert 70 mm wide for anti-roll systems (Leica Biosystems, catalog number: 14047742497)
EprediaTM MX35 PremierTM disposable low-profile microtome blades (Thermo Fisher Scientific, catalog number: 3052835)
Software and datasets
ZEN 2 (Carl Zeiss, https://www.zeiss.com/microscopy/en/products/software/zeiss-zen.html)
Fiji (Schindelin et al. (2012), https://imagej.net/Fiji)
R v4.0.2 (R Core Team (2013), https://www.r-project.org/)
RStudio v1.3.959 (Allaire (2012), https://www.rstudio.com/)
R scripts (https://github.com/tabbassidaloii/ImageProcessing/blob/main/CapillaryDensity/Rscript)
Macros (https://github.com/tabbassidaloii/ImageProcessing/tree/main/CapillaryDensity/Macros)
Procedure
Cryosection of skeletal muscle biopsies
The procedure is detailed in Abbassi-Daloii et al. (2023a). In brief, this step entails the preparation of muscle biopsies for histology and immunofluorescence staining. Following the cleaning of equipment and temperature adjustments, the muscle biopsies are equilibrated inside the cryostat. Subsequently, biopsies are embedded in Tissue-Tek, placed on specimen holders, and cryosections of a specified thickness (10–16 μm) are collected onto SuperFrost slides. Store slides at -20 °C or -80 °C prior to immunostaining.
Immunofluorescence
This step describes immunofluorescence staining using antibodies for CD31 and laminin. Adding anti-CD105 as a second marker for endothelial cells is optional. The antibodies are prepared in PBST.
Air dry slides from -20 °C for 30 min at room temperature (RT).
Outline each section with an immunopen approximately 2–3 mm from the tissue edge. This reduces the required volume of the antibody mix.
Note: Do not draw the line too close to the muscle sections as it can introduce artifacts in the image processing step.
Wash the sections in PBST.
Blocking: Cover each section with PBST + 5% milk (~40 μL) for 30 min at RT.
Wash the slides three consecutive times with a large volume of PBST (~40 μL), each time for 5 min.
Primary antibody incubation: Cover each section with 20 μL of antibody mix containing anti-human CD31-Alexa Fluor® 594-conjugated, rabbit anti-laminin, and anti-human CD105 biotin-conjugated (optional). Incubate for 2 h at RT.
Note: Keep slides in the dark from this step onwards.
Wash the slides three consecutive times with an excessive volume of PBST, each time for 5 min.
Secondary antibody incubation: Incubate sections with 20 μL of mixture of the following secondary antibodies for 1 h at RT: goat anti-rabbit Alexa Fluor® 750-conjugated to detect anti-laminin and Streptavidin-Alexa Fluor® 647-conjugated to detect anti-CD105 (optional).
Wash the slides three consecutive times with an excessive volume of PBST, each time for 5 min.
Nuclei counterstain is carried out by a short incubation (5–10 min) of the section with a DAPI solution (1:1,000 dilution in PBST, ~20 μL per section) in a dark environment. Afterward, gently rinse the sections with PBST to remove excess DAPI solution. DAPI binds to nucleic acids and stains the chromatin.
Mounting: Cover the sections with ProLongTM Gold antifade mountant (~10 μL per section). Cover the slide with a coverslip and fix it with nail polish.
Note: Avoid any air bubbles on the sections as they will affect the image acquisition.
Keep for 24 h at RT in the dark prior to imaging.
Store slides at 4 °C prior to imaging.
Note: Slides can be kept at 4 °C for one month but imaging a week after immunostaining is preferable.
Image acquisition
The image acquisition is detailed in Abbassi-Daloii et al. (2023a). In brief, we utilized a Zeiss Axio Scan.Z1 slide scanner with the ZEN 2 software. Per fluorophore, exposure and intensity were determined to maximize signal-to-noise ratio without bleaching. Imaging was carried out with a 10×/0.45 Plan-Apochromat objective. For high-throughput imaging, we recommend image acquisition with a slide scanner with a stitching option.
Note: It is crucial to optimize the imaging settings on a test slide to determine the appropriate exposure time and intensity for each fluorophore. Adjustments of exposure time and focusing algorithms may affect the visibility of the fluorophore signal. To achieve the best signal-to-noise ratio without causing bleaching, it is necessary to optimize the intensity and exposure time for each fluorophore/channel.
For all channels, utilize single band filters with the following excitation ranges:
Channel 1 (DAPI): 335–355 nm excitation
Channel 2 (CD31, Alexa Fluor® 594): 574–599 nm excitation
Channel 3 (CD105, Alexa Fluor® 647): 650–670 nm excitation (optional)
Channel 4 (Laminin, Alexa Fluor® 750): 672–747 nm excitation
Notes:
It is crucial to maintain consistent image acquisition settings for all slides throughout different batches. This ensures uniformity and allows for reliable comparisons between samples.
To eliminate batch effect, we recommend staining all samples in one batch and then imaging all slides in one session. When staining samples in multiple batches, it is important to conduct imaging in the same order of batches. This consistency ensures that the time interval between staining and imaging remains constant across all batches, promoting accurate and comparable results.
When employing the Axio Scan.Z1 slide scanner, each slide's output will be in the Carl Zeiss Image format (CZI) dataset, containing an image for each section of the slide.
Image processing
Laminin segmentation and object quantification
The procedure for converting image format and laminin segmentation is carried out in ImageJ/Fiji, using five sequential macros. The macros are found in: (https://github.com/tabbassidaloii/ImageProcessing/tree/main/CapillaryDensity/Macros/), macros 2–4. The macros are fully automatic, besides macro number 3, which might require a manual adjustment (as explained in the macro and in Abbassi-Daloii et al., 2023a). The outputs from the five macros are collected in a folder named “check” with mask images after each step, and a folder “ROI” with .txt files reporting the area of the segmented laminin objects that will be used for the calculation of capillary density.
An example of laminin staining and segmentation is in Figure 2.
Figure 2. Visualization of capillary mask and capillary output generation. The images show the entire cross section with a zoom-in insert in the right bottom of each image. Red arrows point to laminin regions that were excluded from the mask after segmentation. Green arrows point to CD31 objects that did not overlap with laminin and were therefore excluded from the capillary mask. The capillary mask is used to obtain CD31 intensity and CD105 intensity.
Segmentation and quantification of CD31 and CD105 objects
This step is executed using macro number 6. We perform the segmentation of the CD31 signal and compute the intersection with the laminin segmentation; these objects are considered as capillary. The output files contain the mean fluorescence intensity, area, and circularity of the capillary objects for both CD31 and CD105.
A mask is made on CD31 images using a Gaussian Blur filter to reduce noise, resulting in a smoother image.
Thresholding is applied using the "Li dark" algorithm to convert the CD31 channel into a binary image that distinguishes between foreground and background pixels.
The Watershed algorithm is then utilized to accurately separate cells that may be touching or overlapping.
The Fill Holes algorithm is employed to fill any empty spaces within the segmented regions.
The laminin segmentation mask is added to identify the overlap between laminin and CD31 objects, resulting in a capillary mask.
Once the capillaries have been segmented, the mean fluorescence intensity is measured in the original CD31 and CD105 channels.
The intensity, circularity, and area of CD31 and CD105 are in the output file that is saved in a folder named “segmentation.” An example of CD31 and CD105 staining, CD31 segmentation, and capillary mask are in Figure 2.
Note: Steps outlined previously can be conveniently executed by running two Windows Batch Files available on GitHub. These files are specifically designed to automate the process, allowing for a streamlined and efficient implementation of the protocol.
https://github.com/tabbassidaloii/ImageProcessing/tree/main/CapillaryDensity/Macros/BatchFiles
Capillary density quantification
In this step, we assess the filtering procedures and quantify capillary density. These analyses are carried out in the RStudio software along with the R statistical software. To facilitate the execution of these steps, we have made the R Markdown file accessible on GitHub: https://github.com/tabbassidaloii/ImageProcessing/blob/main/MyofiberTyping/Rscript/CapillaryDensity.Rmd Within the following steps, we indicate the specific R code chunk within this R markdown file that should be utilized for each task.
Calculate the total myofiber area:
Consolidate myofiber data from all samples by running the "myofibersTotalArea" R code chunk.
Count the number of segmented objects (myofibers) per sample.
Compute the total myofiber area per sample.
Retain the replicate with the highest number of myofibers.
Exclude samples with a small number of myofibers (<100).
Calculate the total CD31 positive area:
Consolidate CD31 and CD105 quantification data from all samples by running the "positiveCD31Area" R code chunk.
Compute and plot the total proportion of CD31 positive area per sample.
Calculate capillary density:
Apply filtering for capillaries by running “capillaryDensity” R chunk code:
• Select objects with positive signals for both CD31 and CD105.
• Choose objects larger than 3 µm2 and smaller than 51 µm2.
• Include objects with circularity larger than 0.5.
Compute and plot capillary density as the number of capillaries per unit (µm) of the total myofiber area.
Note: The capillary density can be alternatively calculated as the number of capillaries per myofibers, which are computed in the "myofibersTotalArea" R code chunk.
Validation of protocol
The procedures outlined in this protocol have been used in the following research article: Abbassi-Daloii et al. (2023b) (Figure 4).
General notes and troubleshooting
General notes
This procedure requires a basic understanding of immunofluorescence, fluorescence imaging, image processing in ImageJ, and data analysis in R.
For basics in immunofluorescence, please refer to the following paper: Im et al. (2019).
For basics in fluorescence imaging, you can refer to the following source: Ogundele et al. (2013).
The basics of using ImageJ can be found in the "ImageJ User Guide" document, available at: https://imagej.nih.gov/ij/docs/guide/user-guide.pdf.
If you encounter problems running the macros, please contact us.
The definition of capillary objects is based on three markers: CD31 & laminin overlap and overlap with CD105. To reduce costs, it is possible to omit CD105. In our experience, CD31 staining was more specific compared with CD105.
When assessing changes in capillary density between tissues or pathological conditions, aim for at least 1,000 capillary masks per sample.
While this procedure is described for human skeletal muscle cross sections, it can be applied to animal models (i.e., mouse, rat). The antibodies we specify should be tested for cross-reactivity in other models. We recommend testing the antibodies before conducting a large experiment.
Acknowledgments
Funding: Association Française centre les Myopathies (AFM Telethon; Grant # 22506).
We acknowledge Hermien E Kan and Peter AC 't Hoen and the funding agency that contributed to the source paper (Abbassi-Daloii et al., 2023b).
Competing interests
All authors declare no competing interests.
Ethical considerations
The muscles used in this study were collected in accordance with an approved ethical protocol. The study received approval from the local Medical Ethical Review Board of The Hague Zuid-West and the Erasmus Medical Centre. The research was conducted under the ethical standards stated in the 1964 Declaration of Helsinki and its later amendments (ABR number: NL54081.098.16). Informed consent was obtained from all subjects, as described in Abbassi-Daloii et al. (2023b).
References
Abbassi-Daloii, T., el Abdellaoui, S., Kan, H. E., van den Akker, E., ’t Hoen, P. A., Raz, V. and Voortman, L. M. (2023a). Quantitative analysis of myofiber type composition in human and mouse skeletal muscles. STAR Protoc. 4(1): 102075. doi: 10.1016/j.xpro.2023.102075
Abbassi-Daloii, T., el Abdellaoui, S., Voortman, L. M., Veeger, T. T., Cats, D., Mei, H., Meuffels, D. E., van Arkel, E., 't Hoen, P. A., Kan, H. E., et al. (2023b). A transcriptome atlas of leg muscles from healthy human volunteers reveals molecular and cellular signatures associated with muscle location. eLife 12: e80500. doi: 10.7554/elife.80500
Allaire, J. (2012). RStudio: integrated development environment for R. Boston, MA 770(394): 165–171.
Andersen, P. (1975). Capillary Density in Skeletal Muscle of Man. Acta Physiol. Scand. 95(2): 203–205. doi: 10.1111/j.1748-1716.1975.tb10043.x
Baum, O., Huber-Abel, F. A. M. and Flück, M. (2023). nNOS Increases Fiber Type-Specific Angiogenesis in Skeletal Muscle of Mice in Response to Endurance Exercise. Int. J. Mol. Sci. 24(11): 9341. doi: 10.3390/ijms24119341
Duscha, B. D., Kraus, W. E., Jones, W. S., Robbins, J. L., Piner, L. W., Huffman, K. M., Allen, J. D. and Annex, B. H. (2020). Skeletal muscle capillary density is related to anaerobic threshold and claudication in peripheral artery disease. Vasc. Med. 25(5): 411–418. doi: 10.1177/1358863x20945794
Gliemann, L. (2016). Training for skeletal muscle capillarization: a Janus-faced role of exercise intensity? Eur. J. Appl. Physiol. 116(8): 1443–1444. doi: 10.1007/s00421-016-3419-6
Hendrickse, P. W., Wüst, R. C., Ganse, B., Giakoumaki, I., Rittweger, J., Bosutti, A. and Degens, H. (2022). Capillary rarefaction during bed rest is proportionally less than fibre atrophy and loss of oxidative capacity. J. Cachexia Sarcopenia Muscle 13(6): 2712–2723. doi: 10.1002/jcsm.13072
Holthöfer, H., Virtanen, I., Kariniemi, A. L., Hormia, M., Linder, E. and Miettinen, A. (1982). Ulex europaeus I lectin as a marker for vascular endothelium in human tissues. Lab Invest 47(1): 60–66. http://europepmc.org/abstract/MED/6177923
Im, K., Mareninov, S., Diaz, M. F. P. and Yong, W. H. (2019). An Introduction to Performing Immunofluorescence Staining. Methods Mol. Biol.: 299–311. doi: 10.1007/978-1-4939-8935-5_26
Lemieux, P. and Birot, O. (2021). Altitude, Exercise, and Skeletal Muscle Angio-Adaptive Responses to Hypoxia: A Complex Story. Front. Physiol. 12: e735557. doi: 10.3389/fphys.2021.735557
McGuire, B. J. and Secomb, T. W. (2003). Estimation of capillary density in human skeletal muscle based on maximal oxygen consumption rates. Am. J. Physiol. Heart Circ. Physiol. 285(6): H2382–H2391. doi: 10.1152/ajpheart.00559.2003
Ogundele, O. M. A., Adekeye, A.O., Adeniyi, P. A., , Ogedengbe, O. O., Enye, L. A., Saheed, S. and Omotosho, D. R. (2013). Basic principles of fluorescence microscopy. World Journal of Young Researchers 3(1): 17–22.
Pestronk, A., Schmidt, R. E. and Choksi, R. (2010). Vascular pathology in dermatomyositis and anatomic relations to myopathology. Muscle Nerve 42(1): 53–61. doi: 10.1002/mus.21651
R Core Team, R. (2013). R: A language and environment for statistical computing.
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. doi: 10.1038/nmeth.2019
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Methodology to Create Auxin-Inducible Degron Tagging System to Control Expression of a Target Protein in Mammalian Cell Lines
AR Amit Rahi
DS Deepika K. Sodhi
CM Christine B. Magdongon
RS Rajina Shakya
DV Dileep Varma
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4923 Views: 1343
Reviewed by: Rajesh RanjanBhuvanasundar RanganathanShashi Kumar Suman
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Jun 2023
Abstract
The auxin-inducible degron (AID) system is a versatile tool in cell biology and genetics, enabling conditional protein regulation through auxin-induced degradation. Integrating CRISPR/Cas9 with AID expedites tagging and depletion of a required protein in human and mouse cells. The mechanism of AID involves interactions between receptors like TIR1 and the AID tag fused to the target protein. The presence of auxin triggers protein ubiquitination, leading to proteasome-mediated degradation. We have used AID to explore the mitotic functions of the replication licensing protein CDT1. Swift CDT1 degradation via AID upon auxin addition achieves precise mitotic inhibition, revealing defects in mitotic spindle structure and chromosome misalignment. Using live imaging, we found that mitosis-specific degradation of CDT1 delayed progression and chromosome mis-segregation. AID-mediated CDT1 inhibition surpasses siRNA-based methods, offering a robust approach to probe CDT1’s mitotic roles. The advantages of AID include targeted degradation and temporal control, facilitating rapid induction and reversal of degradation—contrasting siRNA’s delayed RNA degradation and protein turnover. In summary, the AID technique enhances precision, control, and efficiency in studying protein function and regulation across diverse cellular contexts. In this article, we provide a step-by-step methodology for generating an efficient AID-tagging system, keeping in mind the important considerations that need to be adopted to use it for investigating or characterizing protein function in a temporally controlled manner.
Key features
• The auxin-inducible degron (AID) system serves as a versatile tool, enabling conditional protein regulation through auxin-induced degradation in cell biology and genetics.
• Integration of CRISPR/Cas9 knock-in technology with AID expedites the tagging and depletion of essential proteins in mammalian cells.
• AID’s application extends to exploring the mitotic functions of the replication licensing protein CDT1, achieving precise mitotic inhibition and revealing spindle defects and chromosome misalignment.
• The AID system and its diverse applications advance the understanding of protein function and cellular processes, contributing to the study of protein regulation and function.
Graphical overview
Cdt1–auxin-inducible degron (AID) tagging workflow. (A) Schematic of the cloned Cdt1 gRNA vector and the repair template generated to endogenously tag the Cdt1 genomic locus with YFP and AID at the C-terminal using CRISPR/CAS9-based genome editing. The two plasmids are transfected into DLD1-TIR1 stable cells, followed by sorting and scaling up of YFP-positive single cells. (B) The molecular mechanism of auxin-induced proteasome-mediated degradation of the target protein (CDT1) shown at the bottom of the figure is well worked out.
Keywords: Auxin-inducible degron (AID) Protein regulation Auxin-induced degradation CRISPR/Cas9 Protein tagging Protein depletion TIR1 receptor Proteasomal degradation Mitotic functions CDT1
Background
Conditional protein degradation is an invaluable approach to understand cellular function. Among the various techniques available, the auxin-inducible degron (AID) system stands out. AID is a versatile molecular tool extensively utilized in cell biology and molecular genetics for the conditional destabilization of target proteins, facilitated by CRISPR/Cas9 gene knock-in technology [1, 2]. This mechanism capitalizes on the unique ability of the plant hormone auxin to rapidly degrade specific proteins bearing an AID sequence not only in non-plant cells like DLD1, HCT116, and HeLa but also in organisms like Caenorhabditis elegans, mouse, and yeast [3–6]. The AID assay furnishes a valuable means for probing protein functions and regulating protein expression in live cells. Its precise control over protein degradation addresses limitations of other methods such as RNAi gene silencing, which is both slow and less specific, lacking conditionality. The AID system, on the other hand, operates at the protein level, offering simplicity, rapidity, effectiveness, and reversibility. Central to this system is the interaction between the transport inhibitor response 1 receptor (TIR1), an F-box related protein, and the AID tag (available as 25 or 7 kDa), genetically fused to the protein of interest [7–9]. The half-life degradation of the tag, which is approximately 30 min, can be observed in a mammalian cell line expressing TIR1. Upon auxin binding to the TIR1 receptor, a conformational change occurs, allowing TIR1 to bind to the AID tag on the target protein. This interaction triggers the activation of the SCF (Skp1-Cullin-F-box) E3 ubiquitin ligase complex, comprising Skp1, Cul1, and RBX1. RBX1 associates with the E2 ubiquitin ligase, facilitating the ubiquitination of the protein of interest and its subsequent proteasomal degradation (26S proteasome). This system is advantageous as it leads to rapid degradation of the protein of interest in the presence of auxin, and its removal restores the phenotype [10, 11].
In our study, we employed the AID system as an innovative approach to investigate the mitotic functions of the replication licensing protein, CDT1. Through the AID system, we achieved precise and controlled inhibition of CDT1 during mitosis by inducing rapid degradation through auxin addition [12]. This strategy unveiled significant defects in mitotic spindle structure and chromosome alignment in treated cells. The system’s simplicity, control, and reversibility render it highly valuable for studying essential proteins and unraveling their roles in various cellular processes.
Materials and reagents
Cas9-gRNA expression vector (e.g., pX330, PX458, PX459)
Parent repair template (a gift from Dan Foltz Lab)
Sense oligo for cloning of sg at BbsI site (5'-CACCG [sgRNA Target Sequence]-3')
Antisense oligo for cloning at BbsI site (5'-AAAC [reverse complement sgRNA Target Sequence] C-3')
Biological materials
DLD1-TIR1 cells, colorectal adenocarcinoma cell line (a kind gift from Andrew Holland, Johns Hopkins University, Baltimore, MD, USA)
One ShotTM Stbl3TM chemically competent E. coli (Thermo Fisher Scientific, catalog number: C737303)
Reagents
Dulbecco's modified Eagle's medium (DMEM) (Corning, catalog number: 10-013-CV)
Penicillin-Streptomycin 10,000 U/mL (Thermo Fisher Scientific, catalog number: 15140122)
Trypsin-EDTA, 1× (CORNING, catalog number: 25-052-CI)
Phosphate-buffered saline (PBS 1×) (CORNING, catalog number: 21-040-CV)
Effectene transfection agent (Qiagen, catalog number: 301425)
Dimethyl sulfoxide (DMSO) (Fisher Bio ReagentsTM, catalog number: BP231-100)
Auxin (Indole-3-acetic acid, IAA) (Sigma-Aldrich, catalog number: 15148-2G)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7906)
Anti-GFP mouse monoclonal (Thermo Fisher Scientific, catalog number: A-11120)
Anti-Cdt1 rabbit polyclonal H-300 (Santa Cruz Biotech, catalog number: sc-28262)
Anti-Tubulin anti-mouse monoclonal DM1A (Santa Cruz Biotech, catalog number: sc-32293)
HRP-conjugated rabbit/mouse antibodies (Azure Biosystems, catalog numbers: AC2114 and AC2115)
SuperSignalTM West Pico PLUS Chemiluminescent Substrate (Thermo Scientific, catalog number: 34580)
Plasmid Isolation kit (Qiagen, catalog number: 27104)
Gel Extraction kit (Plasmid and PCR clean-up kit) (Qiagen, catalog number: 28704)
Quick-DNA Microprep kit (Zymo Research, catalog number: D3020)
PCR Master Mix (2×) reagents (Thermo Scientific, catalog number: K0171)
GeneRuler 1 kb DNA ladder (Thermo Scientific, catalog number: SM0311)
TriTrack DNA loading dye (6×) (Thermo Scientific, catalog number: R1161)
T4 DNA ligase enzyme (NEB, catalog number: M0202S)
10× buffer for T4 DNA ligase (NEB, catalog number: B0202S)
Bbs1 enzyme (NEB, catalog number: R0539S)
Ethidium bromide (Bio-Rad, catalog number: 161-0433)
Agarose HS (Denville Scientific Inc., catalog number: CA3510-8)
Ethylenediaminetetraacetic acid (EDTA) (Sigma-Aldrich, catalog number: 03620)
Sodium dodecyl sulphate (SDS) (Fisher Scientific, catalog number: BP166-500)
Tris-base (ChemCruz, catalog number: 77-86-1)
Ammonium persulfate (APS) (Sigma-Aldrich, catalog number: A3678)
TEMED (GE Healthcare, catalog number: 17-1312-01)
Acrylamide/Bis-acrylamide, 30% solution (Sigma-Aldrich, catalog number: A3574)
6× Laemmli SDS sample buffer (Bioland Scientific LLC, catalog number: SAB03-01)
Pierce IP lysis buffer (Thermo Scientific, catalog number: 87787)
Halt protease inhibitor (100×) (Thermo Scientific, catalog number: 87786)
Acetic acid, glacial (Fisher Chemical, catalog number: A38-212)
Glycine 99% (Thermo Scientific, catalog number: A13816.36)
Sodium chloride (NaCl) (Fisher Chemical, catalog number: S271-1)
Magnesium chloride (MgCl2) (Sigma-Aldrich, catalog number: 208337-1KG)
S.O.C. medium (Invitrogen, catalog number: 15544-034)
Ampicillin sodium salt (Fisher Scientific, catalog number: 69-52-3)
Luria broth base (Miller's LB broth base), powder (Invitrogen, catalog number: 12795027)
LB agar (Lennox L Agar), powder (Invitrogen, catalog number: 22700025)
Solutions
1 M Auxin (IAA) solution in Milli-Q water
50× TAE buffer (see Recipes)
1% agarose gel (see Recipes)
SDS-PAGE 10% resolving protein gel (5 mL)
SDS-PAGE 5% stacking protein gel (see Recipes)
Buffers for SDS-PAGE (see Recipes)
100 mg/mL Ampicillin stock solution
Recipes
50× TAE buffer
Reagent Quantity or Volume
Tris-base 242 g
Acetic acid 57.1 mL
EDTA 100 mL (0.5 M, 8.0 pH)
Milli-Q water Adjust volume to 1 L
Total 1,000 mL
Note: Dilute 50× TAE to 1× TAE (1:50) with Milli-Q water when making agarose gel and running the gel.
1% agarose gel (100 mL)
Reagent Final concentration Quantity or Volume
Agarose powder 1% 1 g
1× TAE N/A 100 mL
Ethidium bromide 0.5 µg/mL 2–5 µL
Total 100 mL
In a microwave-safe flask, add 1 g of agarose powder to 100 mL of 1× TAE buffer.
Microwave the flask in short 20–30 s intervals, swirling between each interval. Bring the mixture to a boil until the agarose has completely dissolved and the solution is transparent. Be careful as to not let the solution overboil and evaporate.
Allow mixture to cool before adding 5 µL of ethidium bromide for a 0.5 µg/mL concentration from 10 mg/mL stock. Swirl flask to mix.
Pour solution into casting gel tray and insert well comb. Allow gel to solidify at room temperature (RT).
SDS-PAGE 10% resolving protein gel (5 mL)
Reagent Stock concentration Volume
Milli-Q water N/A 1.9 mL
Acrylamide mix 30% 1.7 mL
Tris-base 1.5 M, 8.8 pH 1.3 mL
SDS 10% 50 µL
APS 10% 50 µL
TEMED N/A 2 µL
Total 5 mL
SDS-PAGE 5% stacking protein gel (2.0 mL)
Reagent Stock concentration Volume
Milli-Q water N/A 1.4 mL
Acrylamide mix 30% 0.33 mL
Tris-base 1.0 M, 6.8 pH 0.22 mL
SDS 10% 20 µL
APS 10% 20 µL
TEMED N/A 2 µL
Total 2 mL
Buffers for SDS-PAGE
Reagent Running buffer (1×) Transfer buffer (1×)
Glycine 14.4 g 2.9 g
Tris-base 3.03 g 5.8 g
SDS 1.0 g 0.33 g
Milli-Q water 1,000 mL 800 mL
Methanol N/A 200 mL
Total 1,000 mL 1,000 mL
Laboratory supplies
96-, 24-, 12-, and 6-microwell plates (Thermo Fisher Scientific, catalog number: 130188)
15 mL conical Falcon tube (Fisher Brand, catalog number: 07-200-886)
1.5 mL Eppendorf tubes (Fisher Scientific, catalog number: 02-682-002)
BioLiteTM cell culture treated dishes, 35, 60, and 100 mm Petri dish (Thermo Scientific, catalog numbers: 130180, 130181, and 130182)
WhatmanTM UnifloTM sterile PVDF syringe filters 0.22 µm (Cytiva, catalog number: 9913-2502)
Sterile polystyrene disposable serological pipettes (Fisher Brand, catalog number: 13-678-11E)
Flow cytometry tubes, Mini 35 µm (Olympus Plastics, catalog number: 28-154)
Sterile glass spreader
Equipment
Centrifuge (Eppendorf, model: 5417C and 5804R)
Thermocycler (Eppendorf, model: GX2)
NanoDrop One (Thermo Scientific, model: Nanodrop One Spectrophotoshot Promo)
Flow cytometer (BD-FACS, model: Melody Cell Sorter)
Western blot developer (Azure biosystems, model: 600)
Agarose gel electrophoresis equipment (Bio-Rad, model: Mini-Sub Cell GT)
Protein gel electrophoresis equipment (Bio-Rad, model: Mini PROTEAN Tetra cell)
Spectrophotometer (Vinmax, model: 721-VIS)
Brightfield cell counter (DeNovix, model: CellDrop BF PAYG)
Inverted microscope equipped with a Yokogawa CSU-X1 spinning disc, an Andor iXon Ultra888 EMCCD camera, and a 60× or 100× 1.4 NA Plan-Apochromatic DIC oil immersion objective (Nikon, model: Eclipse TiE)
Software and datasets
Broad Institute sgRNA design tool (https://portals.broadinstitute.org/gpp/public/analysis-tools/sgrna-design). Alternatively, you can use CHOPCHOP (http://chopchop.cbu.uib.no/) or Crispor (http://crispor.tefor.net/crispor.py)
Procedure
Designing the guide RNA (gRNA)
Download the genomic DNA sequence from NCBI for your gene of interest. In this case, this is Cdt1. Go to website https://www.ncbi.nlm.nih.gov/gene/81620 and http://useast.ensembl.org/Homo_sapiens/Gene/Summary?g=ENSG00000167513;r=16:88803789-8880
Open the full-length gene, including both introns and exons, and locate the translation start (ATG) and stop site (TGA) based on the position in the RefSeq (which is NCBI database).
For N-terminal tagging, select and copy approximately 15 base pairs upstream of the ATG and approximately 150 base pairs downstream from the ATG. This sequence window will provide options to choose the best gRNA near to the point of modification.
For C-terminal tagging, select and copy approximately 15 base pairs downstream of the TGA (stop codon) and approximately 150 base pairs upstream from the stop codon as stated earlier.
Paste the selected sequence into the Crispor or/and CHOPCHOP sgRNA design tool, provided by Broad institute. Do not forget to select appropriate organism genome, which provides SG score, off-target, and valuable parameters.
Download the results of the sgRNA design as a .txt file and import it into a spreadsheet.
Review the sgRNA results obtained from the design tool and prioritize guides that cut within 50 base pairs after the ATG (for N-terminal modification) or close to Stop codon (for C-terminal modification), giving preference to those with none or fewer off-targets.
Focus on specific columns, such as "Position of Base After Cut (1-based)," "sgRNA Target Sequence," "On-Target Rank," and "Off-Target Rank" to select guides with better scores.
Pick at least three high-ranking guides for the cloning into SG backbone plasmid (Table 1).
Table 1. Top rank sgRNA score table
sgRNA # Orientation sgRNA cut position sgRNA sequence sgRNA context sequence PAM sequence On-target rank Off-target rank Combined rank
1 Sense 168 GGCCCACCAGACACGTGCTG GCCTGGCCCACCAGACACGTGCTGAGGAGG AGG 3 5 1
2 Sense 173 ACCAGACACGTGCTGAGGAG GCCCACCAGACACGTGCTGAGGAGGGGCTG GGG 7 9 3
3 Sense 171 CCACCAGACACGTGCTGAGG TGGCCCACCAGACACGTGCTGAGGAGGGGC AGG 13 15 11
Cloning of SG sequence at Bbs1 site into pX330 vector to generate sgRNA constructs
Add the Bbs1 restriction enzyme overhang on 5′ and 3′ ends of the sgRNA sequence.
Upstream cut: 5′ (CACCGG)GTCTTC, 3′ (CC)CAGAAG
Downstream cut: 5′ GAAGAC(CT) 3′, 3′ CTTCTG(GACAAA) 5′
Select sgRNA sequences based on the target region: order oligos for sense and antisense strands, including the sgRNA target sequence and its reverse complement (Table 2 and Table 3).
Sense oligo: 5′-CACCG [sgRNA Target Sequence]-3′
Antisense oligo: 5′-AAAC [reverse complement sgRNA Target Sequence] C-3′
Order the oligos from your company of choice (e.g., IDT) and resuspend them in autoclaved Milli-Q water at a final concentration of 100 µM.
Note: The sequences of SG for Cdt1 were reconfirmed using NCBI blast feature, and the first hit for all three sequences was the Cdt1 gene from Human.
Table 2. Primer list of top three sgRNAs with Bbs1-compatible cohesive ends
sgRNA# sgRNA sequence sgRNA with overhangs
1 GGCCCACCAGACACGTGCTG
Sense 5′ CACCGGGCCCACCAGACACGTGCTG 3′
antisense 3′ CCCGGGTGGTCTGTGCACGACCAAA 5’
2 ACCAGACACGTGCTGAGGAG
Sense 5′ CACCGACCAGACACGTGCTGAGGAG 3′
antisense 3′ CTGGTCTGTGCACGACTCCTCCAAA 5′
3 CCACCAGACACGTGCTGAGG
Sense 5′ CACCGCCACCAGACACGTGCTGAGG 3′
antisense 3′ CGGTGGTCTGTGCACGACTCCAAA 5′
Table 3. Primer list of top three sgRNAs containing Bbs1-compatible cohesive end ready for ordering
sgRNA # sgRNA sequence Forward and reverse primer sequences
1 GGCCCACCAGACACGTGCTG
FP- 5′ CACCGGGCCCACCAGACACGTGCTG 3′
RP- 5′ AAACCAGCACGTGTCTGGTGGGCCC 3′
2 ACCAGACACGTGCTGAGGAG
FP- 5′ CACCGACCAGACACGTGCTGAGGAG 3′
RP- 5′ AAACCTCCTCAGCACGTGTCTGGTC 3′
3 CCACCAGACACGTGCTGAGG
FP- 5′ CACCGCCACCAGACACGTGCTGAGG 3′
RP-5′ AAACCCTCAGCACGTGTCTGGTGGC 3′
Order the CRISPR/Cas9-EGFP sgRNA vector (pX330) and prepare it for cloning.
Plasmid details: pX330, plasmid size: 8,484 bp (https://www.addgene.org/42230/). Dissolve 8 µg of pX330 in 16 µL of Milli-Q water to achieve a final concentration of 0.5 µg/µL.
Linearize the pX330 plasmid by BbsI enzyme.
Milli-Q water 27.00 µL
10× NE buffer 5 µL
BbsI enzyme 2 µL
pX330 backbone 16.00 µL (8 µg)
Total 50 µL
Incubate at 37 °C for 2 h.
Inactivate the enzyme at 65 °C for 20 min.
Purify the digested plasmid by gel purification using the gel extraction kit.
Prepare 1% agarose gel (see Recipes).
Mix 1 µL of ladder with 1 μL of 6× loading dye and 4 µL of Milli-Q water. Load the entire volume (50 µL) on gel. Mix 2 µL of uncut pX330 plasmid sample with 0.4 µL of 6× loading dye. Load the entire volume on gel.
Mix 2.5 µL of BsbI cut pX330 plasmid sample with 0.5 µL of 6× loading dye. Load the entire volume on gel.
Run gel at 90 V for 45 min.
sgRNA annealing and ligation
Prepare stock solutions of the gRNA primers.
The gRNA primer contains 120 µg of DNA and was diluted in 200 µL of Milli-Q water to achieve a concentration of 0.6 µg/µL. Dilute gRNA primer 1:100 to achieve a concentration of 6 ng/µL.
The reverse gRNA primer contains 170 µg of DNA and was similarly diluted in 200 µL of Milli-Q water to achieve a concentration of 0.85 µg/µL. Dilute reverse gRNA primer 1:100 to achieve a concentration of 8.5 ng/µL.
Prepare the annealing reaction of each pair of oligos in annealing buffer as indicated in Table 4 and Table 5.
Table 4. Annealing buffer
Reagent Volume
Milli-Q water 45 μL
1 M HEPES 2.5 μL
1 M MgCl2 0.5 μL
2.5 M NaCl 2 μL
Total 50 μL
Table 5. Annealing reaction
Reagent Volume
Annealing buffer 48 μL
Sense oligo 1 μL (6 mg/mL)
Antisense oligo 1 μL (6 mg/mL)
Total 50 μL
Run the reaction with the following conditions in a PCR machine:
90 °C for 4 min
70 °C for 10 min
Cool from 70 °C to 37 °C at a rate of 2 °C/min
10 °C for 1 min
Hold at 4 °C
Prepare ligation reaction with pX330 and sgRNA as indicated in Table 6. Prepare a negative control without sgRNA as indicated in Table 7.
Table 6. 1:2 pX330:oligos ligation reaction
Reagent Volume
Milli-Q water 4.68 μL
10× buffer for T4 DNA ligase 1 μL
Linearized pX330 2.82 μL
T4 ligase 0.5 μL
Annealed sgRNA oligos 1 μL (0.5893 ng) 1:200 dilution
Total 10 μL
Table 7. Negative control
Reagent Volume
Milli-Q water 5.68 μL
10× buffer for T4 DNA ligase 1 μL
Linearized pX330 2.82 μL
T4 ligase 0.5 μL
Total 10 μL
Incubate ligation reactions at 16 °C overnight or RT for 2 h.
Heat-inactivate the ligation reactions at 65 °C for 10 min (optional).
Transformation of ligated plasmid pX330 + gRNA and vector control
Set water bath to 42 °C.
Thaw Stbl3 cells on ice for 10 min.
Add 3 μL of the ligated plasmid or the vector control to 50 μL of Stbl3 cells.
Incubate bacterial and DNA mixture on ice for 20 min.
Heat shock in a 42 °C water bath for 45 s.
Cool mixture on ice for 2 min.
Add 400 μL of S.O.C. medium to mixture.
Place in a 37 °C shaking incubator at 200 rpm for 1 h.
Spin tube at 13,000 rpm for 1 min at RT to pellet.
Remove 300 μL of media from the pellet. Resuspend pellet in the remaining volume of media.
Plate 100 μL of cells on LB + Amp plate (Ampicillin working concentration 100 μg/mL) using sterilized spreader.
Label plate and incubate in a 37 °C incubator overnight.
Screening of the clones
Examine plate for colony formation.
On a new LB + Amp plate, use a marker to draw a 4 × 5 grid on the backside of the plate. Streak single colonies into their own spot on the grid.
Incubate grid plate in a 37 °C incubator overnight.
Next day, inoculate 4–5 colonies that grew on the grid into 10 mL of LB + Amp broth in 50 mL Falcon tubes to isolate the plasmid for sequencing.
Place Falcon tubes in a 37 °C shaking incubator at 200 rpm overnight.
Next day, isolate the plasmids using Plasmid Isolation kit (Qiagen) following manufacturer’s protocol.
Send plasmid samples (Table 8) for sequencing to confirm the clone.
Table 8. Sample for sequencing
Reagent Volume
Putative pX330 candidate plasmid (20 ng/μL) 10 μL
U6 primer (10 ng/μL) 2 μL
Total 12 μL
Synthesizing Cdt1 gene and the AID+YFP tag into the repair template vector
We synthesized a DNA fragment containing 800 bp (between 500 and 1 kb) homology regions on each side of Cas9 cut site in the Cdt1 sequence; AID with YFP sequence was inserted in between these two homology arms (synthesized by Gene Universal). We then cloned this product into the vector HJURP using KpnI-HindIII restriction sites (a kind gift from Dr. Daniel Foltz, Northwestern University, Evanston, IL, USA). The PAM sequence within Cdt1 homology arms was mutated and replaced with synonymous codons without altering the protein sequence.
Transfect both sgRNA-cloned pX330 plasmid and the repair template (HJURP-AID-YFP) of Cdt1
Grow DLD1-TIR1 cells in complete DMEM media containing 1× penicillin-streptomycin and 10% FBS on a 60 mm plate. Incubate until they reach a confluency of 60%–70% on the day of transfection.
In an Eppendorf tube, add the equimolar ratio needed (1 μg) of each plasmid containing Cdt1 sgRNA/Cas9 and Cdt1 repair template.
Add 150 μL of EB buffer from the Transfection Reagent kit.
Add 8 μL of Enhancer. Vortex for 1 s to mix.
Incubate tube for 2–5 min at RT.
Add 25 μL of Effectene transfection reagent. Vortex for 10 s.
Incubate for 5–10 min at RT.
Add 1 mL of incomplete DMEM media containing 10% FBS without penicillin-streptomycin to the Eppendorf tube. Gently mix by pipetting.
Remove the complete media that is in the 60 mm plate. Wash the plate with 1 mL of incomplete media.
Add the transfection tube contents to the cells drop by drop.
Incubate at 37 °C with 5% CO2.
After 6 h, add 2 mL of incomplete media to the plate.
Incubate the plate for 24 h at 37 °C with 5% CO2.
Remove the incomplete media from the plate.
Add 4 mL of complete media to the plate.
Incubate for 24–48 h at 37 °C with 5% CO2.
When confluent, scale up cells into 100 mm plate.
Five to seven days after transfection, harvest cells for sorting.
Harvesting cells for sorting
Collect the media from the cells growing in the 100 mm plate and filter with a 0.22 μm filter into a 15 mL Falcon tube. This will be used to make the conditioned media.
In a new 15 mL Falcon tube, make conditioned media by combining 5 mL of the filtered media with 5 mL of complete media containing 20% FBS. Filter the conditioned media using a 0.22 μm filter.
Wash the 100 mm cell plate with 1× PBS.
Add 1 mL of 0.05% trypsin and incubate until cells detach.
Add 9 mL of complete media to wash plate and detach cells.
Collect cells into a 15 mL Falcon tube and centrifuge at 200× g for 5 min to pellet.
Remove supernatant and resuspend cells with 5 mL of 1% FBS-1×PBS.
Centrifuge cells at 200× g for 5 min.
Remove supernatant and resuspend cells starting with 1 mL complete media.
Perform cell count; then, add additional complete media such that there are 10 × 106 cells per milliliter.
In a 96-well plate, add 100 μL of conditioned media into each well for single-cell sorting.
Sort single cells into 96-well plate using fluorescence-activated cell sorting with a flow rate of 1,000 events/second (evt/s).
Screening of monoclonal lines by confocal microscopy, PCR, and western blotting
Perform PCR.
Once cells are confluent in the 96-well plate, plate cells into a 24-well plate, followed by a 12-well plate, and then finally to a 6-well plate to ensure enough propagation of the single clones.
Once cells are confluent in the 6-well plate, trypsinize and collect cells of each clone.
Isolate genomic DNA of each clone using the Quick-DNA Microprep kit following manufacturer’s instructions.
Measure the DNA concentration and use as a template for PCR.
Design primers to check modification at targeted endogenous gene locus produced by CRISPR-Cas9 using PCR method (Figure 1).
Figure 1. Strategy to confirm the homozygous and heterozygous knock-in events by PCR. Internal primers, as indicated, were designed for amplifying sequences with the Cdt1 gene (wildtype primers) and the AID-YFP insertion with Cdt1 gene (mutant primer 3).
Homozygosity and heterozygosity of the knock-in event in the clones will also be assessed by PCR. We ordered custom-made primers for the screening of clones from the company of choice (IDT).
Dissolve primers in Milli-Q water as indicated in Table 9 to achieve a concentration of 100 μM.
Table 9. PCR reaction mixture for confirming homozygous and heterozygous clones
Reagent Volume
PCR-Mix (2×) 10 μL (1×)
Primer 1 2.0 μL
Primer 2 2.0 μL
Primer 3 2.0 μL
Genomic DNA X μL (25–50 ng)
Milli-Q water Up to 20 μL
Total 20 μL
Run the reaction with the following conditions in a PCR machine for 25 cycles:
95 °C for 5 min (initial denaturation)
95 °C for 30 s (denaturation)
55 °C for 30 s (annealing)
72 °C for 1 min (extension time)
72 °C for 5 min (final extension)
Hold at 4 °C
Note: Genomic DNA volume will depend on concentration. Annealing temperature will depend on primer TM.
Analyze amplified DNA on a gel and check amplification profile. Control DLD-1 genomic DNA will produce a single band at 0.5 kb. For homozygous knock-in, the single band should be at 1 kb, while two bands at 0.5 kb and 1 kb would be seen for heterozygous knock-in.
For further confirmation of the knock-in events, purify the PCR product and sequence the product.
Clones that are PCR positive can be confirmed by confocal microscopy for GFP-positive cells.
Western blot analysis of PCR-positive candidate strains identifies the best candidate, which expresses the AID-YFP tagged protein.
Grow DLD-1 cells only (for control) and AID-positive homozygous clone in 60 mm plates and incubate until confluent. Trypsinize cells and collect in 15 mL tubes.
Centrifuge cells at 200× g for 3 min to pellet and discard the supernatant.
Lyse cell pellet with 100 μL of Pierce IP lysis buffer containing 1 μL of halt protease inhibitor 100×. Keep on ice.
Incubate cells on ice for 10 min.
Centrifuge cells at 18,000× g for 20 min at 4 °C.
Collect the supernatant into new Eppendorf tubes and determine concentration.
Add 6× Laemmli SDS sample buffer to each sample of equal concentration.
Heat samples at 95 °C for 5 min.
Load samples of equal volume along with pre-stained protein ladder on the SDS-PAGE gel (see Recipes).
Transfer into membrane and block with 2% BSA in 1× PBS. Probe with primary anti-GFP and anti-Cdt1 antibodies followed by the secondary HRP-conjugated anti-mouse or anti-rabbit antibodies. Develop the blot using the chemiluminescent substrate kit following manufacturer’s instructions to confirm integration of AID.
Validate the GFP-positive cell clones via confocal immunofluorescence microscopy and confirm that GFP-AID is degraded in these cells after the addition of IAA. Prepare the AID clones for carrying out functional phenotypic analysis as carried out in Rahi et al. (2023) [12].
Validation of AID system
Split the homozygous GFP-positive clones into in 60 mm plates. DLD-1 cells will be used in a similar way for negative control.
After 24–48 h of growing cells to 60%–70% confluency, treat cells with 500 μM of auxin (IAA) for the desired duration (0, 30, 60, and 120 min) to induce protein degradation.
Note: The used concentration of auxin ranges from 50 to 500 μM, as obtained from previous published protocols, but this should be optimized for particular AID knock-in cell line. The best way to use is to start with the lowest concentration of auxin to prevent potential side effects while still effective in depleting the protein of interest. Treatment duration may vary depending on the AID-tagged protein such that protein levels are depleted without causing loss of viability.
Harvest the cells at various time points (0, 30, 60, and 120 min) after auxin treatment.
Perform western blot using anti-tubulin antibody as a loading control or other appropriate protein analysis techniques to assess the protein degradation.
Frozen stock of clones can be prepared in freezing media (90% FBS and 10% DMSO) and stored in liquid nitrogen.
Data analysis
Analyze the protein degradation kinetics and determine the efficiency of the AID system in controlling protein expression. The AID assay provides a powerful tool for studying protein degradation and its regulatory mechanisms in mammalian cells. With proper optimization and controls, this technique can yield valuable insights into protein turnover and function.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article: Rahi et al. (2023). The Ndc80-Cdt1-Ska1 complex is a central processive kinetochore–microtubule coupling unit. J Cell Biol. (Figure 1, panel B, Figure S1, panel B) [12].
General notes and troubleshooting
General notes
Make sure the PAM sequence is mutated in the donor plasmid/repair template.
Make sure that the TIR1+ mammalian cells (DLD1/HCT-116/HeLa/293T) being used are appropriate for the phenotypic or functional characterization experiments to be adopted.
Troubleshooting
Problem 1: Selection between heterozygous and homozygous clones.
Solution: For homozygous selection (biallelic), use two donor plasmids containing different selections (e.g., one plasmid with GFP and the other with m-Cherry). Cells can be sorted after insertion to confirm they contain two colors.
Problem 2: Difficulty inducing protein degradation by auxins.
Solution: Optimize concentration of auxins, starting with the lowest concentration and then increasing concentrations.
Problem 3: Auto/leaky degradation without the addition of auxins.
Solution: Use mini-AID (7 kD) or mutant form of AID (AID2) that is still able to properly bind to auxins (IAA/5-Ph-IAA).
Acknowledgments
This work was supported by National Institute of General Medical Sciences (NIGMS) grant R01GM135391 to D. Varma and was derived from the original work performed by the Kanemaki laboratory, which has been duly cited. We would like to express out deep sense of appreciation to Shashi Kumar Suman (IGBMC, University of Strasbourg, Strasbourg, France) for his assistance with the preparation of this work.
Competing interests
The authors declare no competing interests exist.
References
Ma, H. T. (2021). The Conditional Knockout Analogous System: CRISPR-Mediated Knockout Together with Inducible Degron and Transcription-Controlled Expression. Methods Mol. Biol. 2329: 323–335.
Natsume, T., Kiyomitsu, T., Saga, Y. and Kanemaki, M. T. (2016). Rapid Protein Depletion in Human Cells by Auxin-Inducible Degron Tagging with Short Homology Donors. Cell Rep. 15(1): 210–218.
Ashley, G. E., Duong, T., Levenson, M. T., Martinez, M. A. Q., Johnson, L. C., Hibshman, J. D., Saeger, H. N., Palmisano, N. J., Doonan, R., Martinez-Mendez, R., et al. (2021). An expanded auxin-inducible degron toolkit for Caenorhabditis elegans. Genetics 217(3): e1093/genetics/iyab006.
Khakhar, A., Bolten, N. J., Nemhauser, J. and Klavins, E. (2015). Cell–Cell Communication in Yeast Using Auxin Biosynthesis and Auxin Responsive CRISPR Transcription Factors. ACS Synth. Biol. 5(4): 279–286.
Shetty, A., Reim, N. I. and Winston, F. (2019). Auxin‐Inducible Degron System for Depletion of Proteins in Saccharomyces cerevisiae. Cu Curr Protoc Mol Biol 128(1): e104.
Brosh, R., Hrynyk, I., Shen, J., Waghray, A., Zheng, N. and Lemischka, I. R. (2016). A dual molecular analogue tuner for dissecting protein function in mammalian cells. Nat. Commun. 7(1): e1038/ncomms11742.
Saito, Y. and Kanemaki, M. T. (2021). Targeted Protein Depletion Using the Auxin‐Inducible Degron 2 (AID2) System. Curr. Protocol. 1(8): e219.
Nishimura, K., Yamada, R., Hagihara, S., Iwasaki, R., Uchida, N., Kamura, T., Takahashi, K., Torii, K. U. and Fukagawa, T. (2020). A super-sensitive auxin-inducible degron system with an engineered auxin-TIR1 pair. Nucleic Acids Res. 48(18): e108–e108.
Yesbolatova, A., Saito, Y. and Kanemaki, M. T. (2020). Constructing Auxin-Inducible Degron Mutants Using an All-in-One Vector. Pharmaceuticals (Basel) 13(5): 103.
Macdonald, L., Taylor, G., Brisbane, J., Christodoulou, E., Scott, L., Von Kriegsheim, A., Rossant, J., Gu, B. and Wood, A. (2022). Rapid and specific degradation of endogenous proteins in mouse models using auxin-inducible degrons. eLife: e476100.
Natsume, T. and Kanemaki, M. T. (2017). Conditional Degrons for Controlling Protein Expression at the Protein Level. Annu. Rev. Genet. 51(1): 83–102.
Rahi, A., Chakraborty, M., Agarwal, S., Vosberg, K. M., Agarwal, S., Wang, A. Y., McKenney, R. J. and Varma, D. (2023). The Ndc80-Cdt1-Ska1 complex is a central processive kinetochore–microtubule coupling unit. J. Cell Biol. 222(8): e202208018.
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YY Yoshio Yamauchi
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4924 Views: 873
Reviewed by: Faraz RashidSuprabhat Mukherjee Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Jan 2023
Abstract
Cholesterol is oxygenated by a variety of cholesterol hydroxylases; oxysterols play diverse important roles in physiological and pathophysiological conditions by regulating several transcription factors and cell-surface receptors. Each oxysterol has distinct and overlapping functions. The expression of cholesterol hydroxylases is highly regulated, but their physiological and pathophysiological roles are not fully understood. Although the activity of cholesterol hydroxylases has been characterized biochemically using radiolabeled cholesterol as the substrate, their specificities remain to be comprehensively determined quantitatively. To better understand their roles, a highly sensitive method to measure the amount of various oxysterols synthesized by cholesterol hydroxylases in living mammalian cells is required. Our method described here, with gas chromatography coupled with tandem mass spectrometry (GC–MS/MS), can quantitatively determine a series of oxysterols endogenously synthesized by forced expression of one of the four major cholesterol hydroxylases—CH25H, CYP7A1, CYP27A1, and CYP46A1—or induction of CH25H expression by a physiological stimulus. This protocol can also simultaneously measure the amount of intermediate sterols, which serve as markers for cellular cholesterol synthesis activity.
Key features
• Allows measuring the amount of a variety of oxysterols synthesized endogenously by cholesterol hydroxylases using GC–MS/MS.
• Comprehensive and quantitative analysis of cholesterol hydroxylase specificities in living mammalian cells.
• Simultaneous quantification of intermediate sterols to assess cholesterol synthesis activity.
Graphical overview
Keywords: Cholesterol Oxysterols Intermediate sterols Cholesterol hydroxylase GC–MS/MS CH25H CYP27A1 CYP46A1 CYP7A1
Background
Cholesterol plays diverse important biological roles, including function regulation of biological membranes and membrane proteins, and is the precursor for steroid hormones and bile acids; thus, cellular cholesterol homeostasis is tightly controlled by multiple mechanisms (Chang et al., 2006; Yamauchi and Rogers, 2018; Luo et al., 2020). Mammalian cells cannot break down the sterol backbone. Instead, cholesterol is converted to cholesteryl ester for storage (Chang et al., 2006) and to various oxysterols (Mutemberezi et al., 2016). Oxysterols exert different functions depending on the hydroxylation site(s). Multiple oxysterols whose side chain is hydroxylated, including 25-hydroxycholesterol (25-HC) and 27-HC, regulate cellular cholesterol homeostasis; they modulate two important transcription factors: sterol regulatory element-binding protein-2 (SREBP-2) and liver X receptor (LXR) (Gill et al., 2008; M.S. Brown et al., 2018). SREBP-2 transactivates most genes involved in cholesterol acquisition (biosynthesis and uptake) (Horton et al., 2002). On the other hand, LXR upregulates the expression of several ATP-binding cassette (ABC) transporters including ABCA1 and ABCG1 that mediate the export of excess cellular cholesterol (Tontonoz, 2011). Mechanistically, side-chain oxysterols bind to Insig-1 and Insig-2—retention factors for SREBP-2 in the endoplasmic reticulum (ER)—and protect them from proteasomal degradation, thereby inhibiting SREBP-2 activation. Side-chain oxysterols also serve as natural LXR ligands when added exogenously.
A series of hydroxylases catalyze the site-specific hydroxylation of cholesterol (Mutemberezi et al., 2016). Although a variety of oxysterols have been identified in the body, 7α-HC, 27-HC, 24S-HC, and 25-HC are the most abundant. These four oxysterols are synthesized largely by CYP7A1, CYP27A1, CYP46A1, and CH25H, respectively, in the ER or mitochondria (Figure 1), while some of these hydroxylases also produce other oxysterols as minor products (Mutemberezi et al., 2016; Saito et al., 2023). Recent studies show that the expression of cholesterol hydroxylases is highly regulated in physiological and pathophysiological conditions (Cyster et al., 2014; A.J. Brown et al., 2021). CH25H expression and 25-HC biosynthesis are markedly upregulated upon infection in immune cells such as macrophages, and 25-HC itself exhibits anti-bacterial and anti-viral effects, protecting cells from infection (Bauman et al., 2009; S.Y. Liu et al., 2013; Zhou et al., 2020). 25-HC can be further hydroxylated at the 7-position by the hydroxylase CYP7B1, generating 7α,25-dihydroxycholesterol (7α,25-diHC). This dihydroxycholesterol is a ligand for the G-protein coupled receptor GPR183 (also known as EBI2) involved in immune cell migration (C. Liu et al., 2011). Higher circulating 27-HC levels associate with the risk of estrogen receptor-positive breast cancer, since 27-HC serves as an endogenous selective estrogen receptor modulator (Nelson et al., 2013). CYP46A1 is a neuron-specific cholesterol hydroxylase that converts cholesterol into 24S-HC for eliminating excess cholesterol in the brain (Russell et al., 2009). CYP7A1 is a liver-specific hydroxylase that serves as the rate-limiting enzyme for bile acid synthesis (Russell, 2009). Although the activity of these cholesterol hydroxylases is biochemically studied using radiolabeled cholesterol as the substrate, comprehensive and quantitative characterization of their specificities remains poorly explored in living cells. Furthermore, precise measurement of the products of these cholesterol hydroxylases is crucial for better understanding their physiological and pathophysiological roles.
Here, we provide a highly sensitive method using gas chromatography coupled with tandem mass spectrometry (GC–MS/MS) to determine various oxysterols synthesized in living mammalian cells where a cholesterol hydroxylase is forcedly expressed or is upregulated by lipopolysaccharide, a component of Gram-negative bacteria known to upregulate CH25H expression in macrophages (Bauman et al., 2009). The current GC–MS/MS protocol enabled us to reveal cholesterol hydroxylase–specific production of oxysterols (Saito et al., 2023). Our protocol can also simultaneously determine the amount of intermediate sterols to monitor the activity of cholesterol biosynthesis. Accordingly, our GC–MS/MS-based sterol analysis, in combination with biochemical and gene expression studies, has suggested that side-chain oxysterols enzymatically synthesized within cells primarily regulate SREBP-2 but not LXR (Saito et al., 2023).
Figure 1. Hydroxylation of cholesterol by the four cholesterol hydroxylases handled in this protocol. CH25H, CYP27A1, CYP46A1, and CYP7A1 mainly produce 25-HC, 27-HC, 24(S)-HC, and 7α-HC, respectively.
Materials and reagents
1.5 mL sampling tubes (WATSON, catalog number: 131-8155C)
Screw-top test tube (Maruemu, catalog number: NR-10)
Mighty Vials (Maruemu, catalog number: 84-0561)
TORAST vial (Shimadzu GLC, catalog number: GLCTV-902)
TORAST vial insert (Shimadzu GLC, catalog number: GLCTV-I04)
TORAST vial cap (Shimadzu GLC, catalog number: GLCTV-903)
Hexane (Fuji Film Wako, catalog number: 085-00416)
2-propanol (Fuji Film Wako, catalog number: 166-04836)
Dibutyl hydroxytoluene (BHT) (Fuji Film Wako, catalog number: 029-07392)
Nitrogen gas
Helium gas
Argon gas
Chloroform (Fuji Film Wako, catalog number: 038-02606)
Ethanol (Fuji Film Wako, catalog number: 057-00456)
Milli-Q water
Pyridine (Fuji Film Wako, catalog number: 161-18453)
Note: Store pyridine in a desiccator with silica gel at room temperature and use within four weeks after opening.
N-methyl-N-trimethylsilyl trifluoroacetamide (MSTFA) (GL Science, catalog number: 1022-11061)
Cholesterol (purity ≥ 99%) (Sigma-Aldrich, catalog number: C8667)
Cholesterol-d7 (purity ≥ 99%) (Avanti Polar Lipids, catalog number: 700041P)
Desmosterol (purity ≥ 99%) (Nagara Science, catalog number: NS460402)
Lanosterol (purity ≥ 99.5%) (Nagara Science, catalog number: NS460102)
Lathosterol (purity ≥ 99%) (Nagara Science, catalog number: NS460502)
7-dehydrocholesterol (purity ≥ 95%) (Sigma-Aldrich, catalog number: 30800)
24,25-epoxycholesterol (purity ≥ 95%) (Abcam, catalog number: Ab141633)
24,25-dihydrolanosterol (purity ≥ 99.5%) (Nagara Science, catalog number: NS460201)
4β-hydroxycholesterol (purity ≥ 95%) (Cayman, catalog number: 19518)
7α-hydroxycholesterol (purity ≥ 99%) (Avanti Polar Lipids, catalog number: 700034P)
7β-hydroxycholesterol (purity ≥ 95%) (Sigma-Aldrich, catalog number: H6891)
7α,25-dihydroxycholesterol (purity ≥ 98%) (Sigma-Aldrich, catalog number: SML0541)
7α,27-dihydroxycholesterol (purity ≥ 99%) (Avanti Polar Lipids, catalog number: 700024P)
25-hydroxycholesterol (purity ≥ 98%) (Sigma-Aldrich, catalog number: H1015)
24(S)-hydroxycholesterol (purity ≥ 98%) (Sigma-Aldrich, catalog number: SML1648)
25-hydroxycholesterol-d6 (Avanti Polar Lipids, catalog number: LM4113-1EA)
27-hydroxycholesterol (purity ≥ 98%) (Sigma-Aldrich, catalog number: SML2042)
Sodium chloride (Fuji Film Wako, catalog number: 196-01665)
Potassium chloride (SIGMA, catalog number: P9541)
Disodium hydrogen phosphate dodecahydrate (Fuji Film Wako, catalog number: 196-02835)
Potassium dihydrogen phosphate (Fuji Film Wako, catalog number: 196-04245)
Potassium hydroxide (Fuji Film Wako, catalog number: 168-21815)
Sodium hydroxide (Fuji Film Wako, catalog number: 192-15985)
D-MEM/Ham’s F-12 with L-Glutamine and Phenol Red (Fuji Film Wako, catalog number: 048-29785)
RPMI-1640 with L-Glutamine and Phenol Red (Fuji Film Wako, catalog number: 189-02025)
Opti-MEMTM I reduced serum medium (Thermo Fisher Scientific, catalog number: 31985070)
Fetal bovine serum (FBS) (lot number: 27419002) (Corning, catalog number: 35-079-CF)
Penicillin G potassium (Meiji Seika Pharma, catalog number: 6111400D3051)
Streptomycin sulfate (Meiji Seika Pharma, catalog number: 6161400D1034)
Lipofectamine LTX reagent (Thermo Fisher Scientific, catalog number: 15338100)
Doxycycline hyclate (LKT Labs, catalog number: D5897)
Kdo2-Lipid A (Avanti Polar Lipids, catalog number: 69500P)
Pierce BCA Protein Assay kit (Thermo Fisher Scientific, catalog number: 23227)
Biological materials
CHO-K1 cells (gift from Dr. Ta-Yuan Chang, Geisel School of Medicine at Dartmouth)
CHO-CH25Htet-on cells (Saito et al., 2023)
J774.1 cells (RIKEN Cell Bank, catalog number: RCB0434)
Expression plasmids encoding cholesterol hydroxylase (Saito et al., 2023): pFLAG-CH25H, pCYP27A1-FLAG, pCYP46A1-FLAG, and pCYP7A1-FLAG
Note: Detailed information on CHO-CH25Htet-on cells and the four hydroxylase expression plasmids are described in the original paper (Saito et al., 2023).
Solutions
70% Ethanol (see Recipes)
75% Ethanol (see Recipes)
10 N KOH (dissolved in 75% ethanol) (see Recipes)
Hexane/2-propanol (3:2) with 0.01% BHT (see Recipes)
0.1 N NaOH (see Recipes)
PBS (see Recipes)
Recipes
70% Ethanol
Reagent Volume
Ethanol (absolute) 35 mL
H2O 15 mL
Total 50 mL
75% Ethanol
Reagent Volume
Ethanol (absolute) 37.5 mL
H2O 12.5 mL
Total 50 mL
10 N KOH (dissolved in 75% ethanol)
Reagent Quantity
Potassium hydroxide 28.1 g
75% Ethanol up to 50 mL
Total 50 mL
Hexane/2-propanol (3:2) with 0.01% BHT
Reagent Quantity
Hexane 60 mL
2-propanol 40 mL
Dibutyl hydroxytoluene 10 mg
Total 100 mL
0.1 N NaOH
Reagent Quantity
Sodium hydroxide 0.2 g
H2O up to 50 mL
Total 50 mL
PBS
Reagent Quantity
Sodium chloride 8 g
Potassium chloride 0.2 g
Disodium hydrogen phosphate dodecahydrate 3.6 g
Potassium dihydrogen phosphate g
H2O up to 1 L
Sodium hydroxide 1 L
Equipment
GC–MS/MS (Shimadzu, model: GCMS-TQ8040NX) equipped with an auto-sampler (AOC-20i)
BPX5 GC capillary column (30 m × 0.25 mm, 0.25 μm) (TRAJAN, catalog number: SGE-054101)
Rxi Guard column (5 m × 0.25 mm) (Restek, catalog number: 10029)
Eppendorf ThermoMixer C (Eppendorf, catalog number: 5382000023)
Hamilton micro syringe 710RN (Hamilton, catalog number: 72-5004)
Pressured gas blowing concentrator (EYELA, model: MGS-2200)
Aluminum block MGB-1540 (EYELA, catalog number: 207580)
Aluminum block MGB-1624 (EYELA, catalog number: 207610)
Low-speed refrigerated centrifuge (TOMY, model: AX-511)
SpectraMax (Molecular Devices, model: M2e)
Software and datasets
GC-MSsolution v4 (Shimadzu)
RStudio software (Posit)
GraphPad Prism 9 (GraphPad)
Procedure
Cell culture
Preparation of CHO-K1 cells expressing cholesterol hydroxylase
Seed CHO-K1 cells in triplicate into 6-well plates at a density of 2 × 105 cells per well and incubate in DMEM/Ham’s F-12 medium supplemented with 7.5% FBS, penicillin (100 unit/ mL), and streptomycin (0.1 mg/mL) at 37 °C and 5% CO2 for 18–24 h.
Change the medium to Opti-MEM 2–3 h before transfection.
Transfect cells with 2 μg of plasmid encoding either CH25H, CYP7A1, CYP27A1, or CYP46A1 (pFLAG-CH25H, pCYP7A1-FLAG, pCYP27A1-FLAG, or pCYP46A1-FLAG, respectively) using Lipofectamine LTX reagent according to the manufacturer’s protocol.
Note: These four hydroxylase expression constructs are described in the original paper (Saito et al., 2023).
Change the medium to DMEM/Ham’s F-12 medium containing 0.1% FBS, penicillin (100 unit/ mL), and streptomycin (0.1 mg/mL) 5 h after transfection.
Incubate cells for 24 h at 37 and 5% CO2 before harvesting cells for lipid extraction.
Preparation of CHO-CH25Htet-on cells
Plate 2 × 105 CHO-CH25Htet-on cells in triplicate into a well of a 6-well plate and incubate for 18–24 h in DMEM/Ham’s F-12 medium with 7.5% FBS, penicillin (100 unit/ mL), and streptomycin (0.1 mg/mL) at 37 °C and 5% CO2.
Treat the cells for 24 h with or without 0.4 or 1 μg/mL of doxycycline hyclate in DMEM/Ham’s F-12 medium containing 0.1% FBS and penicillin/streptomycin to induce FLAG-CH25H expression and harvest the cells to extract cell lipids.
Preparation of J774.1 cells
Seed J774.1 cells in triplicate into 6-well plates at a density of 5 × 105 cells per well and incubate for two days in RPMI-1640 medium supplemented with 10% FBS and penicillin/streptomycin at 37 and 5% CO2.
Incubate the cells with or without 100 ng/mL of Kdo2-Lipid A, a toll-like receptor 4 ligand, in RPMI-1640 medium containing 10% FBS and penicillin/streptomycin for 20 h at 37 before extracting cell lipids.
Extraction of cellular lipids
Wash cells twice in 6-well plates with PBS (1.5 mL/well) and let cells dry at room temperature.
Add 1.5 mL/well of hexane/2-propanol (3:2) with 0.01% BHT (see Recipes) into 6-well plates and place at room temperature for 1 h to extract cellular lipids.
Collect hexane/2-propanol into a glass tube (screw-top test tube).
Add 1 mL/well of hexane/2-propanol (3:2) with 0.01% BHT into the same well and place at room temperature for 30 min.
Transfer hexane/2-propanol into the same glass tube to pool the cellular lipids extracted.
Add cholesterol-d7 (100 ng/tube from 5 μg/mL stock in chloroform) and 25-hydroxycholesterol-d6 (10 ng/tube from 1 μg/mL stock in methanol) to each sample as internal controls.
Evaporate the organic solvent under the nitrogen gas stream at room temperature using a pressured gas blowing concentrator MGS-2200.
Add 1 mL of 100% ethanol and 300 μL of 10 N KOH (dissolved in 75% ethanol) (see Recipes) to each tube, close with a screw cup, and mix gently.
Incubate the tubes at 80 °C for 1 h to saponify lipids.
Cool the tubes on ice.
Add 2 mL of chloroform into each tube and vortex.
Add 2.5 mL of distilled water into each tube and vortex.
Centrifuge the tubes at 2,380× g for 10 min at room temperature using the low-speed refrigerated centrifuge AX-511.
Remove the aqueous phase (upper layer) using an aspirator.
Add 2 mL of distilled water into each tube again and vortex for 1 min.
Centrifuge the tubes at 2,380× g for 10 min at room temperature.
Remove the aqueous layer again using an aspirator.
Add 2 mL of distilled water into each tube again, vortex, and centrifuge at 2,380× g for 10 min at room temperature.
Remove the aqueous layer using an aspirator.
Transfer the chloroform phase (which contains non-saponified lipids, including sterols) to a new glass vial (Mighty Vials) using a Pasteur pipette and dry under nitrogen gas stream.
Derivatization of extracted cellular sterols
Add 50 μL of pyridine and 50 μL of MSTFA into each vial and mix gently.
Incubate the vials at 80 °C for 1 h with agitation at 300 rpm using an Eppendorf ThermoMixer C.
Transfer the derivatized sample to a new TORAST vial with a TORAST vial insert using a Hamilton micro syringe and close the tube with a TORAST vial cap.
Preparation of standard sterol mixtures
Prepare two types of standard sterol mixtures in separate vials, as follows: the standard mixtures of non-hydroxysterols contain cholesterol, cholesterol-d7, desmosterol, 7-dehydrocholesterol, lathosterol, lanosterol, and 24,25-dihydrolanosterol; those of oxysterols contain 4β-hydroxycholesterol, 7α-hydroxycholesterol, 7β-hydroxycholesterol, 24(S)-hydroxycholesterol, 25-hydroxycholesterol, 25-hydroxycholesterol-d6, 27-hydroxycholesterol, 7α,25-dihydroxycholesterol, 7α,27-dihydroxycholesterol, and 24,25-epoxycholesterol. The quantities of each sterol added to each vial are 0.1, 1, 5, 10, 50, and 100 ng for the non-hydroxysterol mixtures and 0.01, 0.1, 0.5, 1, 5, 10, and 20 ng for the oxysterol mixtures.
Dry up under nitrogen gas stream at room temperature.
Dissolve sterols in 50 μL of pyridine and 50 μL of MSTFA.
Note: The final concentrations are 1, 10, 50, 100, 500, and 1000 ng/mL for the non-hydroxysterol mixtures and 0.1, 1, 5, 10, 50, 100, and 200 ng/mL for the oxysterol mixtures.
Incubate the vials at 80 °C for 1 h with agitation at 300 rpm using an Eppendorf ThermoMixer C.
Transfer the derivatized sample to a new TORAST vial with a TORAST vial insert using a Hamilton micro syringe and close the tube with a TORAST vial cap.
GC–MS/MS analysis
Inject 1 μL of sample or the standard mixture into a GCMS-TQ8040 NX equipped with a BPX5 GC column using an AOC-20i autosampler in splitless mode. Detailed conditions for the analysis are described in Table 1.
Table 1. GC–MS/MS conditions
GC parameters
Injection mode Spitless
Column BPX5 GC column (30 m × 0.25 mm, 0.25 mm) Rxi Guard Column (5 m × 0.25 mm)
Carrier gas control Linear velocity (49.5 cm/s)
High pressure injection 250 kPa (1 min)
Injection temperature 275 °C
Column oven temperature 200 °C (1 min) → 25 °C/min → 250 °C → 15 °C/min → 290 °C → 5 °C/min → 320 °C (2 min)
MS parameters
Interface temperature 280 °C
Ion source temperature 230 °C
Loop time 0.3 s
Measurement mode Multiple reaction monitoring (MRM)
Detect the 17 sterols by MS/MS under multiple reaction monitoring (MRM) mode. Retention time, collision energies, quantification ions, and confirmation ions for each sterol are shown in Table 2. Figure 2 shows MRM chromatograms for respective sterols.
Table 2. Retention time, quantification ions, confirmation ions, and collision energy (CE) for the sterol analysis
Sterols Retention time (min) Quantification ions Confirmation ions
MRM transition (m/z) CE (eV) MRM transition (m/z) CE (eV)
7α-hydroxycholesterol 8.48 456 > 208.3 18 456 > 119, 456 > 95.2 30, 33
Cholesterol-d7 8.89 336 > 121.1 18 375 > 145.1, 336 > 109.2 18, 18
Cholesterol 8.94 368 > 145.1 21 329 > 95.1, 329 > 81.1 27, 24
Desmosterol 9.24 129 > 73 15 129 > 57.9, 129 > 127.1 30, 15
7-dehydrocholesterol 9.28 351 > 145.1 30 351 > 128.1 42
7β-hydroxycholesterol 9.38 456 > 233.2 18 233 > 73.1 24
Lathosterol 9.41 213 > 157.2 12 213 > 81.1 15
4β-hydroxycholesterol 9.61 366 > 158.3 18 366 > 129.1 36
24,25-dihydrolanosterol 10.01 395 > 145.2 30 395 > 107 30
7α,25-dihydroxycholesterol 10.21 131 > 73.1 18 544 > 73.2 39
Lanosterol 10.36 393 > 95.2 24 393 > 187.4 12
24,25-epoxycholesterol 10.44 143 > 73.1 18 129 > 73.1, 143 > 128.1 15, 18
24(S)-hydroxycholesterol 10.60 159 > 69.1 9 159 > 73.1 18
7α,27-dihydroxycholesterol 10.60 103 > 73.1 9 544 > 233.2 33
25-hydroxycholesterol-d6 10.75 137 > 73.2 15 137 > 58.2 30
25-hydroxycholesterol 10.79 131 > 73.2 9 131 > 58.1 30
27-hydroxycholesterol 11.26 129 > 73.1 15 456 > 131.2, 417 > 69 42, 42
Notes:
Cholesterol-d7 and 25-hydroxycholesterol-d6 serve as internal standards for non-hydroxylated sterols and hydroxylated sterols, respectively.
The quantification ions are used to identify and calculate the amount of each sterol. The confirmation ions are for assisting in the identification of respective sterols. Several sterols share the same precursor ions, quantification ions, and confirmation ions. When these sterols are not resolved, distinct precursor ions should be selected for the identification and quantification.
Precursor ions for quantification and confirmation ions are carefully determined by the following criterion: higher m/z and stronger intensity. Collision energy is automatically calculated by the instrument.
Retention times for each sterol become shorter after a column is cut for maintenance because it depends on the column length.
Figure 2. Multiple reaction monitoring (MRM) chromatograms of sterols analyzed in this protocol. MRM chromatograms for each sterol are shown according to retention time. Quantification ions (black) and confirmation ions (pink and blue) are presented in each chromatogram. Black lines and pink and blue lines denote quantification ions and confirmation ions, respectively.
Cellular protein assay
After cellular lipid extraction (step B5), add 1 mL of 0.1 N NaOH (see Recipes) into each well and place at room temperature for 20–24 h to solubilize cellular protein.
Mix solubilized proteins using a pipette and transfer 10 μL of a sample to a well of a 96-well plate.
Add 150 μL of BCA assay reagent to each well, incubate the plate for 30 min at 37 °C, and record absorbance at 562 nm using a plate reader (SpectraMax M2e).
Determine the protein concentration of samples with bovine serum albumin as a standard.
Data analysis
Obtain each sterol peak area using GC-MSsolution v4 software.
Create a standard curve for each sterol using GC-MSsolution. Typical standard curves for individual sterols are shown in Supplementary Figure 1.
Determine the concentration of each sterol in a sample using GC-MSsolution based on peak areas and the standard curve.
Note: As described in Procedure D, the ranges of the two standard curves are as follows: 1 ng/mL to 1 μg/mL for non-hydroxysterols and 0.1 ng/mL to 200 ng/mL for oxysterols.
Find the extraction efficiency of cholesterol-d7 and 25-hydroxycholesterol-d6 with the following equation:
where E is extraction efficiency, C is the concentration (μg/mL) of cholesterol-d7 or 25-hydroxycholesterol-d6 detected by GC–MS/MS, V is the final volume (mL) of the sample (0.1 mL in this protocol), and I is the quantity (ng) of the internal standard added at the step B6 (100 ng of cholesterol-d7 or 10 ng of 25-hydroxycholeterol-d6, in this protocol).
Calculate the original amounts of each sterol per sample using the appropriate extraction efficiency.
Note: The extraction efficiencies of cholesterol-d7 and 25-hydroxycholesterol-d6 are used to calculate the original amounts of non-hydroxysterols and oxysterols, respectively.
Normalize the amounts of each sterol to cell protein per well (Figure 3).
All data are represented by the mean ± S.D. of at least three independent biological replicates. Statistical analyses are performed with RStudio software by Student’s t-test or one-way ANOVA with Dunnett or Tukey-Kramer post-hoc test. p < 0.05 is considered statistically significant.
Advantages and limitations
LC–MS/MS, GC–MS, and GC–MS/MS have been employed for sterol analysis, with GC–MS as the most traditional technique (Krone et al., 2010; McDonald et al., 2012; Griffiths et al., 2013; Saito et al., 2023). Although each method has advantages and limitations, the current GC–MS/MS protocol described above has several advantages over other methods. Since our method is highly sensitive, as 1 pg of sterols can be detected, lipids extracted from cells in a well of a 6-well plate (approximately 0.5–1 × 106 cells) are sufficient to measure a series of sterols, including oxysterols and intermediate sterols. The current GC–MS/MS analysis of cellular sterol is much more sensitive than our previous analysis with GC–MS, in which cellular lipids were extracted from a 100 mm dish or a whole 6-well plate (Yamauchi et al., 2007), while the amounts of cellular sterol contents detected are equivalent. Therefore, many samples can simultaneously be handled for studying multiple experimental conditions in a single assay. In addition, the run time for a sample in our method is approximately 15 min. Typical run times for sterol analysis by GC–MS and LC–MS/MS are 20–30 min and 12–20 min, respectively, depending on the columns and conditions used. Thus, our GC–MS/MS method is comparable to LC–MS/MS analysis concerning run time.
Furthermore, GC-based methods generally show better chromatographic resolution than LC–MS/MS. The previous LC–MS/MS method was unable to resolve 7α-HC and 7β-HC (McDonald et al., 2012), whereas our GC–MS/MS protocol distinguished between these two oxysterols (Saito et al., 2023). As such, GC–MS/MS is the superior technology for analyzing small amounts of isomeric oxysterols, while a major limitation of LC–MS/MS methods is that such oxysterols tend to provide similar spectra.
However, GC–MS/MS-based sterol analysis also has disadvantages. In contrast to LC–MS/MS methods, which do not often require derivatization (McDonald et al., 2012), sterol needs to be derivatized for GC–MS/MS analysis, which is a laborious and time-consuming process. In addition, since GC–MS/MS detects a compound of interest with MRM mode like LC–MS/MS, only target molecules with available standards can be measured.
In summary, the current GC–MS/MS protocol provides a rapid and highly sensitive method for quantification of a series of oxysterols synthesized endogenously within cells, determining cholesterol hydroxylase activity quantitatively.
Figure 3. Comprehensive and quantitative determination by GC–MS/MS of oxysterols synthesized by cholesterol hydroxylases in living cells. (A) Oxysterol contents in CHO-K1 cells forcedly expressing CH25H, CYP27A1, CYP46A1, or CYP7A1. The data show that CH25H and CYP7A1 are very specific to synthesizing 25-HC and 7α-HC, respectively, while CYP27A1 and CYP46A1 exhibit broader specificities; CYP27A1 produces not only 27-HC but also 25-HC, and CYP46A1 synthesizes 25-HC and 27-HC in addition to 24(S)-HC. (B) CH25H expression level–dependent production of 25-HC and its effect on intermediate sterol contents in CHO-CH25Htet-on cells. Doxycycline (Dox) induces 25-HC synthesis in a dose-dependent manner, which results in the reduction in intermediate sterol contents. 24,25-DHL, 24,25-dihydrolanosterol; 7-DHC, 7-dehydrocholesterol. (C) 25-HC and 7α,25-diHC contents in J774.1 murine macrophages treated with or without 100 ng/mL of Kdo2-Lipid A (KLA) for 20 h. Stimulation of the cells with KLA induces the expression of CH25H and the production of 25-HC and 7α,25-diHC. Statistical analyses were performed by one-way ANOVA with Dunnett post-hoc test (B) or Student’s t-test (C) (*p < 0.05, **p < 0.01, ***p < 0.001). Data presented in this figure were reproduced from the original paper (Saito et al., 2023).
Acknowledgments
This is a detailed protocol of the GC–MS/MS analysis reported in Saito et al. (2023). We thank Ms. Misato Takata for her assistance with the maintenance of our GC–MS/MS machine. The study was supported by the Japan Agency for Medical Research and Development (AMED)-CREST grants 20gm091008h and 21gm091008h (to Y. Y. and R. S.) from the Japan Agency for Medical Research and Development, KAKENHI grants 19H02908 and 22H02281 (to Y. Y.) and 20H00408 (to R. S.) from the Japan Society for the Promotion of Science, and a research grant from Asahi Group Foundation (to Y. Y.). H.S. was supported by the Japan Society for the Promotion Science Research Fellowship for Young Scientist (20J10181).
Competing interests
The authors declare no conflicts of interest.
Ethical considerations
No human or animal subjects were included in this study.
References
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Supplementary information
The following supporting information can be downloaded here:
Supplementary Figure 1.
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Mesenteric Parametrial Fat Pad Surgery for in vivo Implantation of Hepatocytes in Nude Mice
SS Saloni Sinha *
DN Duc-Huy T. Nguyen *
NH Nora Hassan
QA Qazi Ali *
JS Jason Sethiadi *
ST Sergey Tsoy *
RS Robert E. Schwartz
(*contributed equally to this work)
Published: Vol 14, Iss 2, Jan 20, 2024
DOI: 10.21769/BioProtoc.4925 Views: 607
Reviewed by: Pilar Villacampa AlcubierreAras MattisWilliam C. W. Chen
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Original Research Article:
The authors used this protocol in Science Translational Medicine Jul 2017
Abstract
Cell-based liver therapies utilizing functionally stabilized engineered hepatic tissue hold promise in improving host liver functions and are emerging as a potential alternative to whole-organ transplantation. Owing to the ability to accommodate a large ex vivo engineered hepatocyte mass and dense vascularization, the mesenteric parametrial fat pad in female nude mice forms an ideal anatomic microenvironment for ectopic hepatocyte transplantation. However, the lack of any reported protocol detailing the presurgical preparation and construction of the engineered hepatic hydrogel, fat pad surgery, and postsurgical care and bioluminescence imaging to confirm in vivo hepatocyte implantation makes it challenging to reliably perform and test engraftment and integration with the host. In this report, we provide a step-by-step protocol for in vivo hepatocyte implantation, including preparation of hepatic tissue for implantation, the surgery process, and bioluminescence imaging to assess survival of functional hepatocytes. This will be a valuable protocol for researchers in the fields of tissue engineering, transplantation, and regenerative medicine.
Key features
• Primary human hepatocytes transduced ex vivo with a lentiviral vector carrying firefly luciferase are surgically implanted onto the fat pad.
• Bioluminescence helps monitor survival of transplanted hepatic tissue over time.
• Applicable for assessment of graft survival, graft-host integration, and liver regeneration.
Graphical overview
Keywords: Hepatocyte Hydrogel Nude mice Parametrial fat pad Implantation In vivo imaging
Background
The liver is the largest and an essential internal organ of the human body, as it performs a multitude of detoxification, metabolic, synthetic, and immunologic functions. Most importantly, the liver possesses a unique ability to spontaneously regenerate after being subjected to damaging insults or injuries. Owing to this regenerative potential, orthotopic liver transplantation offers a valuable treatment option for patients suffering from end-stage liver diseases [1]. However, this is the only curative treatment option and remains limited due to organ scarcity and to patients who are not at a very advanced age, do not have any significant comorbidities, and are able to undergo a surgical procedure. Additionally, financial challenges and the long-term immunosuppression required posttransplant often make liver transplantation a last resort. Given the current organ shortage and paucity of living donors, it is becoming increasingly challenging to tackle the burden of liver diseases such as nonalcoholic fatty liver disease, cirrhosis, and hepatocellular carcinoma, which account for ~2 million deaths per year worldwide [2]. Hence, cell-based therapies offer new therapeutic options to restore liver function in patients with chronic liver diseases [3].
The current protocol describes the detailed procedures to perform ectopic transplantation of human hepatic tissue onto the mesenteric parametrial fat pad in an immune-compromised mouse model. Specifically, in this protocol, primary human hepatocytes are transduced with a bioluminescent lentiviral vector and plated in 3D coculture with mouse fibroblasts to form hepatic aggregates. These aggregates are embedded inside a hydrogel to form a hepatic tissue, which is subsequently transplanted onto the mesenteric parametrial fat pad in nude mice. Bioluminescent signal from transplanted hepatic tissue allows monitoring of the tissue over time. This valuable ectopic human hepatocyte transplantation model can be used for understanding important aspects of liver tissue engineering such as graft survival, graft-host integration, and liver regeneration.
Materials and reagents
Biological materials
Plateable cryopreserved primary human hepatocytes (Lonza, catalog number: HUCPG)
Mouse 3T3-J2 fibroblasts (Kerafast, catalog number: EF3003)
Female nude mice, 8–10 weeks (Taconic Biosciences, catalog number: NCRNU)
Reagents
Reagents required for tissue culture
Fibrinogen from bovine plasma (Millipore Sigma, catalog number: F8630)
Thrombin from bovine plasma (Millipore Sigma, catalog number: T4648)
AggreWellTM 400 microwell plates (STEMCELL Technologies, catalog number: 34411)
DMEM, high glucose, pyruvate (Thermo Fisher Scientific, catalog number: 11995065)
Fetal bovine serum (FBS), premium select, heat inactivated (R&D Systems, catalog number: S11550H)
Penicillin-Streptomycin (10,000 U/mL) (Thermo Fisher Scientific, catalog number: 15140122)
CorningTM ITS+ premix universal culture supplement (Fisher Scientific, catalog number: CB-40352)
HEPES (1 M) (Thermo Fisher Scientific, catalog number: 15630080)
Dexamethasone (Millipore Sigma, catalog number: D4902)
Glucagon (Millipore Sigma, catalog number: G2044)
Polybrene, 10 mg/mL, liquid (EMD Millipore, catalog number: TR-1003-G)
CorningTM HepGoTM hepatocyte culture medium (Fisher Scientific, catalog number: MT454670)
Lentiviral vector expressing firefly luciferase under the human albumin promoter (pTRIP.Alb.IVSb.IRES.tagRFP-DEST) (a gift from Charles Rice, The Rockefeller University, New York)
Trypan Blue solution, 0.4%, sterile filtered, suitable for cell culture (Sigma-Aldrich, catalog number: T8154)
Reagents required for surgery
Isoflurane, 3% (vol/vol) (Covetrus, catalog number: 029405)
Puralube vet ointment (Patterson Veterinary, catalog number: 07-888-2572)
Betadine solution 10% (vol/vol) povidone-iodine (Amazon, model number: 104457)
0.25% bupivacaine hydrochloride injection, 2.5 mg/mL (Covetrus, catalog number: 061842)
Meloxicam injectable solution, 5 mg/mL (Covetrus, catalog number: 049755)
Phosphate-buffered saline (PBS) (10×) pH 7.4, RNase-free, 0.9% (wt/vol) (Fisher Scientific, catalog number: AM9625)
Dietgel recovery (ClearH2O, catalog number: 72-06-5022)
D-Luciferin, potassium salt (GoldBio, catalog number: 115144-35-9)
Prescription-only medicines
Ethiqa XR (buprenorphine) extended-release injectable suspension, 1.3 mg/mL, C3 (Covetrus, catalog number: 072117)
Solutions
1× Phosphate-buffered saline solution (see Recipes)
Hepatocyte culture media (see Recipes)
50 mg/mL fibrinogen solution (see Recipes)
200 U/mL thrombin solution (see Recipes)
15 mg/mL D-Luciferin solution (see Recipes)
Recipes
1× Phosphate-buffered saline (PBS) solution
100 mL of PBS (10×) pH 7.4, RNase-free, 0.9% (wt/vol)
900 mL of distilled H2O
Hepatocyte culture media
432 mL of DMEM
50 mL of FBS
5 mL of Penicillin-Streptomycin
5 mL of ITS premix universal culture supplement
7.5 mL of HEPES (1 M)
0.4 µg/mL Dexamethasone
7 ng/mL Glucagon
50 mg/mL fibrinogen solution
1 g of fibrinogen from bovine plasma
20 mL of 1× PBS
200 U/mL thrombin solution
1,000 U thrombin from bovine plasma
5 mL of 1× PBS
15 mg/mL D-Luciferin solution
Reconstitute sterile D-luciferin powder to a concentration of 15 mg/mL in sterile 1× PBS. Aliquot and store at -20 °C. Thaw at room temperature prior to injection.
Laboratory supplies
CytoOne 35 × 10 mm TC dish (USA Scientific Inc, catalog number: CC76823340)
1.7 mL microcentrifuge tubes, PP (VWR, catalog number: 87003-294)
M-fold paper towels (McKesson, catalog number: 858231)
Equipment
Equipment for tissue culture
CellGard® ES Class II type A2 biological safety cabinet (NuAire, Inc., model: NU-475)
HeracellTM 150i CO2 incubator (Thermo Fisher Scientific, catalog number: 50116047)
Equipment for surgery
Dual procedure circuit/anesthesia chamber (VetEquip Inc., catalog number: 921400)
VaporGuard activated charcoal filter (VetEquip Inc., catalog number: NC9270393)
Germinator 500, dry sterilizer (Braintree Scientific, Inc., catalog number: GER 5287-120V)
Gel heating pad (Fisher Scientific, catalog number: 14-370-223)
Round hood to scavenge waste anesthetic vapors (Alsident System, catalog number: 1-7528)
Surgical light source (Stryker Berchtold Chromophare F 300)
Stainless steel digital animal weighing scale (Kent Scientific Corporation, catalog number: SCL-4000)
Towel drape, sterile, 18 × 25 (Dynarex, catalog number: 4410)
Alcohol preps, medium, 2-ply (Webcol, catalog number: 6818)
Self-seal sterilization pouches (VWR, catalog number: 89140-800)
PI polyisoprene surgical gloves, sterile (Protexis, catalog number: 2D72PT80X)
Transparent surgical tape (MD Supplies, catalog number: MPR-62201)
Inguinal hernia repair mesh Bard® flat sheet nonabsorbable polypropylene monofilament 3 × 6 in. rectangle style white sterile (McKesson, catalog number: 413782)
Sterile cotton-tipped wood applicator sticks (Patterson Veterinary, catalog number: 07-847-3562)
1 mL insulin syringes (Fisher Scientific, catalog number: 14-841-31)
Surgical scissors, sharp (Fine Science Tools, catalog number: 14002-13)
Graefe forceps (Fine Science Tools, catalog number: 110049-10)
Flat, round tip tweezers (Electron Microscopy Sciences, catalog number: 78333-33A)
Colibri retractor (Fine Science Tools, catalog number: 17000-03)
Hemostat (Fine Science Tools, catalog number: 13008-12)
Monocryl (poliglecaprone 25) suture (Ethicon, catalog number: Y303H)
Reflex wound clips (7 mm) (Fine Science Tools, catalog number: 12032-07)
Reflex clip removing forceps (World Precision Instruments, catalog number: 500347)
IVIS spectrum CT in vivo imaging system (Perkin Elmer, catalog number: 128201)
Software and datasets
Living Image 3.2 software (IVIS Imaging Systems)
Procedure
Part I: Construction of hepatic hydrogel
Preparation of human hepatocyte and fibroblast aggregates
Thaw cryopreserved human hepatocytes as per manufacturer’s instructions. Immediately after thawing, check hepatocyte viability with 0.4% Trypan Blue. A viability of 70%–90% is acceptable.
Resuspend hepatocytes in basal DMEM medium at 1.0 × 106 cells/mL along with lentiviral particles expressing firefly luciferase under human albumin promoter and 4 μg/mL polybrene in a 15 mL polypropylene conical tube by placing in the tissue culture incubator for 1 h. Place the conical tube horizontally inside the incubator and gently invert five times every 10 minutes to ensure proper mixing of hepatocytes and lentiviral particles.
Note: As hepatocytes quickly settle to the bottom of the tube if placed vertically in the incubator, it is important to place the 15 mL polypropylene tube on the side. Avoid constant shaking or inversion to minimize shear force and mechanical stress to hepatocytes.
Following transduction, spin hepatocytes at 50× g for 10 min and discard the supernatant. Wash cells with 10 mL of DMEM and spin at 50× g for 10 min. Repeat this step twice and resuspend cells in hepatocyte culture medium.
Note: Avoid spinning single-cell suspensions of hepatocytes above 50× g due to their sensitivity to mechanical stress, which could reduce viability.
Resuspend 1.2 × 105 transduced human hepatocytes and 0.6 × 105 mouse embryonic fibroblast J2-3T3 cells in hepatocyte culture media. Plate 1 mL of this cell suspension in each well of the AggreWell microwell plates and culture at 37 °C with 5% CO2. Change media every two days. Aggregates generally form after four days. However, transplantation of aggregates is done seven days post coculture as they are compact and show better survival (Figure 1).
Note: The ability of hepatocytes to form compact aggregates varies depending on the donor background. However, plating hepatocytes with J2-3T3 cells at a ratio of 2:1 always forms aggregates regardless of the donor background.
Figure 1. Hepatocyte–fibroblast aggregates in coculture after seven days. Representative (A) low (4×) and (B) high (10×) magnification images depicting consistent aggregate formation. The size of the aggregates is approximately 200 µm.
Preparation of hepatic hydrogel construct
Collect hepatocyte–fibroblast aggregates from the AggreWell microwell plates using P1000 pipette tips. Rinse the wells with DMEM basal medium twice to ensure all aggregates are collected. Transfer the collected aggregates to a 15 mL conical tube and spin down at 200× g. Remove the supernatant and resuspend the hepatocyte–fibroblast aggregates in DMEM basal medium. Each well of an AggreWell plate contains 1,200 hepatocyte–fibroblast aggregates and each aggregate contains 100 hepatocytes and 50 fibroblasts.
Note: Hepatocyte–fibroblast aggregates are less sensitive to mechanical stress as compared to hepatocytes in single-cell suspension. Hence, aggregates obtained seven days post coculture can be spun down at 200× g. Both P1000 and P200 pipette tips are well tolerated by aggregates.
Prepare the hydrogel by mixing fibrinogen (final concentration 10 mg/mL), thrombin (final concentration 1 U/mL), and the hepatocyte–fibroblast aggregates in DMEM to a total volume of 100 μL. Each hydrogel contains ~4,800 aggregates equivalent to four wells of an AggreWell plate. Thrombin must be added as the last component to trigger gelation of the fibrin gel. Mix quickly by pipetting and transfer to a sterile non-wettable parafilm sheet to form a bubble of the hepatic hydrogel construct.
Note: Avoid using medium containing FBS as it hinders gelation of fibrinogen after addition of thrombin into the gel mixture. We use fibrin as it is one of the many important extracellular matrix proteins present during wound healing to provide both structural support and angiogenic response of the host to enhance vascularization to the hepatic tissue posttransplantation.
Place the hepatic hydrogel construct immediately on a sterile mesh (~size of 5 mm × 5 mm) (Figure 2). Store the construct in a sterile Petri dish filled with hepatocyte culture medium and transport to the mouse facility for the surgical procedure.
Figure 2. Hepatic hydrogel before transplantation. Images of the hepatic hydrogel (A) after preparation and (B) prior to transplantation where it is placed on a sterile surgical mesh (scale bar: 1 cm). C. Zoomed-in image of the hepatic hydrogel showing aggregates inside the hepatic hydrogel (scale bar: 1 mm).
Part II: Implantation of hepatic hydrogel onto mesenteric parametrial fat pad
Preoperative preparations
Autoclave all surgical instruments before use.
Clean the workspace before surgery using a chlorine dioxide (Clidox-S) or an alcohol-based cleaning agent. Prepare the surgical station with dedicated spaces for the anesthesia induction chamber, surgical workspace, sterile surgical instruments, equipment area, and adjustable lights. Place the sterile instruments on the dominant-hand side of the surgeon’s workspace and ensure the lighting is adequate (Figure 3).
Figure 3. Surgical station setup for hepatic hydrogel implantation surgery. A. Surgical station setup. (1) Hood, (2) disinfectant (Clidox-S) spray bottle, (3) gas inlets, (4) timer, (5) isoflurane vaporizer, (6) spotlight, (7) microscope, (8) activated charcoal adsorption filter, (9) anesthesia chamber, (10) Betadine, (11) ophthalmic ointment, (12) alcohol pads, (13) cotton-tip applicators, (14) surgical gloves, (15) surgical tape, (16) towel drape, (17) paper towels, (18) needle uncapper, recapper, and syringe holder, (19) hot bead sterilizer, (20) hepatic hydrogel construct, (21) drug injections, (22) wound clips, (23) sutures, (24) sterile tools, (25) heating pad. B. Zoomed in view of sterile tools required for surgery. (1) Colibri retractor, (2) serrated tip forceps, (3) Graefe forceps, (4) surgical scissors, (5) hemostat, (6) wound clip applier, and (7) wound clip remover. C. Zoomed in view of drugs and supplies. (1) Hepatic hydrogel construct in a sterile Petri dish filled with hepatocyte culture medium and syringes for injecting (2) Bupivacaine hydrochloride, (3) Ethiqa XR, and (4) Meloxicam.
Ensure there is sufficient medical oxygen in the tank to perform surgery and add isoflurane to small-animal anesthesia system vaporizer as required before each case. Set up the anesthetic scavenging machine.
Weigh animals and record weights. Prepare dosages of drugs according to the respective animal’s weight.
Note: For batch surgeries, place a glass bead sterilizer near the aseptic operating field and perform quick dry heat sterilization by inserting tips of tools into the heating chamber for 15 s in between surgeries.
Surgical procedure
Note: Implementation of this protocol requires prior surgical training. Time required for training depends on the ability of the surgeon. With practice, motivated investigators can readily acquire the required surgical expertise.
Induce general anesthesia using inhaled isoflurane vapors by placing the mouse in an induction chamber using 3% (vol/vol) isoflurane and an oxygen flow rate of 1 L/min.
Confirm mouse has reached the surgical plane of anesthesia by gentle toe pinch. Lubricate both eyes with an ophthalmic ointment to prevent drying of the cornea during surgery, place the nose cone of the anesthesia system over the animal’s snout, and lower isoflurane vapors to 2% (vol/vol).
Note: Monitor depth of anesthesia every 15 min throughout the procedure to ensure that there are no changes in respiratory rate associated with surgical manipulation and toe pinch to confirm no movement.
Place the mouse on a heating pad in a supine position with the tail towards the surgeon and secure the forelimbs and hindlimbs with surgical tape. Administer a small volume (0.1 mL) of a local anesthetic agent such as bupivacaine into the tissue adjacent to the intended incision line and subcutaneously inject 2 mg/kg meloxicam and 3.25 mg/kg Ethiqa XR for preemptive analgesia.
Disinfect the surgical site by performing three sets of alternating scrubs of 10% povidone-iodine and 70% ethanol or 70% isopropyl alcohol. Paint the skin with 10% povidone-iodine and cover the mouse with a sterile surgical drape over the intended incision site with a cut of ~2 cm2.
Note: Make sure to put on sterile surgical gloves during the surgery.
Hold the skin using the Graefe forceps and make a lateral abdominal skin and muscle incision (approximately 1–2 cm long) to expose the peritoneum. A sterilized Colibri retractor is used to hold open the peritoneal cavity (Figures 4A and 4B).
Use the scissors to snip open the peritoneum (~1–2 cm). Identify and expose the parametrial fat pad (Figures 4C and 4D).
Note: Perform the incision between the large blood vessels running through the peritoneum to avoid causing unintentional bleeding. Be careful not to damage the abdominal organs during the incision.
Carefully place the sterile mesh carrying the hepatic hydrogel on top of the parametrial fat pad (Figure 4E).
Use a 5-0 Monocryl suture to sandwich the hepatic hydrogel firmly between the parametrial fat pad and surgical mesh by suturing the four corners (Figure 4F). During the procedure, irrigate the area with sterile PBS to prevent dryness and soak any excess buffer using sterile cotton swabs.
Note: Handle the parametrial fat pad as gently as possible to avoid puncturing or tearing the tissue.
Once the hepatic hydrogel is firmly sutured onto the parametrial fat pad, use the Graefe forceps to gently return the fat pad inside the peritoneum cavity (Figure 4G).
Close the peritoneum using an absorbable 5-0 Monocryl suture (Figure 4H) and remove the retractor.
Close the skin using 7 mm wound clips and wipe the skin around the surgical site with betadine (Figure 4I).
Figure 4. Step-by-step visual depiction of the hepatic hydrogel implantation technique for attachment to parametrial fat pad. A. A small incision is made through the skin (indicated by black arrow). B. Colibri retractor is placed to stretch open the skin. The peritoneum is opened by an incision (indicated by black arrow) to (C) locate and (D) expose the parametrial fat pad. E. The fat pad is gently spread out to prepare for implantation. Black arrow points to surgical mesh carrying the hepatic hydrogel. F. The edges of the surgical mesh are sutured on to the fat pad. G. The fat pad is carefully placed back into the peritoneal cavity using the Graefe forceps. H. The peritoneal opening is closed by suturing. I. Skin is closed by applying wound clips.
Postoperative procedures
Allow the mouse to recover in a clean cage by placing it on a warming pad and monitor until it regains consciousness. Check for return of a normal breathing pattern, normal blink reflex, and the beginning of ambulation. Add 2–4 moistened feed pellets to ensure the mouse is eating. Continue to intermittently monitor the condition of the mouse and the incision site for at least 30 min following the procedure and once more before the end of the day. Mice should be housed individually under standard laboratory conditions postsurgery, with food and water available ad libitum.
Fill out details on the surgery card and attach it to the mouse cage.
Administer meloxicam (2 mg/kg; subcutaneously) every 12 h for the next 48 h. Monitor mice at least twice a day for the next 72 h.
Part III. Validation through bioluminescence imaging
Intraperitoneally inject 250 µL of 15 mg/mL D-Luciferin dissolved in sterile 1× PBS into the mouse.
After 15 min, place the mouse in the anesthesia induction chamber with 2%–3% (vol/vol) isoflurane and an oxygen flow rate of 1 L/min (for ~5 min).
Lubricate both eyes with an ophthalmic ointment to prevent drying of the cornea during imaging and place anesthetized mouse in the supine position in the sample stage of the imaging chamber of the IVIS spectrum CT in vivo imaging system. This position helps in obtaining the highest bioluminescent signal.
Acquire images and quantify the bioluminescent signal on the IVIS spectrum CT in vivo imaging system using the Living Image software. Start image acquisition by setting luminescent exposure to 60 s and increment it as required.
Note: Bioluminescence imaging can be performed as early as 72 h posttransplantation and then weekly to monitor survival of the hepatic hydrogel in vivo.
Figure 5. Bioluminescent image of implanted hepatic tissue in nude mice. Control mouse (left) without any hepatic tissue implant and mice with hepatic tissue implants imaged at day 7 posttransplantation.
Data analysis
Bioluminescent signals acquired by the IVIS spectrum CT in vivo imaging system can be quantified by the Living Image software. The intensity of bioluminescence corresponds to survival of engineered tissues and hepatic function. Refer to Stevens et al. [4] or Baranski et al. [5], for representative bioluminescence images.
Validation of protocol
Parts of this protocol have been used and validated in the following research articles:
Stevens et al. [4]. InVERT molding for scalable control of tissue microarchitecture. Nature Communications (Figure 4, panels e and f).
Baranski et al. [5]. Geometric control of vascular networks to enhance engineered tissue integration and function. Proceedings of the National Academy of Sciences (Figure 4, panel e).
Stevens et al. [6]. In situ expansion of engineered human liver tissue in a mouse model of chronic liver disease. Science Translational Medicine (Figure 1, panels a and d).
General notes and troubleshooting
General notes
All reagents are molecular biology grade and must be stored in accordance with the manufacturer’s instructions.
Female mice are used as they have larger fat pads than males.
Make sure to get the surgical procedure approved by the Institutional Animal Care and Use Committee and only use methods that minimize pain and suffering.
All surgery instruments should be sterilized before use.
The surgery takes approximately 20–25 min per mouse. Suturing all four corners of the hepatic hydrogel construct to the fat pad takes approximately 5–8 min. We strongly recommend being very patient while performing this step as the fat tissue is very delicate and one has to be extra careful to not rupture the tissue while suturing.
Additional wound clips and/or sutures can be used if incision is larger than 1–2 cm.
Mice must be monitored twice daily for at least 72 h after surgeries.
Labored breathing, depressed respiratory rate, and/or gasping are indications of a failed surgery.
Troubleshooting
Large blood vessels run across the peritoneum and are visible to the eye. While making the incision, avoid cutting through these vessels to prevent excessive bleeding. However, if any large blood vessels are accidentally cut while making the incision, use a dry sterile cotton swab to stop the bleeding and rehydrate the area afterward using sterile 1× PBS.
If the nude mouse does not have enough parametrial fat pad or the surgical mesh is too large to fit appropriately onto the parametrial fat pad, we recommend carefully trimming the edges of the surgical mesh using sterile scissors before suturing.
If the mouse takes a long time to recover properly from anesthesia, it is possible that the surgical procedure has taken longer than it should. Administer 200 µL of sterile PBS subcutaneously.
If the mouse pulls off the wound clips, it is possible that these were not executed properly. Briefly anesthetize with isoflurane (3% vol/vol for induction, 2% vol/vol for maintenance), wash the incision site with saline solution, and repeat the wound clipping. Disinfect with 10% povidone-iodine solution.
If there are signs of pain or distress (i.e., hunched posture, ruffled hair coat, lack of appetite, dehydration), place the mouse under a heat lamp and administer 1 mL of warm sterile saline subcutaneously and intraperitoneally. Additional analgesic should be administered and DietGel Recovery can be provided for a speedy improvement. Consult with the designated veterinarian if signs of stress persist.
Acknowledgments
This work is based on our previous work published in Stevens et al. (2013) [4] and we acknowledge the work of Baranski et al. (2013) [5], which served as the baseline for adaptation. The authors would like to thank the Research Animal Resource Center (RARC) at Weill Cornell Medicine. This work was supported in part by grants from the NIH to Robert E. Schwartz (R01CA234614, 2R01AI107301, R01AA027327, and R01DK121072), from the DOD W81XWH-21-1-0978, and from The Paul G. Allen Family Foundation UWSC13448. Figures were created with BioRender.com.
Competing interests
Robert E. Schwartz is on the scientific advisory board of Miromatrix Inc. and Lime Therapeutics and is a speaker and consultant for Alnylam Inc. The remaining authors have no conflict of interest to declare.
Ethical considerations
All animal experimental procedures performed were approved by the Institutional Animal Care and Use Committee of Weill Cornell Medicine under protocol #2023-0020.
References
Starzl, T. E., Groth, C. G., Brettschneider, L., Penn, I., Fulginiti, V. A., Moon, J. B., Blanchard, H., Martin, A. J. and Porter, K. A. (1968). Orthotopic Homotransplantation of the Human Liver. Ann. Surg. 168(3): 392–415.
Sinha, S., Hassan, N. and Schwartz, R. E. (2023). Organelle stress and alterations in interorganelle crosstalk during liver fibrosis. Hepatology: e0000000000000012.
Li, T. T., Wang, Z. R., Yao, W. Q., Linghu, E. Q., Wang, F. S. and Shi, L. (2022). Stem Cell Therapies for Chronic Liver Diseases: Progress and Challenges. Stem Cells Transl. Med. 11(9): 900–911.
Stevens, K. R., Ungrin, M. D., Schwartz, R. E., Ng, S., Carvalho, B., Christine, K. S., Chaturvedi, R. R., Li, C. Y., Zandstra, P. W., Chen, C. S., et al. (2013). InVERT molding for scalable control of tissue microarchitecture. Nat. Commun. 4(1): 1847.
Baranski, J. D., Chaturvedi, R. R., Stevens, K. R., Eyckmans, J., Carvalho, B., Solorzano, R. D., Yang, M. T., Miller, J. S., Bhatia, S. N., Chen, C. S., et al. (2013). Geometric control of vascular networks to enhance engineered tissue integration and function. Proc. Natl. Acad. Sci. U.S.A. 110(19): 7586–7591.
Stevens, K. R., Scull, M. A., Ramanan, V., Fortin, C. L., Chaturvedi, R. R., Knouse, K. A., Xiao, J. W., Fung, C., Mirabella, T., Chen, A. X., et al. (2017). In situ expansion of engineered human liver tissue in a mouse model of chronic liver disease. Sci. Transl. Med. 9(399): eaah5505.
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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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Peer-reviewed
Measuring Heart Rate in Freely Moving Mice
JS Jérémy Signoret-Genest *
NS Nina Schukraft *
PT Philip Tovote
(*contributed equally to this work)
Published: Vol 14, Iss 3, Feb 5, 2024
DOI: 10.21769/BioProtoc.4926 Views: 1034
Reviewed by: Xi Feng Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Neuroscience Feb 2023
Abstract
Measuring autonomic parameters like heart rate in behaving mice is not only a standard procedure in cardiovascular research but is applied in many other interdisciplinary research fields. With an electrocardiogram (ECG), the heart rate can be measured by deriving the electrical potential between subcutaneously implanted wires across the chest. This is an inexpensive and easy-to-implement technique and particularly suited for repeated recordings of up to eight weeks. This protocol describes a step-by-step guide for manufacturing the needed equipment, performing the surgical procedure of electrode implantation, and processing of acquired data, yielding accurate and reliable detection of heartbeats and calculation of heart rate (HR). We provide MATLAB graphical user interface (GUI)–based tools to extract and start processing the acquired data without a lot of coding knowledge. Finally, based on an example of a data set acquired in the context of defensive reactions, we discuss the potential and pitfalls in analyzing HR data.
Key features
• Next to surgical steps, the protocol provides a detailed description of manufacturing custom-made ECG connectors and a shielded, light-weight patch cable.
• Suitable for recordings in which signal quality is challenged by ambient noise or noise from other recording devices.
• Described for 2-channel differential recording but easily expandable to record from more channels.
• Includes a summary of potential analysis methods and a discussion on the interpretation of HR dynamics in the case study of fear states.
Keywords: Heart rate ECG Mouse States Freely behaving
Background
Over the past decades, measurements of cardiac function have been exploited not only in cardiovascular research but also in other disciplines like neuroscience due to their tight connection with neural processes and as readouts for emotional states (Carrive, 2000; Leman et al., 2003; Tovote et al., 2005; Stiedl et al., 2009). To take into account the integrated nature of cardiac and motor functions during emotional challenge, we recently developed a novel analytical framework. By analyzing cardiac parameters together with behavioral readouts, we unveiled critical cardio-behavioral states that would not have been detectable with either of the readouts alone (Signoret-Genest et al., 2023).
A variety of non-invasive and invasive methods have been established in order to measure cardiovascular parameters. Non-invasive approaches include surface recording (different electrodes are embedded in the floor and contacted by the individual paws) or external telemetry systems with instrumented jackets, which allow for simultaneous respiratory function monitoring (Chu et al., 2001; Sato, 2019; Fares et al., 2022). However, the signals acquired with these methods are prone to be noisy, compromising the final data quality, and the restricted experimental conditions (e.g., specific testing box) or added burden on the animal (jacket) might hinder the expression of naturalistic behaviors, thereby limiting their scope of application.
An electrocardiogram (ECG) records the electrical charge shifts that occur during a cardiac cycle. Non-tethered telemetry recording systems that record the ECG allow the animal to move freely and are thus ideally suited for long-term recordings (Calvet and Seebeck, 2023). However, they are both invasive, since they require a battery and components for wireless transmission, as well as rather expensive, and may additionally pose data synchronization challenges. On the other hand, tethered systems to record the ECG are inexpensive and easy to establish but must be optimized in order to successfully deal with potential hurdles such as environmental interferences or experimental artifacts.
Tethered ECG recordings, in particular, grant researchers direct access to raw data, which must then undergo proper pre-processing to extract the relevant biological phenomenon, namely heartbeats. As for any other technique, heart rate data then needs to be processed to answer biological questions. However, despite the apparent ubiquity of this readout, the selection of analyses, data sub-selection, treatment, and interpretation are not immune to pitfalls and must be approached with caution.
Here, we present a protocol for cost-effective, highly reliable, and user-friendly ECG recordings in freely behaving mice. The protocol encompasses the fabrication of the patch cable and ECG connector implants, detailed surgical implantation procedures, and subsequent data acquisition and processing steps. We also provide access to a custom graphical user interface (GUI) for heart rate extraction, along with specific (pre)processing strategies. We discuss data processing and interpretation through the scope of recent data, showing that there is more to heart rate–related readouts than simple averages or single heart rate variability (HRV) values, and that such minimalist processing could reduce the statistical power of studies or introduce biases.
Materials and reagents
Reagents
Orthophosphoric acid (Carl Roth, catalog number: 6366.1)
Paladur, liquid component (Anton Gerl, catalog number: 82462)
Buprenorphin (Bayer, catalog number: 14439113)
Isoflurane (cp-pharma, catalog number: 1214)
Depilatory cream (Veet)
Cutasept F (Hartmann AG, catalog number: 9803650)
Naropin (Aspen Germany, catalog number: 02749854)
Vitagel (Bausch & Lomb, catalog number: 1318187)
Braunol (Braun, catalog number: 190971)
Metacam (Boehringer Ingelheim, catalog number: 08890217)
Laboratory supplies
Circular miniature connector, male (OMNETICS, catalog number: A79108-001)
Wire, stainless steel, 7 strand, PFA (AM Systems, catalog number: 793200)
Soldering tin (Stannol, catalog number: 574006)
Glue, transparent (Silisto, catalog number: 71024)
Ultra-flexible microminiature shielded cable (Daburn, catalog number: 2721/5)
Blunt needles, 27 G (Braun Petzold, catalog number: 9180117)
M9 connector, male (Binder, catalog number: 99-0413-00-05)
Heat shrink tubing set (Haupa, catalog number: 267200)
Regular insulated wire (RadioSpare, catalog number: 872-5167)
Circular miniature connector, female (OMNETICS, catalog number: A79109)
Teflon tape (Toolineo, catalog number: 100000000178797)
Aluminum foil
1 mL syringes (Praxisdienst, catalog number: 138322)
Needles, 30 G (Praxisdienst, catalog number: 128330)
Cotton buds
Cotton wipes (Maicell, catalog number: 72100)
Glue, viscous (Pattex, catalog number: 2804443)
Scalpel blade (Hartenstein, catalog number: SJ10)
Suture material, ECG (Serag Wiessner, catalog number: LO07340B)
Suture material, skin, braided silk (SMI, catalog number: 8200518)
Toothpicks
Glue, black (Wekem, catalog number: WK-2400)
Equipment
Third hand (Toolcraft, catalog number: TO-6871371)
Stereomicroscope (Olympus, model: SZ-61)
Forceps, straight (Fine Science Tools, catalog number: 11252-00)
Soldering station (Velleman, model: VTSSC40N)
Scissors (Passau Impex, catalog number: 14060-10)
Cable stripper (Toolcraft, catalog number: TO-4861971)
Lighter
Multimeter (Fluke, model: 179)
Balance (Kern, model: EMB 2200-0)
Anesthetizing box (Hugo Sachs Elektronik, catalog number: 50-0108)
Anesthesia mask (Hugo Sachs Elektronik, catalog number: 73-4858)
Anesthetic vaporizer (Hugo Sachs Elektronik, catalog number: 34-2052)
MiniVac gas evacuation unit (Hugo Sachs Elektronik, catalog number: 73-4910)
Fluosorber filter canister (Hugo Sachs Elektronik, catalog number: 34-0415)
Heating pad (Kent Scientific, model: RightTemp Jr.)
Forceps, curved (Fine Science Tools, catalog number: 11271-30)
Forceps, straight, blunt (Fine Science Tools, catalog number: 11002-14)
Feeding cannula (Fine Science Tools, catalog number: 18061-75)
Needle holder (Passau Impex, catalog number: T220218)
Scalpel handle (Fine Science Tools, catalog number: 10003-12)
Infrared lamp (Medisana, catalog number: 88232)
Amplifier with headstage (NPI Electronic, model: DPA-2FX)
Analog acquisition system with digitizer (e.g., Plexon Omniplex)
Large custom-made box (e.g., 1 m × 1 m × 1 m)
Custom-made Plexiglas cylinder (specifications depend on the conducted behavioral experiment; here, a cylinder with 30 cm diameter, 50 cm height, and a wall thickness of 0.4 cm was used)
Software and datasets
Acquisition software
Dedicated software associated with a system to record analog signals. The data presented here was recorded with a digital Omniplex system (Plexon, Dallas, TX, USA) designed for electrical recordings of neuronal activity.
Analysis software
MATLAB 2022a (back compatibility at least until release 2019a; MathWorks, 09/03/2022)
ECG_Process package (custom-written code); current version can be found on: https://github.com/Defense-Circuits-Lab/ECGanalysis
Prism v9.5 (GraphPad, 26/01/2023)
Procedure
Manufacturing of ECG connectors
See General Note 1.
The circular miniature connector has six pins, from which three are used for manufacturing the ECG connector. Fix the miniature connector in the clamp of a third hand and bend all five peripheral pins to 90°. Pinch the central pin with fine forceps and twist it to break it close to the base (Figure 1).
Note: If there is no use for the additional pins, they can also be removed or bent on the sides to increase support.
Figure 1. Manufacturing of electrocardiogram (ECG) connectors. A. Three pins are bent towards the outside and the three remaining pins are cut short. B. The stripped end of the wire is soldered onto the pin. C. A second ECG and a grounding wire are soldered to the two other pins. D. A layer of glue is applied to the inside of the connector and on all pins and stripped wire parts. Scale bar: 5 mm.
Solder two wires onto the miniature connector pins #1 and #2:
Cut two pieces of wire to a length of 7 cm.
Strip 5 mm from one side of the wire with fine forceps.
Add a very small amount of orthophosphoric acid on the pin.
Align the stripped wire onto the pin.
Take a small amount of soldering tin on the iron and briefly touch the pin/wire ensemble.
Note: Soldering should feel easy—the melted tin should instantly run onto the pin, making a distinctive sound. The result should be a very thin soldering, looking shiny. The tin should not be kept on the iron for too long beforehand, and the iron should not remain on the pin beyond an instant, as it could loosen it internally by heating the connector.
Repeat the steps for pin #2.
Solder a reference wire with a length of 2 cm onto connector pin #6 by repeating step A2.
Test the soldering quality; pulling each wire with a quick and firm motion allows to assess whether the wires are properly soldered onto the pins.
Note: A good soldering can withstand substantial pulling, which is a predictor for high-quality recordings.
Apply glue to all connections and stripped pieces of wire.
Check the connections, fitting a dedicated female connector (same model as for the cable) onto the connector. Strip the very tip of the electrodes’ wires. Contact each pair of female connector’s pin/electrode with fine tools mounted on a multimeter to check for proper connection. Just in case, while still touching each wire, touch the successive pins on the female connector; only the proper one should have a low resistance.
Manufacturing the patch cable
See General Note 2.
Cut the appropriate length of microminiature shielded cable. This should allow the cable to run freely from the headstage to the mouse’s head while being held by a pulley, taking into account possible loops that appear during long recordings.
Note: Potential damage is usually located on the mouse’s side and is due to bites or twisting-induced rupture of a wire inside the insulation. To save time and resources, it makes sense to make the cable much longer so that only the mouse’s side can be remade when necessary. The ideal length depends on how the extra cable length can be handled without risking damage through bending or rolling.
Assemble the headstage side of the cable (Figure 2).
Note: It is advisable to start on this side to be able to check that the sheath properly connects the distal end of the mouse side, shielding all the way to the wire on the headstage side.
Figure 2. Manufacturing of the patch cable’s headstage side. A. Preparation of the miniature shielded cable. The cable is deinsulated (i) and the sheath is carefully unbraided (ii). The individual wires are unwinded, deinsulated (iii), and soldered onto the pins of the connector (iv). B. An extra wire is used as the grounding reference (i) by soldering it onto the sheath of the patch cable (ii). Heat shrink tubes of different sizes are used to insulate the connection (iii).
Begin by sliding the M9 connector casing onto the cable, followed by heat shrink sleeves of varying diameters: one nearly as large as the threaded part of the connector, one that fits the cable loosely, and one of intermediate size.
Carefully remove the insulation over approximately 1 cm and gently unbraid the sheath by using the tip of a blunt needle to wedge between the weave and pull towards the cut side. Loosely bundle the sheath on one side. Unwind the individual wires and arrange them according to Figure 2.
Note: Avoid applying excessive force, as the insulation on the individual wires is delicate and inadvertent damage could lead to discarding the final product due to internal short circuits.
Strip the end of the wires over 1–2 mm.
Maintain tight strand alignment, briefly dipping them into a drop of orthophosphoric acid placed on a surface, and then apply soldering tin.
Note: Ensure that no single strand from any of the individual wires protrudes, as it may complicate the subsequent steps. Additionally, be aware that the wires’ insulation tends to shrink significantly when exposed to excessive heat, so exercise caution during soldering.
Position the cables so that the end to be soldered is placed on top and aligns with the pins of the connector. You can achieve this by securing the cable around one of the binocular knobs and rotating the connector using a third hand tool, positioning the pins in front of the corresponding wires.
Apply a very small drop of acid to each pin.
Note: Use only a minimal amount of acid, as excessive use may lead to excessive corrosion over time.
Insert the first wire into the first pin and apply some soldering tin. The solder should be heated until it becomes fully liquid on the pin but take care not to overheat it, as it could melt the insulation on the thin wire. Repeat this process for the remaining pins.
Adjust the angle of the connector to ensure perfect alignment with the cable. Apply tension to the cable, for example by gently pulling it and adding a weight to prevent it from becoming slack.
Apply glue to the pins and wires, starting from the bottom of the pins and progressing to the top of the stripped part of the cable. Do this in several stages, incrementally adding glue and using Paladur's solvent to quickly cure the adhesive.
Note: Be mindful not to extend the glue beyond the normal dimensions of the connector. Pay attention to the fact that the glue may briefly become more liquid after the application of Paladur's solvent.
Critical: Starting from this point, the connection between the non-deinsulated and insulated parts of the cable becomes extremely fragile, and the wires can easily break. Avoid bending or twisting them at all.
Move the smallest piece of heat shrink tubing closer to the connector.
For the grounding wire, prepare a piece of regular wire by deinsulating ~1 cm and coating the exposed wires with soldering tin.
Insert this wire into the piece of heat shrink tubing (along with the cable that is already inside) and position the exposed wire next to the sheath.
Apply a small amount of acid and solder both components together.
Gently slide the smallest heat shrink tubing all the way down to the glued section and then apply heat.
Critical: Be cautious not to apply too much heat or for too long when shrinking the tubing. The insulation of the individual wires within the cable is very thin and can easily melt.
Successively add and shrink the other two pieces of tubing.
Critical: Aim to keep the assembly as straight as possible. Otherwise, it might make it difficult to put the connector's casing in place.
Carefully slide down the various components of the connector. The cable gland contains teeth that will be tightened by the distal nut to grip the cable, so it is advisable to disassemble the connector into its different parts.
i. Start with the main shaft, which should be screwed onto the part of the connector where the pins were.
Critical: When screwing, ensure that the shaft does not twist the heat shrink and, consequently, the entire cable/wire assembly. Since they are soldered together, twisting could cause them to snap.
ii. Then, add the teeth.
iii. Finally, screw the nut onto the main shaft to secure and tighten the teeth, ensuring a firm grip on the heat shrink section.
Fabricate the mouse side of the cable (Figure 3).
Figure 3. Manufacturing of the patch cable’s mouse side. A. Bent pins of the miniature connector held in a crocodile clamp. B. Thin soldered connection between individual wires and pins of the connector. C. Embedding in glue. D. Two layers of Teflon tape are wrapped around the connection. E. A piece of aluminum foil is prepared to insulate the connection. F. Wrapped aluminum foil. G. Two heat shrink tubings build the cover of the connection.
Begin by sliding two heat shrink pieces of different diameters: one slightly larger than the connector and the other with a loose fit around the cable.
Note: If you want to exercise extra caution or if you have doubts about the headstage side, now is an opportune moment to check if the pins on the connectors at the headstage side are still properly connected to the individual wires by testing for a continuous electrical connection and no between-wires short circuits. Use a multimeter in resistance checking mode for this purpose.
Prepare the cable end as previously instructed but strip less of the insulation.
Secure the connector in a crocodile clamp, bend the pins of the connector outward, and position the cable so that it lays on top of them.
Solder the individual wires according to the provided figure.
Critical: The soldering should be as thin as possible and exhibit a shiny appearance.
Return the connector pins to a straight position.
Note: Ensure there are no short circuits (solder touching another solder or a neighboring pin). If necessary, the pins can be bent laterally as long as the external diameter remains within the connector's limits.
Embed the assembly in glue, trying to bring the wires closer together as they approach the cable.
Cut a piece of Teflon tape and wrap it around the glue, without deforming it first. Apply at least two layers. This step is crucial to ensure proper insulation between the soldering and the shielding in case the glue does not cover the entire area.
Note: To prevent the Teflon from shifting, it is advisable to add a thin film of superglue to the already cured glue just before placing the Teflon.
Prepare a piece of aluminum foil that matches the distance between the bottom of the connector and the stripped part of the cable. Apply glue to the connector and up to the cable (avoiding the sheath), and securely affix one layer of aluminum foil. Trim any excess foil.
Spread the cable sheath over the foil layer. Lower the tightest piece of tubing and shrink it.
Check the reference wire by confirming that the wires’ headstage side is well connected to the aluminum foil layer by using a multimeter.
Carefully lower the other piece until it reaches the edge of the connector (but avoid excessive extension) and then shrink it.
Ensure that all wires are correctly connected. Secure both connectors in clamps and use a multimeter with a fine tool (or a wire) to verify that each pin is linked exclusively to its corresponding counterpart and no other (Figure 4).
Figure 4. Final check of proper wire connection. A. Both ends of the cable are completed. B. A multimeter is used to confirm that all pins of the headstage side are properly connected to the cable’s mouse side connector.
Surgery: Implantation
Prepare the surgical field and all equipment needed throughout the surgery (Figure 5).
Figure 5. Preparation of the surgical field
Prepare the animal (Figure 6A).
Figure 6. Surgical procedure of electrocardiogram (ECG) connector implantation. A. Dorsal (i) and ventral (ii) view of the mouse after preparing it for the implantation of the wires. B. The feeding cannula is inserted subcutaneously from the chest towards the incision on the head. The cable of the ECG connector is threaded through the hollow cannula. C. A small ball of glue is applied to the end of the stripped wire and a knot to suture the wire on the underlying muscle tissue is placed closely to it (i). The incision on the skin is sutured with 2–3 knots (ii). D. Exposed skull surface is dried (i) and roughened (ii, iii). The connector is glued onto the skull (iv) and a layer of black glue is added (v, vi).
Weigh the animal and inject the adequate amount of Buprenorphin (dosage 0.1 mg/kg) subcutaneously in the neck area 20–30 min prior to the start of the surgery for perioperative analgesia.
Anesthetize the animal by placing it into the isoflurane induction chamber (4% induction, 0.6 L/min oxygen flow).
Transfer the mouse on the heating pad and place the head in the anesthesia mask.
Check for paw reflexes to ensure deep anesthesia has been reached.
Maintain anesthesia with 1.5% isoflurane in oxygen.
Apply depilatory cream on top of the head to remove hair from a 1 × 1 cm area.
Turn the animal on its back and apply depilatory cream on the upper right and lower left of the animal’s chest.
Bring it back into prone position to access the head again.
Disinfect the skin by applying Cutasept three times and place 200 µL of Naropin using a 1 mL syringe directly underneath the skin on the top of the skull for local anesthesia.
Open the skin.
With sharp scissors, cut off a 0.5 × 0.5 cm patch of skin on the head.
Turn the animal into supine position and disinfect the skin on the chest by applying Cutasept three times.
Cut a small incision into the skin on the chest with sharp scissors (3–5 mm).
Thread the ECG wire (Figure 6B).
Use blunt forceps to carefully dissect the underlying tissue from the skin.
Insert the feeding cannula into the incision and gently push it dorsally towards the opened skin on the head.
Critical: Always keep the cannula’s angle such that the tip faces outwards to prevent damage to any organs. Sliding the cannula should not require a lot of strength. If it feels otherwise, it is likely that the cannula is not being threaded subcutaneously. Keep the cannula in sterile water before insertion for it to glide smoothly onto the tissues and sterilize it thoroughly afterwards.
Pass the proper wire from the ECG connector through the hollow feeding needle from the head side. When the wire reaches the chest opening of the cannula, carefully pull the cannula out, leaving the wire in place.
Note: Some conjunctive tissue can obstruct the cannula. In that case, just poke it with a needle to free the hole. Once the cannula is out, remove all tissue and sterilize it. Try to consistently use the wire that will be on the right side of the connector for the right chest, and similarly for the left.
Prepare the wire.
Strip 5 mm of the wire tip by using fine forceps.
Apply viscous glue to the very end of the wire tip. Do not spread it too much and make sure you leave enough stripped wire. Polymerize the glue by applying a small amount of Paladur’s solvent.
Note: The tip of the wire can be pressed between some forceps to flatten it (just the tip) so that the individual strands are slightly spread, making it easier to keep glue there.
Fix the wire and skin suture (Figure 6C).
Dissect conjunctive tissues with forceps to expose the muscle and suture the wire onto it, going perpendicular to the muscle fibers. The knot should be placed close to the glue.
Note: Leave the wire long enough that it does not impede the animal’s locomotion afterwards. It is possible and better to keep a small loop on the head side, which is then fitted subcutaneously.
Close the skin with two interrupted sutures and disinfect the wounds with iodine-based disinfectant (e.g., Braunol).
Repeat steps C4–C6 on the upper right of the chest with the second wire of the ECG connector.
Fix the connector on the mouse’s head (Figure 6D).
Turn the animal into prone position again. Remove any remaining tissue from the skull bone by rubbing it with a cotton bud. Carefully roughen the skull surface by slightly scratching it with the back of a scalpel blade.
Note: Proper fixation of the head in a stereotactic frame facilitates these steps, but this can also be performed in the anesthesia mask.
Glue the miniature connector onto the skull and polymerize the glue with a small amount of Paladur’s solvent.
Note: The skull needs to be absolutely dry.
Strip a small portion of the remaining wire and slide it subcutaneously before gluing it into place by applying glue on the bone.
Form a small head cap by adding more glue around the connector and polymerize it.
Apply a layer of opaque black glue.
Critical: Make sure that enough skull surface is exposed and recruited for the cap.
Inject metacam (dosage 2 mg/kg) subcutaneously, transfer the animal to its cage, and let it recover under an infrared lamp.
Monitor the animal until it has fully recovered from the anesthesia.
Monitor the animal’s recovery every 12 h for 7 days post surgery.
For long-term post-surgery care, monitor weight and general state daily.
Note: Pay close attention to the sutures. This surgery is usually very well tolerated and animals fully recover behaviorally after approximately 1 h and regain their pre-surgery weight after 3–4 days.
See General Note 2.
(Optional) At the end of the experiments, retrieve the head cap and place in acetone overnight. Acetone will dissolve the glue but will not damage the connectors if left only for half a day. Rinse with water and dry. Check carefully for pin integrity and general condition. The connector can be reused a few times if handled with care.
Critical: Acetone will dissolve and dilute the cyanoacrylate glue but upon evaporation it could still leave a layer of glue; several head caps can be placed in acetone at the same time but not too many, and the amount of acetone should be consequent. If left too long in acetone, the connectors will also start being damaged; check periodically whether the glue is dissolved and change the acetone solution if needed. Take extra care when checking the connectors before reusing them: connect a female connector and make sure that there is a connection between each pin of the female connector and its corresponding pin on the (reused) male connector. Also check each “ready-to-implant” connectors as described in step A6.
Data acquisition
Let the animal recover for at least one week before data acquisition.
Familiarize the animal with handling and the connection procedure over several days.
Note: Handling the animal for a minimum of three days significantly improves subsequent connection procedures. On the first day, allow the animal to explore the palm of the hand multiple times. On subsequent days, gradually habituate the animal to being held in the hand by gently gripping the head cap with the index finger and thumb while securing the tail with other fingers to prevent escape.
The day of the recording, connect the animal to the patch cable and place it in a behavioral arena, ideally with constant video monitoring.
Notes:
A commutator could be used to prevent entanglement, but a simple pulley system with two grooved wheels, a lightweight thread, and a 5 mL syringe as a counterweight works well. Gently tie the thread to the cable and the syringe head. Adjust the counter pull by modifying the water level in the syringe. The ideal knot position on the cable should be determined empirically (Figure 7).
Try as much as possible to only turn the amplifier on whenever the cable is connected to a mouse to prevent saturation of the system and potential damage.
Figure 7. Recording setup with pulley system. A thread is tied to the electrocardiogram (ECG) cable and runs around two pulleys. An adjustable counterweight is tied to the other end of the thread.
Turn on the acquisition system that receives the amplifier’s output to display it online and save it to a file. In order to preserve enough resolution in the ECG signal and allow for accurate heartbeat timestamping, the sampling rate should ideally be 5 kHz or higher (keep it mind that too high is not useful and will increase file size and processing time).
Note: The current example is based on an NPI amplifier and a Plexon acquisition system (Omniplex/Cineplex).
Note: For file format considerations and pre-processing, see Data analysis section A.
Critical: If experiments involve aligning ECG data with behavioral data, it is essential to ensure perfect synchronization between the ECG signal and the video recording system. This synchronization should be carefully planned before the experiments begin. Recording systems rely on internal clocks to sample signals, and while these clocks are generally quite accurate individually, issues can arise when dealing with two separate systems and their clocks:
a) They may start at different times (one system might be slower to start or require manual triggering), leading to an initial time lag (shift).
b) Their clocks may run slightly faster or slower than each other, resulting in time discrepancies. For example, what appears as t = 10 min on one system could be t = 9 min 56 s on the other (drift).
Addressing these synchronization challenges upfront is crucial for precise data alignment and to prevent any misinterpretation further down the line.
Adjust the amplifier settings (these settings may vary between systems, but the following parameters are commonly present):
Channels: There is a reference channel and one “signal” channel for each electrode. In any case, the signal for each electrode is the voltage difference between the reference and that electrode. However, for each channel, we can also choose to output the difference between the two electrodes.
Note: Unless one electrode's signal is significantly bad (contaminated with noise), it is a good idea to set one channel to output the differential signal and keep the other as a single output, using the better channel for this purpose. This choice should be made individually for each mouse, which should not lead to analytical issues, except for very specific analyses (e.g., interested in heartbeat morphology) where the configuration would matter.
Filters: The low-pass and a high-pass filters can be adjusted. The low-pass filter keeps lower frequencies in the signal while reducing higher frequencies. Conversely, the high-pass filter removes low frequencies and keeps high frequencies.
i. Low-pass filter: Somewhere around 1 kHz.
ii. High-pass filter: A value is set to remove the slow fluctuations in the ECG signal. 30 Hz is a good option. If the amplifier features a 50 Hz (or 60 Hz, depending on the local power grid) notch filter, it can be beneficial to activate it when the signal is affected by prevalent grid-related interferences.
Note: The amplifier’s analog filters do not sharply cut off specific frequencies in the signal. Instead, they have a gradual effect on the frequency spectrum. For example, if a high-pass filter is set at 30 Hz, it will reduce lower frequencies while allowing higher frequencies to pass through to varying degrees. It can be thought of as a gentle slope in how it affects the different frequencies. This means that when setting these filters, it is essential to consider the scope of the study. If the goal is not to study heartbeat morphology, it seems reasonable and more robust to directly adjust the filter settings to improve heartbeat separability from background over wave structure.
Gain: Ensure that the gain is set high enough to clearly visualize the signal without saturating it, which means avoiding the signal hitting the upper or lower boundaries of the acquisition system. If saturation occurs, the signal will appear as a flat line at those extreme values.
Note: Refer to “Troubleshooting Problem 2” for a comprehensive guide and tips on initially setting up the equipment and identifying potential issues.
Offset: The offset allows you to adjust the baseline level of the signal, effectively moving it up or down. Aim to position the baseline (typically found between heartbeats) close to the middle values of the acquisition system, e.g., approximately 0 V if the system’s range goes from -5V to 5V. This adjustment helps to maintain the signal within the system’s optimal operating range.
Data analysis
Heartbeat extraction
In this section, the process of extracting heartbeats using custom MATLAB code is described. This extraction is user friendly and does not require any coding experience. We are providing MATLAB code as regular and live scripts to allow to perform the different steps from the associated data set.
Loading the data.
The GUI accommodates various data sources, including simple times/values series in text/.csv files, .mat files, .pl2 files (Plexon), and .tsq files (Tucker-Davis).
Note: While most systems allow to export data as csv files, the list of supported file formats can easily be expanded as needed. It only requires either the dedicated API for MATLAB to access the proprietary file format or, in other cases where the files are disguised text files, knowledge about the headers and data organization within the file.
Click Start path to select a starting folder (optional, for convenience).
Click Load file to choose a data file. You can also select "_HeartBeats.mat" files (output files) to reload the analyses within the GUI (Figure 8).
Figure 8. Load files into the processing graphical user interface (GUI). As an option, you can define a “start path” to select batch folders. Click load file to choose the first file to be pre-processed.
Raw ECG pre-processing.
Raw ECG signals can undergo pre-processing, which includes:
Detrending: Originally designed for human recordings with significant baseline shifts, this feature could also benefit recordings from other systems. It involves computing a sliding mean with a large window (covering multiple heartbeats) and subtracting it from the raw signal.
Bandpass filtering: A zero-phase bandpass filter can be applied to remove slow fluctuations and noise (zero-phase filtering ensuring no signal shift).
i. Enable detrending and/or bandpass filtering by ticking the corresponding boxes.
ii. Adjust the values in the edit boxes as needed (Figure 9).
Figure 9. Pre-processing of raw electrocardiogram (ECG) signals. Detrending and bandpass filtering options can be enabled by ticking the respective boxes. Adjust the values as needed.
Transformation of the ECG signal.
The ECG signal is elevated to the power of n (where n is an even integer) to amplify heartbeats compared to the background. Typically, elevating the signal to the power of 4 yields optimal results, but it can be experimented with different values empirically. The resulting signal is then smoothed with a Gaussian kernel large enough to encompass one heartbeat, creating smooth peaks.
Note: Some ECG signals may contain noise with similar amplitude and frequency (sharp artifacts). In such cases, reducing the smoothing window size can be beneficial, preventing noise from merging with the peaks. However, this may result in an increased number of detected peaks. In moderate cases, the algorithm can handle this effectively.
Adjust the values in the edit boxes and refine with the following steps if necessary.
Thresholding and putative heartbeat extraction.
The elevated/smoothed signal is thresholded to detect potential heartbeats. The "Window" parameter determines the number of samples before and after the threshold crossing to be selected as a single waveform (potential heartbeat).
Note: This parameter is mainly important for the algorithm step but does not need to be extremely tight around one heartbeat. What matters is that it captures the whole beat without overlapping with another one.
On the ECG plot, identify the window with the smallest amplitude for heartbeats and zoom in on that area. Adjust the blue line to cross the peak corresponding to the smallest heartbeat (approximately 1/5 of its height; Figure 10).
Check the heart rate curve for sharp, unnatural drops indicating missing heartbeats. Zoom in on suspicious areas to confirm the heartbeats were not detected (no filled diamond) and adjust the threshold if necessary.
Note: It is crucial to make sure that all heartbeats are detected at this step. Further steps involve manual cleanup that could be wasted if some beats are missing, since readjusting the threshold will reset the rest.
If this is one of the first recordings in these conditions, tick the Plot waveforms checkbox. This will display, for each thresholded peak, the corresponding waveform directly on the ECG trace. Adjust the values for the Window parameter (number of samples before and after threshold crossing) so that the main features of a single beat are included (as visible on the overlay).
Note: Keeping the waveforms and/or peaks always plotted can lead to a decrease in GUI reactivity.
Figure 10. Thresholding to detect potential heartbeats. Adjust the threshold (blue line) either by dragging the line or by typing in a value to cross the smallest peak in the recording.
Run the algorithm.
The algorithm operates iteratively, systematically analyzing successive putative heartbeats. Here is a concise overview of its operation:
Template Creation: The algorithm begins by constructing a representative waveform derived from all the putative heartbeats. This template serves as a reference for comparing with potential peak candidates, computing correlation values.
Iterating: Starting from an area where at least a few consecutive peaks have a good correlation score, it will progress to the next peak and iteratively do so from peak to peak.
For each peak, the algorithm computes a mean score from the peak’s correlation score and a position score. To obtain the position score, the algorithm creates a Gaussian probability curve based on previous (previously validated) beats. If the following peak falls close to the average from the previous intervals, the score is maximal and slowly decreases for values that deviate.
When either or both scores have critically low values for all putative heart beats in a given time window, the algorithm engages in parallel paths. In each path, it selects one of the closely located peaks as the valid heartbeat and discards the others. For each path, the algorithm processes the corresponding ECG signals for a few seconds of recordings, as mentioned here. The algorithm then computes a global score for the segments obtained from each possibility, considering an aggregated score derived from correlation and position scores of all the peaks in the segment. Based on the global score, the algorithm decides which of the initial candidates to retain as valid heartbeat and keeps processing the signal further.
Note: The algorithm is purposely tuned to clean sections where peaks can be safely identified as heartbeats and ignore the rest so that it can be processed manually.
Manual post-processing.
The heart rate curve is inspected for suspicious patterns. In cases where the algorithm struggles to distinguish between noise and actual heartbeats, but the experimenter can confidently identify true heartbeats, it is safe to discard the noise peaks and retain the authentic heartbeats. However, if the experimenter has any doubt, we recommend excluding the time period starting from (and including) the last clearly identified valid heartbeat up to (and including) the first subsequent valid heartbeat.
Choose a zoom magnitude that allows to see anomalies in the heart rate curve (typically approximately 200 s). Browse the recording with the slider line on top or by pressing left/right arrows on the keyboard.
Check for anomalies (Figure 11).
i. A typical example is when a noise peak is taken instead of a very close heartbeat; because the noise peak is too early, it creates a sharp and very transient rise in heart rate, and when the sliding window used for computing the heart rate goes past it, it then creates an equally sharp dip. This biphasic noise is probably the most common and easiest to identify.
ii. Most of the time, when there are sections that the algorithm could not successfully clean, the manual processing consists in unclicking the noise peaks and/or setting the whole section to be excluded. Excluded ranges can then be set as NaN values for further processing. To exclude a range, click on Add exclusion range and draw a rectangle on the ECG plot. Overlapping ranges are automatically merged, and ranges can also be deleted by clicking on them and then clicking Delete selected range.
Notes:
1). Except when dealing with occasional recordings of suboptimal quality (poor signal-to-noise, artifacts, etc.), manual post-processing is extremely fast. It takes only a few recordings to get familiarized with heart rate curve and how they typically look and efficiently spotting problematic ranges with a quick glance.
2). It is better to use a small window for heart rate processing at this step, as it does not smooth out the anomalies as a bigger window would. 0.6 s is usually enough to not create gaps in the heart rate and allows for good identification of problematic ranges.
3). When manually ticking/unticking several peaks at once, it is better to untick AutoUpdate while (de)selecting and then tick it back afterwards. This saves time by preventing the GUI from refreshing after each operation.
4). Excluded ranges are displayed as filled grey areas. The empty periods following then, and ending with a dashed line, represent the periods for which heart rate cannot be processed because of the excluded section, but are not, per se, excluded ranges.
Critical: It is highly advisable to pre-process each recording perfectly as it ensures that they can be used safely for any kind of subsequent analyses, without compromising the results by adding noise or biases to the quantifications.
Figure 11. Check for anomalies. Example of a noisy recording snippet. Noise peaks are easily visible by sharp, transient rise and dips on the heart rate curve (top). Unselect the falsely picked peaks or exclude ranges by clicking Add exclusion range.
Saving.
The GUI exports two files:
A “_HeartBeats.mat” file, containing heartbeat timestamps and exclusion ranges.
An “_ECGLog.mat” file, containing the analysis parameters and information regarding the experimenters and the date.
Note: An option can be enabled within the function to export the heartbeats and exclusion ranges as a .csv file.
Obtaining the readout from the raw heartbeats
The same method as the one used by the GUI can be implemented to derive heart rate from heartbeats.
Choose a fixed window size, like 0.6 s.
For each heartbeat, count the number of heartbeats that occurred in that 0.6 s window before the current beat.
Divide this count by the time difference between the first beat in the window and the current beat.
Parse the exclusion ranges and set all the values that fall in the corresponding time windows (plus the size of the averaging window) to NaN. This allows to average between mice/periods without having artifacts from the bad ranges.
(Optional) This method offers a quick and efficient way to derive heart rate from heartbeats. However, it results in irregularly spaced data points because there is one data point for each heartbeat. To make it suitable for averages or when creating peristimulus time histograms, it is essential to resample the data to a fixed sampling rate so that the data is evenly spaced.
Note: When using this method to process heart rate from heartbeats, it is important to understand that the window ends precisely at each beat. This means that the heart rate curve shows immediate increases in heart rate with each beat. However, it is crucial to recognize that the decrease in heart rate is more gradual. Different alignment options could be chosen, such as centering the window on each beat or starting it at each beat, among others, depending on your technical preferences. Regardless of the chosen alignment, it is essential to keep this in mind when interpreting the data, as it can affect the perception of heart rate changes.
Use the GetHeartRate.m function to replicate this step.
Summary of analyses/quantifications that can be performed
Global averages.
Why: To globally compare a particular readout between general conditions (e.g., species, context).
How: Perform a long recording, pre-process the data, and obtain one average per individual.
Statistics: Depends on the characteristics of what is compared (Figure 12).
If the values come from a single group from measures repeated at different time points (for instance during the development of a pathology) or under different conditions (sated vs. fed): repeated measures one-way test + post-hoc tests.
If the values come from a single group but different cohorts (for instance different drugs are tested): a one-way test + post-hoc tests.
If the values come from several groups (for instance wild-type vs. mice with specific knocked-out genes): a one-way test + post-hoc tests.
If the values come from several groups recorded at different time points (for instance comparing the development of a model at different weeks between mice that received the treatment vs. control mice): a repeated measures two-way test + post-hoc tests.
If the values come from several groups receiving different treatments, with one treatment per cohort: a two-way test + post-hoc tests.
Interpretations: Depends on the test.
Note: For all these cases, one important aspect is to properly inform the statistical software about the matched conditions. The exact choice of the test depends on the number of groups and whether certain conditions are fulfilled (e.g., normality, homoscedasticity), which falls beyond the scope of the current protocol. Same goes for the interpretation: considering main effects/interactions when available and when to interpret them, and post-hoc comparisons.
Figure 12. Typical data holder table. (i) n groups could be recorded under t conditions or time points each (the whole table, grey contour), allowing the use of two-way tests. (ii) In case n groups were recorded for a single condition (yellow contour), or (iii) if a single group was recorded under t conditions, a one-way test would be required. Both cases (i) and (iii) would require taking into account repetitions if the same individuals were undergoing the different conditions.
Average whole traces.
Why: To observe the evolution of a signal during a recording, potentially between two groups/conditions.
How: Perform a long recording, pre-process the data, and obtain average curve(s) from matching conditions.
Statistics: If the statistical analyses are to match the curves (which could still only be showed as example/descriptive data), time needs to be one of the dimensions, which de facto changes all statistics to repeated measures tests. Post-hoc tests could then inform about specific time periods that are significantly different (within groups or across). Same principles as above apply—but this time with potentially more factors, which can lead to three-way analyses and/or sub-selecting the data by conditions and operating statistical tests on meaningful subsets of factors. Alternatively, group-matched fitting and associated statistical comparison could be used.
Peri-stimulus time histograms (PSTH).
Why: To observe changes happening around a synchronizing event (stimulus, behavior, etc.), potentially between two groups/conditions.
How: Perform a recording, pre-process the data by aligning it around the events, and obtain average curve(s) from matching conditions.
Note: Data can be normalized, typically to a baseline, by subtracting the mean and, optionally, dividing by the standard deviation (resulting in a z-score from the baseline). However, it is crucial to consider whether the baseline itself exhibits intrinsic differences, as this could either obscure or introduce an artificial effect during the normalized period (see section D for more details and examples).
Quantifications/statistics:
One way is to treat PSTHs like the average whole traces and perform RM analyses.
Another is to compare before/after periods directly with paired tests.
Finally, more tailored analyses could be used when they make sense. For example, a single peak or trough value can be extracted in a specific time window and group comparisons can be conducted on these individual per-mouse values.
Note: When constructing PSTH, particularly when presenting data dispersion on the curve (using either standard error or standard error of the mean) and choosing/reporting statistics, the precise method of averaging and counting is decisive and should be transparently documented.
Pooling all events from all animals is justifiable in specific cases but inherently reduces the apparent data dispersion while increasing statistical significance. Furthermore, the number of events per mouse can vary based on experimental designs or quantification methods, especially when the synchronizing event involves behavior. Consequently, relying solely on a global average can skew the results towards specific mice within the study. It is advisable to obtain an average per experimental unit (one recording and/or one animal) to use as a value for the PSTH and statistics. The range of total number of events could still be mentioned where appropriate as average number ± deviation.
Interpreting the data
Direct interpretation of the analyses presented above should not pose any significant challenge beyond typical statistical results interpretation. However, the method of choice and data sub-selection require careful consideration.
For studies based on event-induced changes, it is tempting to focus exclusively on PSTH analyses; conversely, for studies comparing groups, it seems straightforward to compare average heart rate from group A with average heart rate from group B, and so on. However, before performing any quantification or further analysis, it is often useful to take a global look at heart rate curves. To illustrate the importance and benefits of such approach, we provide below a few examples of key principles and potential confounds or insights that might be overlooked during a blind analysis (examples in Figure 13, summarized in Table 1) before discussing their underpinnings and then generalizing and giving general directions on how to explore new datasets.
Note: These examples primarily revolve around heart rate changes associated with behavioral immobility, as it is the central focus of the study that generated the data set from which they were selected. Nonetheless, the underlying principles and key takeaways should be relevant and applicable to a wide range of conditions.
Figure 13. State-dependency of heart rate values. A. Representative heart rate trace for a single mouse over the course of a second fear-conditioning day. Black lines on top depict immobility bouts, vertical dashed lines represent the compound conditioned stimulus (CS), and vertical dark lines the US presentations. (1) At first, it could be interpreted that heart rate variability (HRV) is increasing over time, but (2) heart rate fluctuations are tightly correlated to immobility behavior. (3) There is a slow rise in heart rate at the beginning of the recording that could be attributed to context-re-exposure and therefore fear. Additionally, while similar values can be reached during e.g., (1) as during the early phase (3), they are unlikely to have the same mechanistic underpinnings. B. Average heart rate for n = 33 mice during conditioning day 2. (1) It looks as though there is a peak only during CS phase, and (2) baseline-like levels in between. While this undeniably captures an average phenomenon, it does not reflect single mouse curves (e.g., in A.), in which there seems to be a toggle between bounded high and low values. (3) Such theoretical maximum (ceiling) is represented onto the average curve. C. Average heart rate curve for n = 25 mice in an open-field recording. (1) A similar early increase as in conditioning day 2 [B. (3)] can be seen. This suggests that such increase is unrelated to conditioning itself. D. Detrending heart rate based on the theoretical maximum in B (3) allows for a more robust and faithful interpretation: (1) heart rate decreases between CSs and peaks during the second half of the CS and the shock. (2) This also leads the early rise to be normalized, which prevents wrongful interpretation about absolute heart rate values. It also homogenizes the values between mice as visible by the very low variability when compared to B (3).
Table 1. Examples of potential observations of heart rate characteristics and their direct and adjusted interpretations
Observation Direct interpretation Closer look Adjusted interpretation
A(1) HRV is high Fear conditioning increases HRV A(2) HRV here actually reflects immobility bouts that are correlated with HR decrease Fear conditioning conditions increase immobility bout probability
A(1) HRV is increasing over time Fear conditioning increases HRV over time The amplitude of immobility-related decreases in heart rate increases over the course of the recordings Some latent process over the course of the recordings globally affects the amplitude of heart rate changes (Rigidity). This is not specific to threat exposure, but threat exposure changes the magnitude of this phenomenon
A(3), B(3) HR increases at the beginning of the recording Re-exposure to the conditioning context increases HR C(1) HR increases at the beginning of the recordings in other contexts Something in the recording procedure leads to an early increase in HR
B(1) HR increases during CS/US pairings, but B(2) remains at a constant level in between CS/US increases HR D(1), A(1) HR decreases during immobility bouts that are frequent outside the CS/US pairings, and increase during the second half of the CS/the US HR changes reflect immobility probability changes as well as rigidity changes
These few points highlight several key principles:
State dependency
It is common, particularly in human studies, to discuss heart rate in terms of a constant value and compare general metrics across individuals with diverse ages, fitness levels, and health statuses. However, it is evident that heart rate is a highly dynamic parameter, influenced by a spectrum of biological processes and environmental elements, operating on different timescales. Circadian rhythms modulate HR, HR increases to match the body’s needs when exercising, and most importantly, emotions drastically affect HR. Hence, when considering these average values, it is often implied that they correspond to the resting heart rate obtained under equivalent and somewhat controlled circumstances. Similarly, in mouse studies, a single recording should yield a valid per-individual average as long as the conditions remain consistent between the values being compared. Nevertheless, for mice, interpretations can quickly become delicate, with the slightest variations in the conditions. While avoiding excessive determinism, the aim is to work within a framework that acknowledges the accessible factors, enabling a balanced interpretation and robust conclusions.
Behavior
Behavior is a critical determinant of heart rate values in mice. For instance, entering or leaving a defensive immobility bout, commonly termed freezing, leads to rapid and drastic changes in heart rate. Rearing causes a transient increase in heart rate, and many other behaviors are likely associated with heart rate fluctuations. Interestingly, the increase in heart rate associated with locomotion is often masked in many typical recording conditions (see Section B). In general, the most pronounced heart rate changes can be traced back to a behavioral change.
Generally, the most significant short-term heart rate changes can be attributed to behavioral shifts. Even within the same behavior, the cardiac response can depend on intrinsic properties, such as the duration of the behavioral bout (for example, immobility bout duration is positively correlated with the heart rate decrease amplitude).
• When averaging over extended periods, the resulting average heart rate may heavily depend on the proportion of certain behaviors (e.g., if the mouse displays many immobility bouts, the average will be low) and their respective characteristics, which can be influenced by different factors.
• Analytical power and meaningfulness of the results can be enhanced by comparing heart rate values during similar behaviors and even accounting for specific behavioral bout characteristics.
→ Investigate if individual behaviors are associated with consistent, time-locked heart rate changes (i.e., one behavioral bout leads to a stereotypical heart rate change).
→ Explore if there are time periods enriched in certain behaviors that exhibit overall higher or lower heart rates.
→ Incorporate these factors into the analyses by matching behaviors and periods accordingly.
Context
The diffuse threat level of the context, and consequently, the anxiety it generates, affects average heart rate values (even for matched behaviors; Signoret-Genest et al., 2023). This is true both for a paradigm as a whole (e.g., open field vs. small arena) as well as within specific areas of certain paradigms (e.g., closed arms vs. open arms in an elevated plus maze). This perception of diffuse threat is shaped by inherent species-specific evolutionary responses (e.g., open spaces being associated with an increased risk of predation) and may be influenced by other factors such as prior history or individual traits.
Surprisingly, the level of contextual threat also influences the magnitude of heart rate changes associated with the same behavior: bradycardia associated with immobility is reduced in higher-threat environments.
Notably, the baseline threat level/stimulus induced by standard recording procedures can be sufficient to obscure any increase in heart rate associated with locomotion, as the baseline is already elevated (Andreev-Andrievskiy et al., 2014).
In extreme cases, presenting the same stimulus in different contexts can elicit opposing heart rate responses (see also e.g., Signoret-Genest et al., 2023).
• If two different recordings are performed to compare two conditions (e.g., two drugs) but in two areas that are too different, it could reflect an influence of the area instead of that of the treatment.
• When averaging across a session recorded in a paradigm with areas associated with different threat levels, the time spent in each area will influence the resulting average.
• Again, accounting for the potential impact of context can increase statistical power by controlling for the variability it otherwise induces, which also constitutes a meaningful result in itself.
→ Take these considerations into account for the analyses (ensure context/areas are matched, assess context/areas' independent influence).
Heart rate modulation at different time scales
Changes in heart rate related to behavior operate on the timescale of a behavioral episode—ranging from fractions of a second to a few dozen seconds. Contextual effects may extend over an entire recording or specific periods within a recording when subareas present differences. Additionally, slower processes can manifest, as demonstrated in the provided example (an increase in heart rate due to the recording procedure and a gradual augmentation in the amplitude of cardiac responses).
→ Again, it is advisable to account for these factors in the analyses, seeking out patterns and correlations across different scales. Decomposing the signal in such a way may allow for the mitigation of these factors’ influence on the data as well as their comprehensive characterization.
History/individual characteristics
Exposure of mice to fear conditioning, for example, is likely to lead to subsequent heightened levels of anxiety. Beyond the observable behavioral shifts and associated direct heart rate changes, one would anticipate consequential alterations in heart rate patterns. Metabolic challenges at earlier time points may also have a lasting impact on cardiac responses, and various other biologically significant states can reasonably be expected to induce or modulate changes in heart rate (e.g., satiated vs. fed states). Taking these factors into account in specific research areas is crucial as they have the potential to significantly influence outcomes.
→ In some cases, conducting longitudinal analyses and recognizing the individuality of mice entering a session (beyond the obvious treatment differences) can provide valuable insights by introducing a level of complexity that helps elucidate variability.
Preexisting state
Baselines are also a non-negligible source of variability, often hidden in the fine methods details. Leaving mice to habituate before starting the recording/procedure or not can lead to major differences—up to the point of inverting the heart rate responses. But even at the level of smaller scale events, such as an individual behavioral bout or the presentation of a stimulus, the pre-state of the mouse is a critical determinant of the following cardiac changes. In some cases, the relevant toggle for heart rate is entering or exiting a specific behavior, in which case transitioning from or to another one, respectively, does not matter.
→ It is sometimes worth thinking of transition from one state to another rather than simply entering/exiting one state (e.g., a behavior).
→ A corollary to this is that normalization of the signal to baseline is a double-edged sword since the baseline might not be homogenous between episodes and/or conditions and present intrinsic meanings on its own. For example, if a behavior is correlated to a decrease in heart rate globally, but one episode occurs during a state where the heart rate is already low, it might not decrease any further. Normalizing by the baseline would look as though there is no change, when the most relevant information is that the heart rate is (still) low.
Interaction
Different aspects could interact with each other, yielding potentially paradoxical results if not disentangled. For instance, if a certain heart rate change occurs at a very specific time point, and if mice happen to be probabilistically in a specific area around the same period, the area-effect could modulate the otherwise non-causally related heart rate change.
→ As a last step, the different elements should be pieced together.
Umbrella terms and semantics
Trying to fit an observation “into a box” too early on can be problematic, as it leads to skewed data exploration and interpretations that are biased by higher level concepts and semantics and taken away from the low-level data description.
A prime example is the HRV, a widely used metric for studying human heart rate, making it a potentially valuable tool for mouse studies with translation in mind. However, it is worth noting that HRV encompasses a wide range of analyses. While descriptive and comparative results can be useful for diagnostics and revealing hidden states, the field faces a challenge in terms of lacking a solid mechanistic grounding and understanding. This can make biological interpretations somewhat precarious, especially as the increasing popularity of HRV can sometimes lead to overreaching interpretations that are not grounded in facts.
Most HRV analyses rely on long recordings, for which they produce a single value per readout. So-called time domain analyses look directly into beats intervals. The Root Mean Sum of Squared Successive Differences (RMSSD) is one of the most commonly used (Shaffer et al., 2014). However, because these are so closely related to beats intervals, they are inherently strongly correlated to changes in heart rate at a small-time scale. For that reason, even larger time scales, particularly in mice, should be controlled for the points mentioned in step D1.
Frequency-based analyses of HRV look at the representation of certain frequency bands in the heart rate signal (that is, how much it oscillates at certain rhythms). While they provide sometimes useful biomarkers, their biological underpinnings are unclear and there is no clear consensus. We recently applied frequency analyses to our signals to quantify a macroscopic, visible oscillation: the equivalent of the Mayer-Waves in humans, which are the oscillations hypothesized to originate from baroreflex loops. Because their amplitude was affected in a similar manner as some of the other, simpler, heart rate changes (increases/decreases), it provided us with a bidirectional validation of our hypothesis—that a latent mechanism related to a baroreflex curve tuning was constraining heart rate changes.
A last type of HRV analyses exists: the non-linear analyses, which suggest that heart rate is inherently fractal (it repeats patterns at different scales). While showing promises for diagnosis, it suffers from a lack of biological interpretability.
→ We would advise to use HRV analyses either with translatability to humans in mind, or to characterize and quantify a very obvious oscillatory pattern in heart rate that cannot be explained by other means, as many changes could otherwise be inadvertently confused for HRV.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Signoret-Genest et al. (2023). Integrated cardio-behavioral responses to threat define defensive states. Nature Neuroscience (all figures).
We are providing a small data set along with the current protocol, allowing to access examples of raw ECG data as well as the corresponding pre-processing and analysis procedures. Additionally, we provide open-access code that can be applied to any steps for testing and experimentation.
General notes and troubleshooting
General notes
We chose to use 6-pin Omnetics connectors because of their minimal size and weight, which allows to combine ECG recordings with techniques requiring additional elements to be fixed onto the skull, and because we might occasionally require additional channels. These connectors are of high-end quality and provide perfect connections but can increase the per-mouse costs, even though they are reusable with proper care (see section C). Simpler three-pin connectors might be used instead; the general procedure and important tips remain the same.
Manufacturing the patch cable may initially pose challenges due to the fragility of the wire. In terms of selecting the cable model, the chosen reference is relatively costly but offers two significant advantages: a) it is exceptionally thin and lightweight, and b) it is shielded, making it suitable for recording in conditions where noise cannot be avoided (or simply dealing with usual ambient noise). On the other hand, this comes at the cost of being pretty delicate. Considering these points, the decision could be made to switch for a cheaper/sturdier reference if it seems more beneficial.
The ECG implantation described here is the least invasive possible in terms of number of implanted electrodes and providing the most stable recordings over several weeks. It allows for accurate and robust heartbeat detection and therefore reliable heart rate extraction. However, with the setup and parts proposed here, it can effortlessly be expanded to four channels for applications that might need more in-depth analyses of the heartbeats themselves (waves). Since the cable is already made so that it can feed the preamplifier signals from more than two channels, it only requires to solder two more wires to the connectors and implant the four resulting electrodes at the desired place on the mouse. It is then probably best to record only the direct output from each channel and take care of the specific ECG derivations offline.
Troubleshooting
Problem 1: The ECG signal is not visible
Possible cause #1: There is too much noise to see the ECG.
Solution #1: See Problem 2.
Possible cause #2: Something is wrong with the amplifier parameters or the cabling, etc.
Solution #2: See section D from Procedure.
Possible cause #3: Something is wrong with the implantation.
Solution #3: Assess the surgery quality, if needed post-mortem, to understand what could have gone wrong.
Possible cause #4: Something is wrong with the connector preparation or the cable integrity.
Solution #4: Check the cable for shortcuts or lack of connection and check any new connector before implanting (making sure the soldering is of good quality and that each pin connects to the end of its corresponding wire).
Problem 2: The signal is noisy
Possible cause #1: The signal and recording system are fine but there is no ECG, so the baseline looks as though we have only noise.
Solution #1: Troubleshoot for no ECG (implantation issue, cable/connector issue).
Possible cause #2: Something is wrong with the amplifier parameters.
Solution #2: See section D from Procedure.
Possible cause #3: The system is picking noise, in particular the ambient “hum” (noise coming from AC-powered equipment). The shielded cable decreases the impact of that common issue with electrophysiological recordings but might not prevent it entirely. This is very likely to happen for a new setup. Check for 50 Hz (or 60 Hz depending on the power-grid characteristics) and harmonics in the spectrogram of a recording.
Solution #3: The system will need to be grounded in such a way that the noise is gone or at least decreased. There are general principles as to how to ground a system but no unique solution. In case nothing works, it might be best to contact the support from the amplifier company (NPI in the case of this setup).
Problem 3: The signal looks fine sometimes, but some sharp artifacts render the rest of the recordings unusable
First, assess whether it occurs for a single individual or all of them.
If it happens for a single individual:
Possible cause #1: One or several wires are not properly fixed (anymore). This is likely to be due to a problem during the surgery. The wires could have been slightly too short, leading to some tension, ultimately leading them to move away, or the stitches on the muscles were suboptimal, or the ball of glue at the tip was not big enough/stable enough, or a mix of several factors.
Solution #1: Once the technique is implemented and the experimenter is experienced enough, these particular issues should become rare. In most cases, it seems better to address the problematic elements upstream (e.g., having wires long enough, properly stitched), than to resort to a second surgery to try and fix things, which might not even be possible.
Possible cause #2: Similar to cause #1, but a single wire is affected.
Solution #2: Change the settings to disable differential recordings and check each wire’s individual signal. If the environment is relatively noise-free, differential recording is not mandatory and it could be that the differential signal looks bad because one wire has an issue, while the single channel’s signal from the other would look fine.
Possible cause #3: The electrode picks a lot of muscular signals (electromyogram, EMG), which typically looks like bursts on the recording. They are present to varying degree in some animals, but it becomes problematic when their amplitude is greater than that of the ECG.
Solution #3: That particular animal was implanted at a suboptimal location and/or the electrode was stripped too much or not enough; it cannot be solved but can serve as feedback for later implantations.
If it happens for all individuals:
Possible cause #1: The head-to-headstage cable is touching something.
Solution #1: Make sure the cable is hanging from the middle of the arena as much as possible and not touching anything else than the thread holding it up via the pulley.
Possible cause #2: The cable is damaged. The cable used is ideal because it is shielded and light weight, but it is also fragile. A moment of inattention leading to a mouse grabbing and biting the cable can lead to some bite marks that can produce various effects depending on what is damaged:
- Breaching the shielding can let some environmental noise through.
- Breaching the individual wires’ insulation can lead them to sometimes contact other wires or the shielding sheath, creating artifacts as motion is applied to the cable.
- If a single wire is partially severed, it could intermittently disconnect.
Note that bending the cable too hard or repeatedly on the same place can also lead to damage that is not visible, with similar consequences.
Solution #2: Check for bite marks along the cable and assess how bad the damage is. In case of no external damage, carefully palpate the cable on all its length to locate potential breaks inside the insulation.
If there is no time to make another cable (an experiment is running and cannot be postponed), using tape to tightly wrap the cable can hold the wires in place for some time, which provides a temporary solution. Regardless, the cable needs to be changed; it is wise to always have a second cable ready just in case.
And as a more general fix: the setup could maybe be improved so that the cable cannot be easily grabbed (enough tension to pull it when needed and keeping it from hanging too low) and making sure the experimenter keeps a close eye on what is happening, so that he can intervene whenever required.
Possible cause #3: There are some rather slow and somewhat cyclic drifts at play, but the filtering is set in such a way that it looks like artifacts during some turning points.
Solution #3: Check Problem 3.
Possible cause #4: The electrode picks a lot of muscular signals (electromyogram, EMG), which typically looks like bursts on the recording (Figure 14). If this happens more or less systematically and not anecdotally and is not restricted to bursts of very small amplitude (in theory, if the amplitude is smaller than the heartbeat’s signal, it should be workable but still denotes a systematic issue), two things could be checked:
Solution #4a: The implantation sites are not optimal and must be readjusted.
Solution #4b: The electrodes’ material might not have ideal properties. Try to use the reference provided here, or at least, change from what is currently used.
Figure 14. Electrocardiogram (ECG) contamination by electromyogram (EMG) signal. Representative raw ECG trace (only filtered online by the amplifier), showing heartbeat signal and EMG contamination (looking like thicker baseline noise, e.g., prominent at t = 218 s). In this case, the amplitude is smaller than the ECG signal, but in some recordings the EMG signal could prevent ECG analysis.
Problem 4: The signal looks fine sometimes, but some slow oscillations are superimposed
Possible cause: The recordings are combined with another device and the shielding is either damaged or insufficient.
We encountered such issues in two different cases, which can help understand others:
The ECG cable is bundled with a data transfer cable for a different type of recordings.
In our case, we bundled the ECG cable with the cable for a miniaturized head-mounted microscope (miniscope). That microscope relies on a wet-lens to achieve focus on the region of interest and can quickly shift between several depths of focus. This requires reshaping the wet lens with different currents that cause specific offset when the focus is held.
Solution #1: We implemented the current shielded cable, which works fine into preventing all noise contamination. If the contamination comes back, that is a sign that the shielding is damaged.
The animal is placed in an electromagnetic field.
For some other experiments, we needed to deliver electromagnetic stimulations. The stimulations were ranging between 10 and 50Hz, 50% duty cycle. At each pulse, a biphasic deflection could be visible on the ECG (a rise then a decrease).
Solution #1: The shielded cable was not enough to prevent noise contamination, as the entire subject was bombarded with it. We were however able to record the current pulses driving the stimulation to retrieve their timing. This allowed us to determine and fit a model for the noise by looking at the signal at each onset/offset of the stimulation. Fitting the model to each stimulation then allowed to subtract the estimated noise and retrieve an almost clean ECG signal that could then be pre-processed as usual. This can in theory be extended to different types of noise contamination that follow a discernable pattern. Alternatively, under certain conditions, one could implant the reference wire closer to the other electrodes, so that the noise would be more similar and therefore just subtracted.
Problem 5: The head cap falls off
Possible cause #1: The surface of the cap on the skull is not enough.
Solution #1: Remove a bigger patch of skin to build a larger head cap.
Possible cause #2: The glue was applied on a wet skull.
Solution #2: Dry the bone before adding the glue. Some minor bleeding can sometimes occur from the bone itself; do not use bone wax as it would decrease glue’s adhesion but gently scrap the bleeding area with a blunt tool instead (consider using an appropriate rugine).
Possible cause #3: The surface of the skull was not scratched enough.
Solution #3: On the dry bone, do not hesitate to draw a very clear grid pattern with the back of a scalpel blade. This is critical for the head cap to be firmly fixed.
Acknowledgments
We thank Dr. Giorgio Rizzi for discussions and valuable suggestions for the cable reference when we first encountered cross-contamination of our ECG signal, and Jens Looser from NPI Electronics for initial help and advice.
We are grateful to J. Deckert (Center for Mental Health, University Hospital Würzburg) for his support on this project (to J.S.G.). This work was supported by the Deutsche Forschungsgemeinschaft [Heisenberg professorship and project funds to P.T.: (TO 1124/1,2,3), TRR 295: (446022135), (446270539)] and received funding from the European Union’s Horizon 2020 research and innovation program under the Marie Skłodowska-Curie grant agreement No. 956414 (to P.T.). It was supported by a NARSAD Young Investigator Grant of the Brain and Behavior Foundation to P.T.
Competing interests
The authors declare no competing interests.
Ethical considerations
All animal procedures were approved by the local veterinary authorities and animal experimentation ethics committee (Regierung von Unterfranken, authorization 2532-2-509). ARRIVE guidelines were followed (Animal Research: Reporting of In Vivo Experiments).
References
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Calvet, C. and Seebeck, P. (2023). What to consider for ECG in mice—with special emphasis on telemetry. Mamm. Genome 34(2): 166–179. doi: 10.1007/s00335-023-09977-0
Carrive, P. (2000). Conditioned fear to environmental context: cardiovascular and behavioral components in the rat. Brain Res. 858(2): 440–445. doi: 10.1016/s0006-8993(00)02029-1
Chu, V., Otero, J. M., Lopez, O., Morgan, J. P., Amende, I. and Hampton, T. G. (2001). Method for non-invasively recording electrocardiograms in conscious mice. BMC Physiol. 1: 6. doi: 10.1186/1472-6793-1-6
Fares, R., Flénet, T., Vial, J., Ravaz, M., Roger, V., Bory, C. and Baudet, S. (2022). Non invasive jacketed telemetry in socially-housed rats for a combined assessment of respiratory system, electrocardiogram and activity using the DECRO system. J. Pharmacol. Toxicol. Methods 117: 107195. doi: 10.1016/j.vascn.2022.107195
Leman, S., Dielenberg, R. and Carrive, P. (2003). Effect of dorsal periaqueductal gray lesion on cardiovascular and behavioural responses to contextual conditioned fear in rats. Behav. Brain Res. 143(2): 169–176. doi: 10.1016/s0166-4328(03)00033-0
Sato, S. (2019). Multi-dry-electrode plate sensor for non-invasive electrocardiogram and heart rate monitoring for the assessment of drug responses in freely behaving mice. J. Pharmacol. Toxicol. Methods 97: 29–35. doi: 10.1016/j.vascn.2019.02.009
Shaffer, F., McCraty, R. and Zerr, C. L. (2014). A healthy heart is not a metronome: an integrative review of the heart's anatomy and heart rate variability. Front. Psychol. 5: e01040. doi: 10.3389/fpsyg.2014.01040
Signoret-Genest, J., Schukraft, N., L. Reis, S., Segebarth, D., Deisseroth, K. and Tovote, P. (2023). Integrated cardio-behavioral responses to threat define defensive states. Nat. Neurosci. 26(3): 447–457. doi: 10.1038/s41593-022-01252-w
Stiedl, O., Jansen, R. F., Pieneman, A. W., Ögren, S. O. and Meyer, M. (2009). Assessing aversive emotional states through the heart in mice: Implications for cardiovascular dysregulation in affective disorders. Neuroscience & Biobehavioral Reviews 33(2): 181–190. doi: 10.1016/j.neubiorev.2008.08.015
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Seed Collection in Temperate Trees—Clean, Fast, and Effective Extraction of Populus Seeds for Laboratory Use and Long-term Storage
NB Naima Bhutta
ON Oscar F. Nunez-Martinez
CM Carmen Mei
KB Katharina Bräutigam
Published: Vol 14, Iss 3, Feb 5, 2024
DOI: 10.21769/BioProtoc.4927 Views: 661
Reviewed by: Samik BhattacharyaSimab KanwalTie Liu
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Abstract
Seeds ensure the growth of a new generation of plants and are thus central to maintaining plant populations and ecosystem processes. Nevertheless, much remains to be learned about seed biology and responses of germinated seedlings to environmental challenges. Experiments aiming to close these knowledge gaps critically depend on the availability of healthy, viable seeds. Here, we report a protocol for the collection of seeds from plants in the genus Populus. This genus comprises trees with a wide distribution in temperate forests and with economic relevance, used as scientific models for perennial plants. As seed characteristics can vary drastically between taxonomic groups, protocols need to be tailored carefully. Our protocol takes the delicate nature of Populus seeds into account. It uses P. deltoides as an example and provides a template to optimize bulk seed extraction for other Populus species and plants with similar seed characteristics. The protocol is designed to only use items available in most labs and households and that can be sterilized easily. The unique characteristics of this protocol allow for the fast and effective extraction of high-quality seeds. Here, we report on seed collection, extraction, cleaning, storage, and viability tests. Moreover, extracted seeds are well suited for tissue culture and experiments under sterile conditions. Seed material obtained with this protocol can be used to further our understanding of tree seed biology, seedling performance under climate change, or diversity of forest genetic resources.
Key features
• Populus species produce seeds that are small, delicate, non-dormant, with plenty of seed hair. Collection of seed material needs to be timed properly.
• Processing, seed extraction, seed cleaning, and storage using simple, sterilizable laboratory and household items only. Obtained seeds are pure, high quality, close to 100% viability.
• Seeds work well in tissue culture and in experiments under sterile conditions.
• Extractability, speed, and seed germination were studied and confirmed for Populus deltoides as an example.
• Can also serve as template for bulk seed collection from other Populus species and plant groups that produce delicate seeds (with no or little modifications).
Graphical overview
Keywords: Temperate trees Populus Seed collection Seed extraction Seed storage Viability Laboratory use Tissue culture
Background
Seeds are an innovation of Spermatophyta (seed plants). They are specialized dispersal units that safely package the plant embryo to allow for dispersal in space and time. Seeds are critical for successful propagation of the individual and maintenance of populations within a species but also for terrestrial ecosystems including forest ecosystems. Seeds can vary drastically in their characteristics, such as size, mass, color, or durability [1]. The smallest orchid seeds, for example, weigh less than 1 μg, while the large seeds of the double coconut palm (Lodoicea maldivica) can reach up to 25 kg in weight [2–4]. Poplar seeds lose viability within a few days or weeks after release, while seeds of a date palm from an archeological site were reported to have germinated after 2,000 years [5,6]. This means that seed collection, handling, processing, and storage need to be tailored carefully to the species or taxonomic group of interest to ensure seed viability and successful plant growth from the seed. Here, we focus on the seeds of temperate forest trees of the genus Populus.
Despite their importance for healthy forests, much remains to be learned about seed traits and biology in forest trees, ranging from the molecular mechanisms of seed production or seed performance under climate change to applications in forest genetic resources preservation and long-term forest management [7–11].
Trees in the genus Populus are among the most widespread trees in North America, including some of the largest and fastest growing hardwoods in this large geographic area [5,12,13]. This taxonomically complex genus comprises poplars, cottonwoods, and aspens. They are early successional in natural plant communities but are also used widely in intensive culture for biomass, pulp, and paper production, for lumber, as shelterbelts or urban trees, or in land reclamation [5,13]. Populus species are dioecious, reach sexual maturity at 10–15 years of age, and produce seeds every year once matured. The number of seeds produced is remarkable: 28–54 million seeds have been reported for individual trees of Eastern cottonwood (P. deltoides) and European aspen (P. tremula) [5,14,15]. The seeds themselves are small and covered with a significant amount of hair (cotton or white fluff) that allows for easy wind dispersal. Seeds do not show dormancy, germinate quickly under suitable conditions, and can lose viability rapidly under natural conditions [5,16]. These characteristics impose challenges on seed collection, cleaning, and storage. Seed collection, therefore, needs to be timed precisely with the seasons and seed ripening on the trees. Moreover, extraction of seeds from the difficult-to-separate seed hair requires careful handling to prevent damage to the thin seed coat. Finally, seeds need to be stored properly to ensure longer lasting viability.
Agroforestry equipment such as seed macerators or grinders and fanning mills can be used to extract large numbers of Populus seeds [5,17]; however, seed viability declines strongly within two days, possibly due to damage of the delicate seed coat. Similarly, brushing against a coarse screen released seeds from the white cotton, but the extraction efficiency was low (20%), and seeds remained viable only for a few days [18]. Smaller quantities can be collected using a vacuum cleaner, and separation from cotton can be done in a nested set of soil sieves and applying a stream of air [19–21]. Based on this rationale, we established a method for seed extraction that leaves the thin seed coat intact and that yields very clean seeds, suitable for molecular analyses. The method is fast, simple, and effective. It uses household items that can be obtained easily, that are used in every lab, and importantly, that can be cleaned and sterilized easily. Storage of the extracted seeds was tested under different conditions. Germination tests confirmed high viability of seeds directly after extraction as well as after extended storage time. Growth in tissue culture without contamination furthermore confirmed purity of seeds.
Due to the high quality, intactness, and purity of seeds extracted with this protocol, these lend themselves well to analyses of seed responses under controlled conditions in tissue culture, also germinating well on soil and other substrates. The protocol was specifically developed for Populus to separate seeds from the attached tufts of hair but could equally be adjusted for the extraction of small, delicate seeds in other species that need separation from attached structures or impurities.
Materials and reagents
Biological materials
Female trees from the genus Populus that carry catkins with fruits that are near maturity (see Figure 1). Nearly mature catkins develop between late spring and earlier summer in temperate climates. The precise timing of seed maturation will depend on geographic location and local weather conditions.
The reproductive material used in this protocol was collected from Populus deltoides Bartr. ex Marsh. trees growing in Ontario, Canada in the Greater Toronto Area (GTA) and at the University of Toronto Mississauga campus. Details on location of the trees and on seed collection are given below in Table 1.
Table 1. Trees and material used in this study
Genotype (tree) Species Latitude Longitude Elevation (m) Collection date
POP31 P. deltoides 43.6311319 -79.4715803 75 2023
POP32 P. deltoides 43.6310720 -79.4713711 75 2023
POP50 P. deltoides 43.6294418 -79.474405 70 2023
POP50 P. deltoides 43.6294418 -79.474405 70 2022
POP29 P. deltoides 43.5464906 -79.6599571 110 2021
POP19 P. deltoides 43.5461274 -79.6605435 114 2023
Reagents
Murashige & Skoog (MS) basal salt mixture (PhytoTech Labs, catalog number: M524)
Plant preservative mixture (PPM) (Plant Cell Technology, catalog number: PPM)
Agar (BioShop, catalog number: AGR003)
Potassium hydroxide (KOH) (BioShop, catalog number: PHY202)
Solutions
MS medium (100 mL) (see Recipes)
Recipes
MS medium (100 mL)
Adjust pH to 5.8 using 2.5 M KOH and autoclave. Cool medium to approximately 60 °C and add 200 μL of PPM (0.2% final concentration) immediately before pouring plates. The given volume yields four plates.
Reagent Final concentration Quantity or Volume
MS 0.5× 0.2166 g
Agar 0.7% (w/v) 0.7 g
H2O n/a to 100 mL
PPM 0.2% (v/v) 200 μL
Total n/a 100 mL
Laboratory supplies
Petri dish, round, 100 mm × 15 mm (Fisherbrand, catalog number: 2071-FB0875712)
Petri dish, square, 100 mm × 15 mm (Fisherbrand, catalog number: 08-757-11A)
Borosilicate glass vial (VWR, catalog number: 66011-085)
50 mL conical centrifuge tube (FroggaBio, catalog number: TB50-500)
Splinter forceps (Almedic, catalog number: 7737-A10-600)
Micropore tape (3M Micropore, catalog number: 1533-0)
Household items
Paper lunch bags (1.88 L) (Canadian Tire, catalog number: 053-0200-8)
Plant propagation tray with lid (The Grow Depot, model: Mondi Propagation Tray 10 × 20)
Wire mesh pencil cup (10 cm × 8 cm × 8 cm); pencil cups can be obtained at any office supply store (e.g., Grant & Toy, Dollarstore)
Round bamboo skewer, 5 mm in diameter and cut into 2–2.5 cm pieces. Round bamboo sticks can be obtained at any retail or grocery store
Stainless steel fine mesh kitchen strainer (mesh size 30 and 40)
White paper sheets
Equipment
Micro bead sterilizer (Fisherbrand, catalog number: 14-955-342)
Laminar flow hood (Thermo Scientific, model: Heraguard ECO)
Growth chamber (Conviron, model: MTPS144)
Software and datasets
R Studio v 2023.03.0+386 [22]
R v4.3.x [23]
Procedure
Seed collection
Identify mature female trees and monitor reproductive development carefully.
Notes:
Trees in the genus Populus are dioecious. Copious amounts of seeds are produced on mature female trees (Figure 1A).
Flowers are arranged in flower clusters (catkins). After fertilization, catkins lengthen, and flowers develop into green fruit capsules carrying the developing and maturing seed (Figure 1B). Capsules split open when ripe and release seed with trichomes (cotton) for wind dispersal (Figures 1C and 1D).
Depending on geographic location and prevailing weather conditions, trees flower in spring, and seeds mature within 4–6 weeks [13,16].
Collect maturing catkins when the first few capsules begin to open (Figure 1C). Gently pick catkins from branches by breaking them off at the base. Store catkins in paper bags.
Notes:
If catkins are collected too early (Figure 1B), seeds will not mature properly, and the extracted seeds will not be viable. If catkins are left too long on the tree (Figure 1D), wind can disperse the whole seed crop in a very short period of time (e.g., an afternoon).
Therefore, close monitoring is required when catkins mature in late spring or early summer, ideally on a daily basis.
Figure 1. Tree habit and seed-bearing structures in Populus. A. Populus deltoides tree (eastern cottonwood) in late spring carrying female catkins (flower clusters). Each catkin comprises several capsules; each developed from an individual flower. B–D. P. deltoides catkins at different stages of capsule opening. B. Immature catkins with developing seeds fully enclosed in capsules. C. Catkin with first capsules split open upon gradual maturation, exposing trichomes and seeds for wind dispersal. D. All capsules on the catkin are fully open. They release copious amounts of seeds attached to tufts of long hair (cotton). Scale bars: 2 cm.
When catkin collection is completed, bring material indoors and spread out in trays. Evenly distribute catkins in a single layer and avoid overlap of catkins to allow for rapid drying. Keep in a dry place. Drying in the lab, a general storage area, or a climatized office at room temperature (20–24 °C) works well. No special drying room is required in temperate climates.
Note: Without drying, the material can become moldy and seeds will not mature properly.
Cover trays very loosely with a lid to keep emerging cotton inside the tray while allowing for proper ventilation.
Note: It will take between one and seven days for seeds to shed, i.e., for cotton to emerge.
Seed extraction should be done as soon as all capsules are open and cotton emerges.
Note: Tufts of long cellulose-rich hair are attached to the small, light seeds to enable wind dispersal. The hair needs to be separated from the seeds without damaging the delicate seed coat.
Yield: Harvesting and processing 15–25 catkins will yield approximately 10,000 clean seeds (see also Validation section 2 of this protocol).
Note: Seed production per catkin can vary between individual trees (Figure 3).
Seed extraction—mechanical extraction by shaking
Disinfect the surface of the work area and cover it completely with clean sheets of paper. This will help later with gathering the extracted seeds.
Assemble the seed shaker (Figures 2A and 2B) by placing 10–15 fresh pieces of bamboo sticks inside an autoclaved mesh pencil cup.
Notes:
The added bamboo stick pieces facilitate the mechanical seed extraction.
Various objects of different shape, density, and material have been tested for use in mechanical seed extraction. These included pieces of wood, small aquarium rocks, steel balls, or bamboo sticks. Moreover, bamboo pieces of variable length have been tested. Pieces of bamboo sticks 2–2.5 cm in length yielded best results.
Figure 2. Materials required for seed extraction, cleaning, and storage. A. Our protocol was designed to use exclusively (i) simple and (ii) easy-to-clean items that are (iii) readily available in every household and lab. B. Our assembled seed shaker. Cotton from dried and mature catkins (seeds embedded in hair) is placed inside a mesh pencil cup and covered with the lid of a Petri dish. Added bamboo stick fragments aid with the mechanical extraction. Shaking easily releases seeds, some of which are shown around the cup. C. Intact, clean seeds are stored in glass vials. Scale bars: 2 cm.
Transfer 0.5–1 g of cotton into the shaker and cover it with a Petri dish lid (Figure 2B).
Notes:
The cotton is the white fluff released from dry and mature catkins (Figure 1D, A.4), which comprises seeds and attached hair.
Avoid adding parts of the seed capsules or any other hard pieces from the catkin to the shaker.
Keep the fluff loose and avoid pressing it down. Compressed cotton will reduce the efficiency of seed extraction.
Adding more than 1 g of cotton will also reduce extraction efficiency and prolong the shaking and clean-up process.
Tilt the seed shaker (Figure 2B) to the side and shake vigorously by hand. Seeds will fall out of the shaker and onto the paper sheets. Seeds’ hair (white cotton with most seeds removed) will stay inside.
Continue shaking until only a few seeds are being extracted or until the trichomes inside the seed shaker form small, felt-like clumps.
Gather the extracted seeds from the paper sheets on the work surface, place them in a small container, and set them aside for subsequent cleaning steps.
Cover the work surface again with the paper sheets and remove the clumped trichomes from the mesh cup but leave bamboo sticks inside the pencil cup.
Repeat steps B3–B7 until all cotton from one genotype (tree) has been processed.
To avoid sample mix-up and cross-genotype contamination, process cotton from one genotype at a time. Sterilize mesh pencil cups before processing a new genotype, use a fresh set of bamboo stick pieces, and thoroughly clean the working area.
Seed cleaning
Place the extracted seeds into a fine mesh kitchen strainer and sieve the material in a gentle circular motion. Small debris will pass through the sieve.
Note: We tested several household items with different mesh sizes for suitability. A typical fine mesh kitchen strainer worked best. It retains Populus seeds but allows small debris to pass through.
Next, place seeds onto a single sheet of paper and manually remove larger pieces of debris such as residual trichomes or fragments of dry capsules with tweezers.
Some very small pieces of debris can be static and will adhere to the seed surface. These are removed by transferring seeds from one sheet of paper to another. Small static debris stays behind and remains on the sheet of paper. Repeat two or three times.
Transfer the seeds into a clean glass vial and store at -20 °C.
Notes: Seed storage conditions are critical.
The seeds have a delicate seed coat, hardly contain endosperm, and do not exhibit dormancy.
Fresh mature Populus seeds germinate rapidly under favorable conditions, but seed quality deteriorates quickly within weeks when released from the capsule under natural conditions or when seeds are stored at room temperature [17,18,24–26].
Based on seed storage characteristics, Populus seeds are considered suborthodox, and dry mature seeds can be kept at cold or subfreezing temperatures for prolonged preservation of seed viability [27].
Seeds extracted with our protocol that were stored immediately at -20 °C retain high viability for weeks and years (see Validation section 5). They had a seed moisture content (SMC) of approximately 10%, which is comparable to reported SMC values for mature, dry Populus seeds [27].
Germination test—to assess seed quality
Prepare MS plates as detailed under Recipes. Work in a laminar flow hood and allow plates to solidify and dry.
Notes:
Poplar seeds can be damaged easily by surface sterilization. Instead of surface sterilization, 0.2% PPM (final concentration) is added to the medium to prevent microbial contamination.
Alternatively, moist filter paper can be used for germination tests.
Set aside at least 100 seeds per condition for the germination test.
Sterilize tweezers using a micro bead sterilizer and let it cool down prior to handling seeds. Place seeds on a plate using sterilized tweezers and seal plates with micropore tape. Here, we used 50 seeds per plate and included at least two replicates. Work under a laminar flow hood.
Place plated seeds in a climate-controlled chamber under long-day conditions (16:8 h light/dark and 21/19 °C day/night).
Count germinated seeds three days after plating and calculate germination efficiency as number of germinated seeds per total seeds plated multiplied by 100.
Notes:
A seed is considered germinated when the radicle (embryonic root) has penetrated the seed coat.
Seed lots collected in previous years were included to study the effect of longer-term seed storage.
Plant growth and survival after germination was further recorded (see Validation section 5).
Data analysis
All data presented in the validation section were visualized using R and R Studio.
Validation of protocol
Selection of biological material
To test our seed extraction protocol, we selected the eastern cottonwood (Populus deltoides Bartr. ex Marsh.), a typical species from the genus Populus that is native to eastern North America. P. deltoides plants grow quickly into large trees on moist, well-drained sites, provide ecosystem services, and are used in commercial forestry and urban environments. Female mature trees were identified and closely monitored for seed development and maturation during spring and early summer between May and June of 2023 (Figure 1).
Characterization of reproductive structures and seed production in P. deltoides
To assess our seed extraction protocol for effectiveness, we first aimed to characterize reproductive structures and seed production in P. deltoides by studying three trees from our local study population in detail (Table 1, Materials and Reagents section). Seeds develop in capsules on female catkins. We counted the number of seeds per capsule and the number of capsules per catkin and determined the number of seeds per entire catkin (Figure 3). This was done by manually picking and individually counting all seeds that had developed inside a capsule, for each capsule on each of the catkins studied.
On average, we counted 13, 14, and 19 seeds per capsule for each of the three genotypes, with minimum and maximum values for seeds per individual capsule ranging from 1 to 38 (Figure 3A). Note that not every ovule will eventually produce a mature seed, leading to variable numbers of seeds per capsule. We further observed on average 41, 44, and 40 capsules per catkin, depending on genotype (Figure 3B). This amounted to approximately 516, 631, and 768 seeds per catkin on average for each of the three genotypes, with extreme values for individual catkins ranging from 391 to 921 seeds at maturity (Figure 3C). For comparison, Bessey [14] reported 32 seeds per capsule and 27 capsules per catkin for a single mature P. deltoides tree. This would amount 864 seeds per catkin. Similarly, our observations are well within the range described for P. deltoides in the Flora of North America [28]. The characterization of our material (Figure 3) provides now an exact reference point for our seed extraction protocol.
Figure 3. Characterization of seed production in eastern cottonwood (P. deltoides). Trees in the genus Populus produce catkins carrying several capsules and each capsule contains multiple seeds. The reproductive material was characterized thoroughly by manual counting. A. Number of seeds per individual capsule. B. Capsules per catkin. C. Seeds per entire catkin. Data is shown for three different genotypes (trees) and six randomly selected catkins per tree. Each dot represents an individual data point. Within each box in the boxplot, horizontal lines indicate the median, and boxes extend from the 25th to 75th percentile (i.e., the 1st and 3rd quartile). Whiskers represent 1.5 times the interquartile range between the 25th and 75th percentile.
Seeds are extracted effectively from P. deltoides reproductive structures using our protocol
Not all produced seeds can be extracted easily. Maisenhelder [18] reports, for example, that only 20% of eastern cottonwood seeds were readily extractable from seed-containing structures in Populus using the method described in his publication. Despite its potential relevance, effectiveness of extraction procedures for Populus seeds are, however, rarely reported.
We randomly assigned catkins to either manual seed counting (see also Figure 3) or mechanical extraction by shaking as described herein. Shaking allowed to extract on average 402, 591, and 635 seeds per catkin for the three genotypes studied (Figure 4, left panel). This means that between 78% and 83%–94% of all the seeds per catkin were extracted easily by shaking from the genotypes studied. Across all genotypes, 85% of the seeds present in a catkin were extracted easily with our protocol (Figure 4, right panel).
Figure 4. Extraction of seeds from P. deltoides reproductive structures using our protocol. The protocol presented herein allows to extract clean and healthy seeds effectively from P. deltoides cotton (dark grey, label: “Extracted”). For comparison, the total number of seeds that is present in the reproductive material was determined by manual handpicking and counting (white, label: “Present”). Catkins were used as reference unit for reproductive material. Data is given for three different genotypes (trees) and six randomly selected catkins per tree and extraction method (left panel). The right plot summarizes the data across the different genotypes.
Clean seeds are extracted quickly and efficiently from P. deltoides cotton
Time requirements are also of practical relevance. Therefore, we recorded the time spent on seed extraction and seed cleaning when using our protocol. This was compared with the time needed for manual seed picking. We selected one genotype, POP50, and documented extraction times in parallel for several lab members with different levels of work experience (undergraduate, graduate, research scientist). Very clean seeds were obtained, at a rate of 36 seeds per minute on average when using our protocol compared to only five seeds per minute, approximately, when seeds were picked manually (Figure 5).
When testing different materials for our manual shaker (Figure 2B), we observed that the shape of the mesh pencil holder had a clear influence on both seed yield and seed extraction speed. Round cups were less efficient than square mesh pencil cups. Therefore, we proceeded with square cups and recommend them for future work.
Figure 5. Clean seeds are extracted quickly from P. deltoides cotton. Seed extraction was timed for our extraction protocol and for handpicking using cotton from genotype POP50. Four to five extractions each were timed. The timing included both extraction and cleaning steps.
Extracted seeds are viable and suitable for tissue culture and long-term storage
Seeds need to be viable for assays and downstream experiments. To assess seed quality, viability was tested in a standard germination test. Populus seeds do not enter dormancy and germinate rapidly under favorable conditions, often within 24 h. However, seed longevity is limited in nature [5].
Seeds extracted with our protocol were subjected to two different storage conditions and tested at different time intervals (Figure 6). Fresh seeds that were tested directly after extraction and cleaning exhibited a high germination efficiency of 98% (Figure 6B), and the emerging seedlings were healthy and strong (Figure 6A). Viability declined by 5% when seeds were stored at room temperature for 15 days (93% germination efficiency), and the trend continued. After 30 days of seed storage at room temperature, germination efficiency dropped to 84% (Figure 6A and 6B). In contrast, seeds stored at -20 °C for the same period of time (30 days) retained a high germination efficiency of 94%, and seeds germinated vigorously (Figure 6A and 6B). Longer-term effects on seed viability were only studied for -20 °C storage conditions (Figure 6C and 6D). After one year of seed storage at subfreezing temperatures, seeds retained viability and germinated with a high efficiency of 95%. A germination efficiency of 79% was still observed after two years of storage at -20 °C, although for a different tree. It should also be noted that, after successful germination, almost all plants survived, continued to grow well, and established healthy plants (Figure 7), further highlighting seed quality and seed health.
The germination data confirm that the seeds extracted with our protocol are, despite of their delicate nature, intact and viable and germinate vigorously. They are very clean and suitable for plating, tissue culture, and experiments under sterile conditions. Seeds lose viability quickly under ambient conditions and should not be stored at room temperature. Storage at -20 °C, however, retains viability well for weeks, months, and years.
Figure 6. Viability of extracted P. deltoides seeds and effect of storage conditions. Seeds extracted with our protocol were subjected to standard germination tests. A. Images of representative plates with germinated cottonwood seedlings. Seeds were stored for 0, 15, and 30 days at either -20 °C or room temperature (RT). B. Germination efficiency is shown for the conditions described in A. C, D. Seeds collected in previous years were tested after storage at -20 °C over longer periods of time: one and two years. Again, typical plates with germinated seedlings as well as germination efficiencies are given. A–D. Seeds were plated on MS medium, and germination (radicle emergence) was scored three days after plating. Fifty seeds per plate were prepared in duplicate for the germination test. All data represent genotype POP50, except for the seeds that were stored for two years (POP29). Of note: fresh seeds and seeds stored at -20 °C for 15 days gave identical germination values across replicates. Thus, error bars are not visible for these conditions.
Figure 7. Populus deltoides plants continue to grow well after successful germination. Following a standard germination test, plants show healthy seedling growth on plates (A) and in tissue culture (B). A. Two-week-old plants, genotype POP50. B. Eight-week-old plant, genotype POP19. Scale bar: 1 cm.
Concluding remarks
Most plant life starts with a seed. Here, we describe a protocol for the collection and storage of seeds for Populus trees. This genus is of economic and ecological importance and serves as scientific model for deciduous trees. Seeds are delicate, short-lived, and attached to dense tufts of hair, needing to be separated and processed quickly. Our protocol describes a fast and effective way of collecting, extracting, and cleaning these seeds using simple laboratory and household items only. All equipment used in this protocol can be cleaned and sterilized easily. Seed quality and, critically, storage conditions have been tested. Germination tests confirm lasting quality of the obtained seeds for weeks, months, and up to years of storage. The obtained seeds are very clean and suitable for tissue culture and experiments under sterile conditions.
Acknowledgments
We thank Dana Al Refai and Stefan Heinen for their contributions to the seed counting data and seed plating. This work was generously supported by competitive funding awarded to KB from the National Sciences and Engineering Research Council of Canada (NSERC) (RGPIN-2017-06552), the Canadian Foundation of Innovation (CFI, 36678) and the University of Toronto (RSAF).
Competing interests
The authors declare no competing interests.
Ethical considerations
Ethics approval was not required for this study.
References
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Live Imaging Transverse Sections of Zebrafish Embryo Explants
EP Eric Paulissen
BM Benjamin L. Martin
Published: Vol 14, Iss 3, Feb 5, 2024
DOI: 10.21769/BioProtoc.4928 Views: 592
Reviewed by: Alberto RissoneMikiko Nagashima Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Feb 2022
Abstract
Vertebrate embryogenesis is a highly dynamic process involving coordinated cell and tissue movements that generate the final embryonic body plan. Many of these movements are difficult to image at high resolution because they occur deep within the embryo along the midline, causing light scattering and requiring longer working distances. Here, we present an explant-based method to image transverse cross sections of living zebrafish embryos. This method allows for the capture of all cell movements at high-resolution throughout the embryonic trunk, including hard-to-image deep tissues. This technique offers an alternative to expensive or computationally difficult microscopy methods.
Key features
• Generates intact zebrafish explants with minimal tissue disturbance.
• Allows for live imaging of deep tissues normally obscured by common confocal microscopy techniques.
• Immobilizes tissues for extended periods required for time-lapse imaging.
• Utilizes readily available reagents and tools, which can minimize the time and cost of the procedure.
Graphical overview
Keywords: Zebrafish explants Mesoderm Fluorescent microscopy Deep tissues Morphogenesis Imaging
Background
A central question in developmental biology is understanding how embryonic tissues organize themselves into the correct form. Embryos go through a series of cell/tissue interactions and broad dynamic movements to adopt their final shape. However, observing embryonic tissues in three-dimensional space can be challenging. Zebrafish are an excellent model system for visualizing living tissues. The relative transparency of zebrafish embryos compared to other model systems facilitates imaging of deeper tissues, and transgenic reporter lines allow for the visualization of specific cell or tissue types (Kimmel et al., 1995; Tonelli et al., 2017).
Despite the advantages of the zebrafish model, deep tissues can still be difficult to image at high resolution due to light scattering and the long working distance that is required (Jonkman and Brown, 2015). These issues can prevent detailed analysis of cell and tissue movements deep within a zebrafish embryo, such as the notochord or other tissues at the embryonic midline. Imaging of transverse sections can alleviate these problems, but prior methods require fixation and thus prohibit analyzing dynamic tissue movements in real time. We therefore developed a novel method of imaging living transverse sections of zebrafish using explants.
Zebrafish explants have been used frequently to understand various aspects of development (Langenberg et al., 2003; Picker et al., 2009; Simsek and Özbudak, 2021). Often, explants are used to separate tissues away from signaling sources or to conform tissues for specific analyses (Manning and Kimelman, 2015; Schauer et al., 2020). However, our methodology provides a novel way to orient intact embryonic trunk explants to image tissues of a transverse section. We first used this method to make time-lapse movies of the migration of angioblasts from bilateral regions of the embryo to the midline, where they differentiate into the dorsal aorta and cardinal vein (Paulissen et al., 2022). An advantage of this method is that it keeps most of all trunk tissues intact including the portions of the yolk to help stabilize the embryo. It allows for straightforward observation of cellular activity along the entire dorsal–ventral and medio–lateral axes, which would otherwise be difficult to capture. It also has an added benefit of using serum-less culture media that can be heated without worry of denaturation, which is relatively inexpensive compared to other methods that require serum-based media. Taken together, this new method provides an inexpensive, straightforward approach for visualizing deep-tissue movements at high-resolution during zebrafish embryogenesis.
Materials and reagents
Biological materials
Wild-type embryos used in this study were from hybrid adults generated from an inbred strain of locally acquired pet store fish (which we call Brian) crossed to the TL line (to generate TLB)
The tg(hsp70l:CAAX-mCherry-2A-NLS-KikGR)sbu104 transgenic strain was maintained on the TLB background (Goto et al., 2017). Our lab can provide this strain through resource sharing; however, many commercially available reporter lines can be used to label tissues of interest, including through the Zebrafish International Resource Center (ZIRC)
Reagents
NaCl (Millipore Sigma, catalog number: S9888)
KCl (Millipore Sigma, catalog number: P3911)
CaCl·2H2O (Millipore Sigma, catalog number: C7902)
KH2PO4 (Millipore Sigma, catalog number: P5655)
NaHPO4 (Millipore Sigma, catalog number: S5011)
MgSO4·7H2O (Millipore Sigma, catalog number: M1880)
Modified Barth's saline (1×) (MBS), liquid, without FicollTM 400 (Millipore Sigma, catalog number: F-04-B)
Agarose, low-gelling temperature (Millipore Sigma, catalog number: A4018)
Solutions
Embryo growth medium (see Recipes)
1.2% low-gelling agarose in MBS (see Recipes)
Recipes
Embryo growth medium
Reagent Final concentration Quantity
NaCl 15 mM N/A
KCl
CaCl·2H2O
KH2PO4
NaHPO4
MgSO4·7H2O
0.5 mM
1.3 mM
0.15 mM
0.05 mM
1.0 mM
N/A
N/A
N/A
N/A
N/A
H2O n/a Desired volume (usually 50 mL per clutch of eggs laid)
1.2% low-gelling agarose in MBS
Reagent Final concentration Quantity
MBS N/A 100 mL
Low gelling agarose 1.2% 1.2 g
Total N/A 100 mL
Laboratory supplies
Pyrex reusable Petri dishes, 100 mm (Fisher Scientific, catalog number: 08-747B)
Fisherbrand polystyrene Petri dishes, 100 mm × 15 mm (Fisher Scientific, catalog number: FB0875713)
Glass-bottom dish, 50 mm No. 1.5 coverslip 30 mm glass diameter uncoated (Mattek, catalog number: P50G-1.5-30-F)
NuncTM 15mL conical sterile polypropylene centrifuge tubes (Thermo Scientific, catalog number: 339650)
General purpose water bath set to 42 °C (Thermo Scientific, catalog number: TSGP20)
Microknives plastic handle, 22.5 degree cutting angle (Fine Science Tools, catalog number: 10316-14)
Dumont biological-grade forceps (Dumont, catalog number: 72700-D)
Bel-Art pipette pump 10 mL pipettor (Bel-Art, catalog number: F37898-0000)
FisherbrandTM disposable borosilicate glass Pasteur pipettes (Fisher Scientific, catalog number:13-678-20D)
Equipment
Custom assembled spinning disk confocal microscope consisting of an automated Zeiss frame, a Yokogawa CSU-10 spinning disc, a Ludl stage controlled by a Ludl MAC6000, and an ASI filter turret mated to a Photometrics Prime 95B camera
Zeiss Plan-Apochromat 40× Dipping Microscope Objective (Zeiss, catalog number: 421462-9900-799)
Leica S9E stereoscope with a Leica KL300 LED light source (Leica)
Software
The spinning disc microscope was controlled with Metamorph microscope control software and images were obtained with Metamorph (V7.10.2.240 Molecular Devices)
Procedure
Preparing embryos to generate trunk explants
This process details the preparatory steps for generating trunk explants. This includes the steps up to the point at which the embryo is cut.
Collect and store freshly laid embryos in polystyrene Petri dishes in embryo growth medium until roughly 2 h before the desired stage for sectioning.
Pyrex reusable dishes should be clean and sterile before sectioning process begins. Brand new dishes are preferable.
Prepare modified Barth’s saline (MBS) with low-gelling agarose. Melt 1.2% agarose powder into MBS in a laboratory microwave, avoiding boiling off the liquid as much as possible.
Place melted MBS/agarose mixture in a 15 mL polypropylene conical tube in a 42 °C water bath to prevent solidification. Allow at least 30 min for the MBS/agarose mixture to equilibrate to 42 °C.
Add approximately 25 mL of MBS to a clean Pyrex reusable dish.
Transfer embryos to be sectioned from embryo growth media to the Pyrex dish containing MBS using a fire-polished glass Pasteur pipette. Low amounts of embryo growth media (< 100 μL) are tolerated in the MBS. If this cannot be achieved, MBS should be changed to fresh media.
Using clean Dumont forceps, manually remove chorions from the embryos in the MBS media.
Sectioning and mounting trunk explants
This step is for the sectioning and mounting of embryo explants. The following steps require both speed and precision to prevent excess damage to the explant. Individual explants should be mounted soon after sectioning before continuing to the next explant. Our protocol maintains the yolk to allow for more accurate morphology and to prevent distention during mounting. All subsequent steps should be performed under a dissection stereomicroscope. We utilize the Leica S9E stereomicroscope, but most commercial stereomicroscopes are sufficient.
Using two Dumont forceps, carefully open a section of the yolk away from the sectioning area (see Figure 1A). This allows yolk to vent away from the sectioning area to prevent obscuring of the trunk region during imaging as well as preventing endoderm damage (Figure 1A and 1B).
Figure 1. Schematic of embryo section. A. Opening a puncture in the yolk of the embryo allows venting of yolk after sectioning. This reduces the need to remove yolk manually, which can damage the endoderm. Use forceps to puncture yolk. B. Image of puncture in an 8-somite-stage embryo (black arrows). C. Schematic showing the position of forceps and microknife during sectioning. D. Image of sectioned explant. Tail of the explant (T) and head of explant (H) are visible.
Immediately afterward, using the forceps to immobilize the embryo, section the area cleanly using a microknife (Figure 1C and 1D).
Remove the agarose/MBS mixture from the 42 °C water bath. Add a droplet of agarose/MBS (roughly 100 μL) to a Mattek glass-bottom dish. Quickly and carefully transfer the explant to the droplet using a fire-polished Pasteur pipette while avoiding adding too much MBS during the transfer (Figure 2A and 2B).
Figure 2. Schematic of embryo mounting in glass-bottom dish. A. Place a liquid drop of agarose/MBS on a glass-bottom dish. B. Immediately transfer the desired explant to the drop using a fire-polished pipette. C. Using forceps, gently orient the embryo until you adopt the proper position. A good marker of correct orientation is seeing a visible round notochord through the dissecting scope, as shown in the image (black arrow). D. Add MBS to cover the surface of the solidified agarose/MBS droplet. Using dip lenses may be necessary to image explant.
Prior to the agarose solidifying, carefully orient the explant using your Dumont forceps such that the transverse section is visible through the stereomicroscope (Figure 2C). Hold the embryo in place until the agarose cools.
Carefully remove your forceps from the droplet. Add additional liquid agarose/MBS to the periphery of the droplet if further stabilization of the explant is required; this can range from 100 to 500 μL as desired.
Let the agarose cool to solidification. Add additional MBS over explant until it is completely submerged; this is usually 2 mL but can depend on agarose volume added in previous step. The more agarose added, the less volume of MBS is required.
Imaging trunk explants
Imaging explants requires an inverted microscope or an upright microscope using water-immersion dipping objectives. Each methodology has strengths and weaknesses. With an inverted microscope, there is no need for specialized objectives when using the glass-bottom dishes. However, orienting the explant can be more challenging when using an inverted microscope, and the explant must also be pressed close to the coverslip. By contrast, an upright microscope works best in this context when using water immersion lenses that allow submersion in culture media. In this case, it is easier to correctly orient the explant (Figure 2D). Take care to not add excessive MBS to the glass-bottom dish to avoid spilling into the base of the microscope when the lens enters the media. Explants mounted by this method can last 6–8 h during timelapse.
Using this method can bring to light cellular movements that were previously difficult to document. This method was instrumental in our ability to visualize the migration of angioblasts from bilateral positions to the embryonic midline (Paulissen et al., 2022). Here, we provide another example of deep tissue imaging using this explant strategy. For our example, we image the developing neural tube as it transitions from the neural keel to the neural rod (Cearns et al., 2016). The developing neural tube is a large structure that can be difficult to image at high resolution without sectioning. Using a transgenic embryo that labels the cell membranes, tg(hsp70l:CAAX-mCherry-2A-NLS-KikGR)sbu104, we were able to observe cells as they transitioned from an unpolarized to polarized state (Figure 3A-3C, Supplemental Video 1).
Figure 3. Time-lapse imaging of cell polarization in the neural keel. A. Image of the neural keel starting at the 10-somite stage. B. Image of the neural keel after 60 min. Note the polarization of cells at the midline (red arrow). C. Image of neural keel after 120 min observation during the transition to the neural rod. Note the cells have polarized at the midline (red arrow ). Scale bars = 50 μm.
Considerations and limitations
This technique allows for long-term imaging of embryo explants starting from early somitogenesis stages. Explants from early to mid-somitogenesis stages can be imaged over an 8 h period before they begin to lose their fidelity. Explants from older embryos (24 hpf or older) can survive longer (up to 18 h) after being generated. The initial survivability of explants can vary depending on sectioning precision and the amount of agitation that occurs when transferring explants. If transferred carefully with minimal agitation, you can expect survivability of >50%. Explants, particularly at earlier stages, will still undergo broad morphogenetic movement including axial extension, which can cause the region of interest to move out of focus during time-lapse imaging. Removing unnecessary tissues like the tailbud can prevent this movement, which we demonstrated in a previous study (Paulissen et al., 2022). Anterior half explants containing the head, rather than the posterior half containing the tailbud, have more limited anterior–posterior axis extension and are preferable for imaging. We also note that these spatial issues can occur even with whole, intact embryos, and are not unique to this explanation method (Hirsinger and Steventon, 2017). Sectioning and explant generation can be performed at variable positions along the anterior-posterior axis as well as multiple positions. However, we recommend limiting this to two sectioning positions within one embryo as smaller explants are more difficult to transfer and orient.
Validation of protocol
The protocol presented here was performed previously in the reference listed below. A total of 12 explants were imaged (4 explants on three separate occasions).
• Paulissen, E., Palmisano, N. J., Waxman, J. S. and Martin, B. L. (2022). Somite morphogenesis is required for axial blood vessel formation during zebrafish embryogenesis. eLife 11: e74821. (Figure 7, panel O, Video 8, Video 9).
Acknowledgments
We thank Stephanie Flanagan for excellent fish care. We also thank members of the Matus lab for their helpful microscopy advice and generous use of their facilities. This work was supported by a National Institutes of Health training grant [T32 GM008468] to E.P., and National Science Foundation [IOS1452928] and NIGMS [R01GM124282, R35GM150290] grants to B.L.M.
Competing interests
The authors declare no competing or financial interests.
References
Cearns, M. D., Escuin, S., Alexandre, P., Greene, N. D. E. and Copp, A. J. (2016). Microtubules, polarity and vertebrate neural tube morphogenesis. J. Anat. 229(1): 63–74.
Goto, H., Kimmey, S. C., Row, R. H., Matus, D. Q. and Martin, B. L. (2017). FGF and canonical Wnt signaling cooperate to induce paraxial mesoderm from tailbud neuromesodermal progenitors through regulation of a two-step EMT. Development. e143578.
Hirsinger, E. and Steventon, B. (2017). A Versatile Mounting Method for Long Term Imaging of Zebrafish Development. J. Vis. Exp. 119:55210.
Jonkman, J. and Brown, C. M. (2015). Any Way You Slice It—A Comparison of Confocal Microscopy Techniques. J. Biomol. Tech. 26(2): 54–65.
Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B. and Schilling, T.F. (1995). Stages of embryonic development of the zebrafish.Dev. Dyn. 203, 253–310.
Langenberg, T., Brand, M. and Cooper, M. S. (2003). Imaging brain development and organogenesis in zebrafish using immobilized embryonic explants. Dev. Dyn. 228(3): 464–474.
Manning, A. J. and Kimelman, D. (2015). Tbx16 and Msgn1 are required to establish directional cell migration of zebrafish mesodermal progenitors. Dev. Biol. 406(2): 172–185.
Paulissen, E., Palmisano, N. J., Waxman, J. S. and Martin, B. L. (2022). Somite morphogenesis is required for axial blood vessel formation during zebrafish embryogenesis. eLife 11: e74821.
Picker, A., Roellig, D., Pourquié, O., Oates, A. C. and Brand, M. (2009). Tissue Micromanipulation in Zebrafish Embryos. Methods Mol. Biol. 546: 153–172.
Schauer, A., Pinheiro, D., Hauschild, R. and Heisenberg, C. P. (2020). Zebrafish embryonic explants undergo genetically encoded self-assembly. eLife 9: e55190.
Simsek, M. F. and Özbudak, E. M. (2021). A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish. J. Vis. Exp. e61981.
Tonelli, F. M., Lacerda, S. M., Tonelli, F. C., Costa, G. M., de França, L. R. and Resende, R. R. (2017). Progress and biotechnological prospects in fish transgenesis. Biotechnol. Adv. 35(6): 832–844.
Supplementary information
The following supporting information can be downloaded here.
Supplemental Video 1
Article Information
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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Quantification of Macrophage Cellular Ferrous Iron (Fe2+) Content using a Highly Specific Fluorescent Probe in a Plate-Reader
PG Philipp Grubwieser
NB Natascha Brigo
MS Markus Seifert
MG Manuel Grander
IT Igor Theurl
MN Manfred Nairz
GW Günter Weiss
CP Christa Pfeifhofer-Obermair
Published: Vol 14, Iss 3, Feb 5, 2024
DOI: 10.21769/BioProtoc.4929 Views: 759
Reviewed by: Alka Mehra Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Frontiers in Microbiology Aug 2023
Abstract
Macrophages are at the center of innate immunity and iron metabolism. In the case of an infection, macrophages adapt their cellular iron metabolism to deprive iron from invading bacteria to combat intracellular bacterial proliferation. A concise evaluation of the cellular iron content upon an infection with bacterial pathogens and diverse cellular stimuli is necessary to identify underlying mechanisms concerning iron homeostasis in macrophages. For the characterization of cellular iron levels during infection, we established an in vitro infection model where the murine macrophage cell line J774A.1 is infected with Salmonella enterica serovar Typhimurium (S.tm), the mouse counterpart to S. enterica serovar Typhi, under normal and iron-overload conditions using ferric chloride (FeCl3) treatment. To evaluate the effect of infection and iron stimulation on cellular iron levels, the macrophages are stained with FerroOrange. This fluorescent probe specifically detects Fe2+ ions and its fluorescence can be quantified photometrically in a plate reader. Importantly, FerroOrange fluorescence does not increase with chelated iron or other bivalent metal ions. In this protocol, we present a simple and reliable method to quantify cellular Fe2+ levels in cultured macrophages by applying a highly specific fluorescence probe (FerroOrange) in a TECAN Spark microplate reader. Compared to already established techniques, our protocol allows assessing cellular iron levels in innate immune cells without the use of radioactive iron isotopes or extensive sample preparation, exposing the cells to stress.
Key features
• Easy quantification of Fe2+ in cultured macrophages with a fluorescent probe.
• Analysis of iron in living cells without the need for fixation.
• Performed on a plate reader capable of 540 nm excitation and 585 nm emission by trained employees for handling biosafety level 2 bacteria.
Graphical overview
Keywords: Salmonella Typhimurium Macrophages FerroOrange Iron Iron quantification Fluorescence
Background
Human typhoid fever, a severe and often life-threatening infectious disease, is caused by the facultative intracellular Gram-negative bacterium Salmonella enterica serovar Typhi, leading to major health loss globally (Stanaway et al., 2019). For murine infection models, the strain Salmonella enterica serovar Typhimurium (Salmonella Typhimurium, S.tm), which causes a systemic disease in mice but a self-limiting gastroenteritis in humans, is frequently used.
In the case of a bacterial infection, invading pathogens are phagocytized by macrophages, the first line of innate immune defense (Weiss and Schaible, 2015). S.tm, however, thrives within macrophages. Despite its ability to invade various types of cells, virulence is dependent on intramacrophage proliferation (Fields et al., 1986; Leung and Finlay, 1991). One factor critically affecting the outcome of this host–pathogen interaction is the availability of nutrients, as intramacrophage S.tm depends on the acquisition of essential nutrients to sustain efficient proliferation. This includes amino acids or trace metals, like iron. On one hand, S.tm–driven metabolic reprogramming grants the pathogen access to intracellular nutrients (Liss et al., 2017). On the other hand, macrophages exploit this nutrient demand by withdrawing iron from the spatio-temporal localization of the pathogen in a process termed nutritional immunity (Nairz et al., 2007; Murdoch and Skaar, 2022). Appropriately, a state of systemic or cellular iron excess is associated with increased bacterial proliferation (Khan et al., 2007; Porto and De Sousa, 2007; Kao et al., 2016).
Macrophage cellular iron metabolism and its adaptation to an infection have been extensively studied. In the case of an infection with an intracellular pathogen like S.tm, regulation of the key iron metabolism proteins Ferroportin-1 (FPN1) and the Transferrin receptor-1 (TFR1) facilitates a decrease of cellular iron content and thus leads to an improved infection control of intracellular bacteria (Nairz et al., 2008; Fritsche et al., 2012; Wessling-Resnick, 2015; Abreu et al., 2020). Furthermore, transport of iron into the cytosolic lumen is accomplished by the divalent metal transporter-1 (DMT1) and by the natural resistance-associated macrophage protein-1 (NRAMP1 or SLC11A1), both of which have also been implicated in bacterial iron withdrawal (Forbes and Gros, 2003; Fritsche et al., 2012; Grander et al., 2022). DMT1 is responsible for the uptake of non-transferrin-bound iron (NTBI) from outside the macrophage and for transporting transferrin-bound iron (TBI) from the early endosome into the lumen of the phagocyte. NRAMP1 transports iron out of the late phagosome. As both are only capable of binding Fe2+, the function of the six-transmembrane epithelial antigen of prostate 3 (STEAP3) in the late endosome to reduce Fe3+ to Fe2+ is indispensable (Wang and Pantopoulos, 2011).
Another factor at play is the acute phase protein hepcidin (HAMP1), a liver-derived hormone regarded as the systematic master regulator of iron metabolism. During an infection, hepcidin targets the iron exporter FPN1, leading to its degradation and thus iron sequestration, causing hypoferremia (Nemeth et al., 2004). However, its role in an infection with intracellular pathogens is not yet completely understood (Chlosta et al., 2006; Lim et al., 2018).
During infection studies, cellular iron quantification is frequently relevant. Standard procedures like the usage of radioactive 59Fe isotopes are elaborate, expensive, and might be inapplicable in certain experimental settings. Iron quantification with the help of the quenchable probe Calcein-AM is easily available and often used but has several disadvantages. Acquiring fluorescence by flow cytometry needs extensive preparation of samples that exposes cells to mechanical stress; furthermore, as Calcein does not pass into the cellular membrane compartments (e.g., lysosomes), which are rich in labile iron (chelatable), total cellular iron is most likely drastically underestimated when this method is applied (Tenopoulou et al., 2007).
Herein, we report a simple and powerful tool to accurately determine alterations in cellular iron levels upon diverse stimuli, which can be employed for cultured immune cells, here exemplified in a murine macrophage infection with intracellular bacteria. As the cellular iron trafficking machinery primarily utilizes Fe2+, monitoring metabolically active intracellular ferrous iron levels is most insightful (Hentze et al., 2010; Moroishi et al., 2011; Cronin et al., 2019). We apply a fluorescent probe that specifically detects Fe2+, based on N-oxide chemistry (RhoNox-4; commercial name: FerroOrange) (Hirayama et al., 2020). By using this approach, cellular levels of the trace metal can be quantified in a plate reader without additionally exposing cells to stressors.
Materials and reagents
12-well plate (Falcon, catalog number: 353043)
Acridine orange/propidium iodide stain (Biocat, catalog number: F23001)
Agar-Agar Kobe I (Roth, catalog number: 5210.3)
Aqua bidest (Fresenius Kabi, catalog number: 16.231)
CASY Cup (OMNI Life Science, catalog number: 5651794)
CASY Ton buffer (OMNI Life Science, catalog number: 5651808)
Cell scraper (Sarstedt, catalog number: 83.3951)
CoolCellTM LX freezing container (Merck, catalog number: BCS-405G)
Cryo vial with silicone washer, 2 mL (Simport, catalog number: T311-3)
Dimethylsulfoxide (DMSO) (Roth, catalog number: A994.1)
Disposable cuvette (BRAND, catalog number: 759015)
Disposable pipettes, 5 mL, 10 mL, and 25 mL (Falcon, catalog number: 606180, 607180, and 357525, respectively)
Dulbecco’s modified Eagle’s medium (DMEM) (Pan BiotechTM, catalog number: P04-01500)
Eppendorf tubes, 0.5 mL (Eppendorf, catalog number: 0030121.023)
Erlenmeyer flask, 250 mL (Stoelzle Medical, catalog number: 21226368000)
Ferric chloride (FeCl3) (Sigma, catalog number: 236489)
FerroOrange (GERBU Biotechnik GmbH, catalog number: F374-10)
Fetal bovine serum (FBS) (Pan BiotechTM, catalog number: P30-3031)
Gentamicin (Gibco, catalog number: 15750-037), stock: 50 mg/mL
Glass beaker, 150 mL (Ruprechter, catalog number: 102113729)
Glycerol (Sigma, catalog number: G5516-100ML)
Iscove’s modified Dulbecco's medium (IMDM) (Pan BiotechTM, catalog number: P04-20150S3)
L-Glutamine (Lonza, catalog number: BE17-605E)
Luna cell counting slides (Biocat, catalog number: L201B1C3GB)
Lysogeny broth (LB) medium Lennox (Roth, catalog number: X964.2)
Penicillin/streptomycin (Capricorn Scientific, catalog number: PS-B)
Phosphate buffer saline (PBS) (Lonza, catalog number: 17-515 F)
Pipetman L Starter Kit, 2, 20, 200, and 1,000 μL pipettes (GILSON, catalog number: F167370)
Pipette tips, 10, 200, and 1,250 μL (STARLAB, catalog number: S1110-3700, S1120-3810, and S1112-1720, respectively)
Polypropylene tube, 50 mL (Falcon, catalog number: 352070)
Salmonella enterica serovar Typhimurium ATCC14028 (ATCC)
Sartorius Midi Plus pipetting controller (Sartorius, catalog number: 710931)
Tissue culture flask, 750 mL, straight neck (Falcon, catalog number: 353028)
LB medium (see Recipes)
LB medium with 30% glycerol (see Recipes)
Cell culture medium (see Recipes)
Cell culture medium for infection with S.tm (see Recipes)
Staining solution (see Recipes)
Iron (III) chloride solution (see Recipes)
Cell line
J774A.1 (ATCC TIB-67TM) is a macrophage cell line isolated in 1968 from a female BALB/c mouse with reticulum cell sarcoma.
Recipes
LB medium
200 mL aqua bidest
2 g LB medium powder
Autoclave (20 min at 121 °C and 10 min at 50 °C)
LB medium with 30% glycerol
Add 300 μL of glycerol to 700 μL of LB medium
Cell culture medium
500 mL of DMEM
50 mL of FBS
5 mL of L-Glutamine
5 mL of Penicillin/Streptomycin
Cell culture medium for infection with S.tm
500 mL of DMEM
5 mL of FBS
5 mL of L-Glutamine
Staining solution
5 mL of IMDM
1 μL of FerroOrange (1 mmol/L)
Iron (III) chloride solution
135 mg of FeCl3
50 mL of Aqua bidest
Equipment
Automated multimode microplate reader (TECAN Spark, catalog number: 1912001805)
Casy counting system CASY TT (OMNI Life Science, catalog number: TT2QA2571)
Centrifuge (Hettich Micro 200R and Rotanta 460R)
Freezer, -80 °C (Thermo Fisher Scientific, catalog number: 15788587)
CO2 incubator (Thermo Fisher Scientific, model: Heraeus® HERAcell®)
Laminar flow cabinet (EuroClone Sicherheitswerkbank Safe Mate Eco 1.2) (Politakis Laborgeräte, catalog number: EN 12 469)
Liquid nitrogen storage tank (CryoShop, catalog number: CS-79105601)
LUNA automated cell counter (Biocat, catalog number: L10001-LG)
Millivac-Maxi vacuum pump (Merck, catalog number: SD1P014M04)
Photometer (Eppendorf, catalog number: BioPhotometer D30)
Shaking incubator (VWR, catalog number: GFL 3031)
Software and datasets
GraphPad Prism 9.1 (GraphPad Software)
SparkControlTM (Spark Method Editor V.3, release date 2021 06 01) (TECAN Trading, Ltd.)
Procedure
Thawing of the J774A.1 macrophage cell culture aliquot
Note: Perform the next steps in a sterile laminar flow cabinet.
Preheat a water bath to 37 °C.
Preheat the cell culture medium (see Recipes).
Prepare a 50 mL polypropylene tube with 25 mL of preheated cell culture medium.
Take the J774A.1 cell culture aliquot from the liquid nitrogen storage and incubate the cells in the water bath until the aliquot is almost completely thawed.
Pipette the thawed cell culture aliquot into the previously prepared 50 mL polypropylene tube.
Centrifuge the cells at 300× g for 5 min.
Discard the supernatant in a 150 mL glass beaker.
Resuspend the cell pellet in 25 mL of cell culture medium (see Recipes).
Pipette the cells into a 750 mL tissue culture flask.
Place the cell culture flask into a cell incubator at 37 °C with 5% CO2 and 95% humidity as growth conditions.
Splitting of the J774A.1 cell culture
Note: Perform the next steps in a sterile laminar flow cabinet.
Remove the complete cell culture medium using a peristaltic pump.
Wash the cells with 10 mL of PBS.
Remove the PBS using a peristaltic pump.
Add 10 mL of cell culture medium .
Use a disposable cell scraper to scrape off the cells.
Prepare a new cell culture flask with 24 mL of cell culture medium.
Pipette 1 mL of the scraped off cells into the previously prepared cell culture flask.
Place the cell culture flask into a cell incubator at 37°C with 5% CO2 and 95% humidity as growth conditions.
Check the density of the cells every second day.
If the cells cover up to 90% of the cell culture flask’s surface, further split the cells into a new cell culture flask.
Freezing and storage of the J774A.1 cells
Note: Perform the next steps in a sterile laminar flow cabinet.
Perform steps as described in Section B1–5.
Pipette the scraped off cells into a 50 mL polypropylene tube.
Pipette 9 μL of the cell suspension into a 0.5 mL Eppendorf tube.
Mix 1 μL of the acridine orange/propidium iodide stain solution to the 9 μL cell suspension.
Pipette the 10 μL of the stained cells into a Luna cell counting slide.
Measure the cell number using the LUNA-FL fluorescent and brightfield automated cell counter.
Centrifuge cells at 300× g for 5 min.
Prepare the freezing mix (DMEM containing 10% FBS, 1% L-glutamine, and 1% penicillin/streptomycin + 8% DMSO).
Discard the supernatant into a 150 mL glass beaker.
Resuspend the cell pellet in FBS to a concentration of 1 × 107 cells/800 μL.
Mix 800 μL of the cells in FBS with 800 μL of freezing mix.
Aliquot 800 μL of this cell suspension into Cryo tubes.
Label the Cryo tubes with the cell type, the number of cell passages, and the date of storage.
Put the Cryo tubes into a Cool CellTM freezing container and leave it at -80 °C for two days.
Place the Cryo tubes on liquid nitrogen for long-term storage.
Seeding of the J774A.1 cells on 12-well plates
Note: Perform the next steps in a sterile laminar flow cabinet.
Remove the complete cell culture medium using a peristaltic pump.
Wash the cells twice with 10 mL of PBS.
Remove the PBS using a peristaltic pump.
Add 10 mL of DMEM containing 1% FBS and 1% L-glutamine.
Note: 1% FBS is used to reduce the amount of iron in the cell culture medium. This reduction of FBS is necessary to achieve the iron overload with the iron stimulus.
Use a disposable cell scraper to scrape off the cells.
Pipette 9 μL of the cell suspension into a 0.5 mL Eppendorf tube.
Mix 1 μL of the acridine orange/propidium iodide stain solution to the 9 μL cell suspension.
Pipette the 10 μL of stained cells into a Luna cell counting slide.
Measure the cell number using the LUNA-FL fluorescent and brightfield automated cell counter.
Seed 2.5 × 105 cells in 1 mL of DMEM containing 1% FBS and 1% L-glutamine in a 12-well plate.
Incubate cells overnight in a cell incubator at 37°C with 5% CO2 and 37°C and 95% humidity as growth conditions.
Treatment of the J774A.1 cell culture with iron
Note: Perform the next steps in a sterile laminar flow cabinet.
Treat the cells with 50 μM of FeCl3 [iron (III) chloride solution; see Recipes] or leave them untreated for 5 h (Figure 1).
Figure 1. Timeline and 12-well plate layout of the experimental procedure described in this protocol. A. Timeline of the experimental procedure depicting distinct experimental phases. B. 12-well plate layout depicting the arrangement of treatments and replicates used in this protocol.
Incubate cells for 5 h in a cell incubator at 37 °C with 5% CO2 and 95% humidity as growth conditions.
Preparation of Salmonella typhimurium (S.tm) stock as described in Brigo et al. (2022)
Take an aliquot of Salmonella enterica serovar Typhimurium ATCC14028 from -20 °C storage.
Thaw the aliquot at room temperature.
Note: Perform the next steps in a sterile laminar flow cabinet.
Pipette 10 μL of S.tm into 10 mL of LB medium in a 250 mL Erlenmeyer flask and cover the top of the flask using a tin foil.
Incubate at 37 °C overnight in a shaking incubator at 200 rpm.
The following day, pipette 50 μL of the overnight culture into fresh 10 mL of LB medium in a 250 mL Erlenmeyer flask and cover the top with tin foil.
Dispose of the remaining overnight culture of S.tm in an appropriate designated biosafety level 2 waste and wash and sterilize the 250 mL Erlenmeyer flask.
Incubate the culture at 37 °C for 1–2 h at 200 rpm in a shaking incubator.
Measure OD600 to check if S.tm reached 0.5:
Calibrate a photometer by measuring the blank with 500 μL of LB medium in a disposable cuvette.
Pipette 500 μL of the S.tm culture in a new disposable cuvette and measure.
S.tm reaches the optimal logarithmic growth phase when OD600 is between 0.5 and 0.7.
Note: If the OD600 value is below 0.5, continue the incubation of the culture in the 250 mL Erlenmeyer flask as described above until the OD600 value of 0.5 is reached. Of note, S.tm density duplicates every 20 min. If the OD600 value is above 0.7, dilute the culture 1:1 with LB medium and incubate the culture in the 250 mL Erlenmeyer flask until reaching an OD600 value of 0.5.
Transfer the culture into a 50 mL polypropylene tube or continue with counting of S.tm (Section G); keep the bacteria on ice for counting using the Casy counting system and afterwards until infection of the cells.
Centrifuge the S.tm culture at 5000× g for 5 min at room temperature.
Remove the supernatant by using a peristaltic pump.
Resuspend the pellet in 1 mL of freshly prepared LB medium with 30% glycerol (see Recipes).
Prepare 50 μL aliquots in 0.5 mL Eppendorf tubes.
Store the aliquots at -20 °C.
Counting of viable S.tm using a Casy counting system as described in Brigo et al. (2022)
Note: The setting up procedure and the programs used for the Casy counting system have been described in (Pfeifhofer-Obermair et al., 2022; Brigo et al., 2022). Alternatively, another method of counting/quantifying bacteria may be used (i.e., manual counting via microscopy).
Use the 45 μm capillary.
Measure the background by placing a new Casy cup with fresh 10 mL of Casy Ton buffer under the measuring unit.
Select the program for the background measurement.
Measure the background. It should be below 30 counts and 1 μm size. Otherwise, wash the system.
Prepare a new Casy cup with 10 mL of Casy Ton buffer and add 5 μL of S.tm OD600 0.5.
Shake gently.
Place the sample under the measuring unit.
Select the program for measuring between 1 and 3 μm.
Measure.
Click Next to get the number of viable counts/mL = viable S.tm /mL.
Note: Viable counts from a freshly prepared S.tm culture with an OD600 of 0.5 are between 2.5 × 108 and 3 × 108 viable counts/mL.
Infection of the J774A.1 macrophage cell line with S.tm on a 12-well plate
Note: Perform the next steps in a sterile laminar flow cabinet.
Infect the desired wells with a cell density of 2.5 × 105 cells with S.tm using a multiplicity of infection (MOI) of 10; therefore, 10 times more S.tm than cells are added to each well.
Example for calculation of S.tm:
To gain a MOI of 10, multiply the cell number × 10: 2.5 × 105 × 10 = 2.5 × 106.
Divide the calculated cell number against the gained viable Salmonella count (e.g., 2.5 × 108):
2.5 × 106 viable counts/mL: 2.5 × 108 viable counts/mL = 0.01 mL = 10 μL
Add the calculated amount of S.tm directly into the desired cell culture wells containing cell culture medium for infection with S.tm.
Incubate the cells for 1 h in a cell culture incubator (37 °C, 5% CO2, 95% relative humidity).
Gentamicin neutralization assay and continued treatment of the J774A.1 cell culture with iron
Pre-warm the medium and PBS in a water bath at 37 °C to avoid additional stress to the cells.
Remove the medium containing non-phagocytosed S.tm using a cell culture peristaltic pump.
Wash the cells twice with 1 mL of PBS + 25 μg/mL gentamicin.
Add 1 mL of DMEM supplemented with 1% FBS, 1% L-glutamine and 25 μg/mL gentamicin.
Treat the cells again with 50 μM FeCl3 [Iron (III) chloride solution] or leave them untreated.
Incubate the cells for up to 24 h in a cell culture incubator (37 °C, 5% CO2, 95% relative humidity).
Staining of the J774A.1 cells with FerroOrange
Thaw an aliquot of FerroOrange.
Prepare staining solution (see Recipes).
Remove the cell culture media using a peristaltic pump.
Gently wash the cells twice with 1 mL of PBS.
Remove the PBS using a peristaltic pump.
Add 200 μL of the prepared staining solution into each cell culture well.
Incubate the cells for 30 min in a cell incubator at 37 °C with 5% CO2 and 95% humidity as growth conditions.
Place the 12-well plate into a preheated and pre-set 5% CO2 Spark plate reader.
Setup of the Spark plate reader
A captured image of the setup screen is displayed below (Figure 2).
Figure 2. Screenshot of Spark plate reader method editor setup
Open the Spark plate reader method editor.
Select a 12-well plate.
Note: Select the correct model of a 12-well plate, depending on its commercial source, to avoid inaccurate measurements in “bottom read mode” and to avoid damaging the detector.
Select measurement Fluorescence Intensity.
Set the instrument temperature to 37 °C.
Set the CO2 flow to 5%.
Change the Mode to Bottom for fluorescence intensity bottom reading.
Change the Fluorophore Setting to other.
Set the fluorophore parameters to:
Monochromator 540 nm excitation; bandwidth 20 nm.
Monochromator 585 nm emission; bandwidth 20 nm.
Leave Flashes at 30.
Change the gain to manual and use 140.
Set the z-position to manual.
Note: The z-position can be calibrated right before the measurement and changed if needed.
Select multiple reads per well:
User defined.
Type: filled circles (4 × 4).
Border: 1450.
Save the setup.
Open Spark plate reader control.
Load the FerroOrange measurement setup.
Confirm that the temperature is at 37 °C and CO2 is at 5%.
Place the 12-well plate into the spark plate reader.
Calibrate the z-position in the machine (Figure 3):
Select z-position in the menu on the bottom left.
In the Scan sub-menu, select wells for signal detection.
Click the Scan button on the bottom left.
In the resulting graph, signal strength for different z-positions is visible.
Figure 3. Screenshot of Spark plate reader calibrating the z-position. In the z-position panel of the SparkControlTM software, the user is able to measure the signal strength (x-axis) over multiple z-positions (y-axis) to determine the optimal z-position for the assay.
Note: The z-position determines how far apart the measuring device is from the microplate's surface. You can change it by moving the plate up and down. When light bounces off the liquid on the sample, adjusting the Z-position helps to maximize signal-to-noise ratio.
Apply automatically suggested z-position to the entire plate or enter a manual value.
Click Start.
After the measurement is completed, an Excel file will open automatically.
Save this file and proceed to data analysis.
Data analysis
Open the generated Excel file. The Excel file shows all the performed actions in the experimental setup.
Scroll down until the measured data shows up.
Each individual measurement of the 4 × 4 multiple reads per well is shown in the table. The position of each individual measurement is displayed in a map, also found in the Excel file (see Figure 4).
Figure 4. Position of the multiple reads per well as found in the Excel table (left) and an illustration of multiple measurements in one well (right). Reprinted from Brigo et al. (2023).
In the table, in the lines below the assay information (i.e., time and temperature), the mean and standard deviation of all individual measurements of one well are displayed (exemplary output Excel available under the Validation section below).
Open GraphPad Prism.
Select a column graph in GraphPad Prism.
Label the columns according to the groups.
Add the technical replicates of each group into each column.
Check the graph.
Label the y-axis as fluorescent intensity of FerroOrange.
Label the x-axis as 24 h stimulation.
Note: An example graph of the measured 24 h time point is shown in Figure 5.
Figure 5. Fluorescent intensity of FerroOrange after 24 h of iron treatment with or without infection. Data from three experiments, done in triplicates (n = 9), are shown as mean fluorescent intensity ± SD, normalized to uninfected control conditions (dashed line). * denotes p < 0.05 and *** denotes p < 0.001, as evaluated by one-way ANOVA.
General notes and troubleshooting
General notes
Here exemplified in an infection model of the macrophage cell line J774A.1, this protocol may be applied to other macrophage models like bone marrow–derived macrophages or the human THP-1 cell line.
As the manufacturer implies in the technical information sheet (DOJINDO, product code F374), FerroOrange fluorescence intensity may be observed and quantified using a confocal fluorescence microscope (Cy3-channel) or flow cytometer (Weber et al., 2020). To the best of our knowledge, the FerroOrange dye is not fixable.
Troubleshooting
Potential problems include low signal, implausible measurements, or inappropriate variance between replicates or individual experiments. To counteract this, assay details should be verified: cells should be seeded uniformly in the well. Although this protocol suggests multiple measurements per well, a homogeneous distribution of cells will allow most consistent measurements. The correct z-position of the 12-well plate is vital for optimal signal-to-noise ratio. This calibrated z-position may need to be changed if 12-well plates from other manufacturers are used. As FerroOrange is time and light sensitive, accurate results may only be obtained with fresh staining aliquots thawed and used immediately. At last, as FerroOrange may leak out of cells, the medium should not be changed before the measurement.
Validation of protocol
This protocol was adapted and modified after Grubwieser et al. (2023).
For this protocol, we used a 12-well plate layout as illustrated in Figure 1. This enabled simultaneous measurement of three technical replicates (wells) for each experimental condition. Furthermore, the experiment was repeated independently three times, to increase the robustness of findings (Figure 5). An exemplary output Excel file of one experiment is shown in Figure 6.
Figure 6. Exemplary output Excel file of one experiment. In the output Excel file, details of the performed measurement are listed (panel 1, blue). Beneath, the result table shows data from multiple measurements (panel 2, orange). The experimental conditions and user calculations are depicted in panel 3 (green).
Acknowledgments
G.W. is supported by grants from the Christian Doppler Society and the Austrian research Funds (FWF doctoral program W1253 HOROS; and FWF, DOC 82 doc.fund). P.G. and I.T. were supported by the Austrian Science Fund (FWF, DOC 82 doc.fund). N.B. was supported by FWF doctoral program - W1253 HOROS. M.N. was funded by the Austrian Science Fund (FWF, P33062). This protocol has been used in Grubwieser et al., 2023.
Competing interests
The authors declare no conflicts of interest.
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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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Immunology > Immune cell function > Macrophage
Cell Biology > Cell metabolism > Other compound
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