Isolation and Characterization of Single Cells from Zebrafish Embryos

This protocol requires the use of live, adult zebrafish to produce embryos. The embryos are harvested for tissue collection. It is essential to obtain approval from appropriate ethics review boards to conduct this experiment.

1. Obtain Staged Embryos

1. The day before the experiment, prepare healthy, adult zebrafish for breeding. Place one male and one female on opposite sides of a clear divider in a breeding tank.
2. Repeat 1.1 for as many breeding tanks as necessary for sufficient embryo production for the downstream application. Obtain embryos from both wild type fish and transgenic fish that express fluorescent proteins in the cell type of interest.  
   NOTE: The number of embryos needed for downstream applications in Steps 2-8 depends on the relative abundance of the cells of interest at the time point of interest. Though this may vary by cell type, 200 embryos produce 2,000-5,000 sorted cells when the cells of interest represent <1.0% of the total cells at 24 hpf (hours post-fertilization).
3. The next morning, change the water in the breading tank by transferring fish to a fresh breeding tank and remove the divider. Tilt the tank at an angle to encourage breeding.
4. Collect staged embryos.
   1. Every 15 min, collect embryos by transferring the adults to a fresh breeding tank and passing the eggs which are left behind through a tea strainer.
      NOTE: Zebrafish embryos develop synchronously when maintained at comparable densities and temperatures.
   2. Rinse the eggs with Egg Water (0.21 g/L Instant Ocean salts in 1 L double distilled water) and transfer to a petri dish. Transfer the petri dish to a humid incubator at 28.5 °C with air circulation.
5. Two hours after the last collection, sort fertilized, multicellular embryos into 10 cm petri dishes and reduce density to 50 embryos per dish. Select embryos from a single, 15 min time window of collection for downstream application. Incubate embryos at 28.5°C.  
   NOTE: For example, collect embryos at 8:30, 8:45, 9:00, 9:15, 9:30, 9:45, 10:00 and 10:15 AM. Comparing across time points, if the largest number of fertilized embryos are from the clutches collected at 9:00, then use only these embryos for downstream applications.

2. Set Up for Single Cell Dissociation

1. Approximately 30 min prior to the time point of interest (18 hpf) remove embryos from their chorion manually with fine forceps.
2. Collect and label the following for each condition: two 2 ml microcentrifuge tubes, one 40 µm cell strainer, one 35 mm cell culture dish, and two FACS tubes topped with a 35 µm cell strainer.
3. Chill the following reagents on ice: Egg Water containing 0.21g/L Instant Ocean salts in 1L double distilled water; De-yolking Buffer containing 55 mM NaCl, 1.8 mM KCl, and 1.25 mM NaHCO₃; and FACS Buffer containing Leibovitz's L-15 media supplemented with 5% heat-inactivated fetal bovine serum.
4. Bring 1 ml per sample of Cell Dissociation Reagent 1 to RT. Thaw 1 ml Cell Dissociation Reagent 2 per sample on ice. Bring to RT immediately before use.

3. Single Cell Dissociation

1. Using a wide bore glass pipette, transfer 100-300 embryos to a 2 ml micro centrifuge tube in a minimum volume of Egg Water.
2. Euthanize embryos by replacing water with 1 ml ice-cold Egg Water and submerging tube in ice for 20 min.
   CAUTION! It is critical to obtain approval from proper institutional animal care oversight committee for this euthanasia method (chilling on ice followed by cell dissociation as secondary euthanasia). Standard euthanasia typically requires that embryos <3 days post-fertilization are bleached after they are chilled, which is not appropriate for obtaining viable cells.
3. Wash the embryos two times with 1 ml ice-cold Egg Water. To wash, use a wide bore glass pipette to remove Egg Water and a P1000 to add Egg Water.
4. Remove the yolk by replacing the embryo water with 1 ml Deyolking Buffer and triturating 8-12 times with a P1000 tip, or until the yolk is dissolved and only the bodies of the embryos are visible.
5. Collect the tissue by centrifugation at 300 x g for 1 min. Use a pipette to gently remove the supernatant without disrupting the tissue pellet. Re-suspend in 1 ml Egg Water.
6. Repeat step 3.5 for a total of three washes, but on the final wash, re-suspend in 1 ml RT Cell Dissociation Reagent 1.
7. Incubate in Cell Dissociation Reagent 1 for 10 min at RT with tubes placed horizontally. Every 2-3 min, gently triturate with a P1000 pipette to prevent clumping.
   NOTE: Handle samples gently during trituration steps. Sometime a single, large clump will form in the tube from a tangle of embryo bodies. This will disperse with additional digestion aided by gentle trituration. DO NOT VORTEX.
8. Collect tissue by centrifugation at 300 x g for 3 min. Remove supernatant and resuspend in 1 ml Cell Dissociation Reagent 2.
9. Incubate for 5-15 min at RT. Place the tubes horizontally. Every 2-3 min, gently triturate to prevent clumping. Every 5 min, assess digestion progress.
   1. To assess digestion progress, dilute 2 µl supernatant into 18 µl FACS Buffer and pipette as a droplet onto a cell culture dish.
   2. Place a coverslip over the sample and observe under a tissue culture microscope at 10X and 20X magnifications.
      NOTE: The preparation should appear as a mix of single cells, small clusters, large clusters, and occasional mostly-intact embryo body.
   3. Visually assess the proportion of the preparation that is single cells and small clusters.  
      NOTE: Over-digestion will reduce viability; under-digestion will reduce single cell yield.
10. Collect the cell preparation by centrifugation at 300 x g for 5 min. Discard the supernatant and re-suspend in 1 ml cold FACS Buffer.
11. Moisten a 40 µm cell strainer with FACS Buffer. Pass the cell suspension through the 40 µm cell strainer onto a 35 mm cell culture dish. Wash the cell strainer one time with 1 ml FACS sorting buffer.
12. Transfer the flow-through to a 2 ml microcentrifuge tube. Collect cells by centrifugation at 300 x g for 5 min and re-suspend in 100 µl FACS Buffer.
13. Count cell yield using a hemocytometer. Further dilute samples to 5×10⁶ cells per ml, or optimal concentration recommended for FACS machine of choice. Optional-When counting cell yield on the hemocytometer, counterstain dead cells with trypan blue to confirm viability of the cell preparation prior to FACS sorting.  
    NOTE: Preparations that are too dilute will take excess amount of time to sort. Preparations that are too concentrated tend to clump into multimers and are suboptimal for sorting a pure population.
14. Reserve 10-20% of each sample to use as unstained controls. For remaining samples, stain dead cells by adding a fluorescent live/dead (L/D) discrimination dye into each sample. DO NOT wash.  
    NOTE: Any fluorescent dye that penetrates cells with compromised cell membranes and stains nucleic acids may be used as the L/D dye, provided that the fluorescence spectra is compatible with the FACS machine of choice and fluorophore labeling cells of interest.
    1. Do not wash the preparation after adding the dye as some cells will die in transit to FACS purification and should be excluded from the sorted population.
15. Moisten the 35 µm cell strainer capped FACS tube with 20 µl FACS buffer. Add cells to the strainer and collect by gravity. Store on ice.

4. FACS Enrichment

1. Use FACS to enrich for single, live cells expressing the fluorescent marker.
   1. Set a gate to distinguish cells from debris on a scatter plot of forward scatter (FSC-A) amplitude vs side scatter amplitude (SSC-A), both with linear scaling.
      Caution! Zebrafish cells are typically smaller than mouse or human cells. This is reflected in a lower basal separation between cells and debris.
   2. From the gate set in 4.1.1, set a gate to enrich for single cells and exclude multimers using a scatter plot of forward scatter height (FSC-H) and low side scatter height (SSC-H), both with linear scaling.
      NOTE: Cells with disproportionately high FSC-H and low SSC-H are likely multimers and are excluded from sorting.
   3. From the gate set in 4.1.2, using single color controls, set a gate to include only live cells using a scatter plot of FSA-A with linear scaling and amplitude of the channel used to detect L/D stain with log scaling. Include L/D negative cells.
   4. From the gate set in 4.1.3, using single color controls, set a gate to include only live cells with positive fluorescence using a scatter plot of FSA-A with linear scaling and amplitude of the channel used to detect fluorescent protein cell marker with log scaling. Include positive cells.
   5. Set any compensation controls, if necessary, to account for interference between the L/D stain and fluorescent protein spectra.
2. Set sort logic to cells that fall into all of the gates set in step 4.1.
   1. Using double labeled cells, verify gating for cell sorting.  
      NOTE: Cells for sorting will be cells rather than debris, single cells rather than multimers, L/D negative and fluorescence positive cells.
3. Sort 2,000-4,000 cells from the population of interest into 5 µl of cold FACS Buffer in a microcentrifuge tube on ice. To minimize shearing and strain on cells during sorting, use the lowest pressures possible for the cell sorter.  
   NOTE: The 5 µl droplet of FACS Buffer serves as a cushion for cells exiting the FACS machine. Collecting 2,000-4,000 cells directly into FACS Buffer eliminates the need for centrifugation prior to loading onto the IFC chips. This strategy is recommended because centrifugation can lead to formation of multimers if cells adhere to one another in the pellet.
4. Assess the post-sort viability analysis.
   1. Transfer 1 µl sorted cells to a fresh FACS tube with 100 µl FACS sorting buffer and 1:1,000 dilution of L/D stain. Pass the cells through the FACS sorter with the gating strategy in Step 4.1. Use the proportion of L/D negative to positive to estimate viability of sorted cells.

5. Load Cells onto Microfluidics Chip

1. Using a hemocytometer, measure both the concentration and size of sorted cells.
   1. Dilute sorted cells to at least 10 µl. Dilute an aliquot of 5 µl cells with 5 µl trypan blue. Load onto hemocytometer and apply coverslip.
   2. Collect bright field images of all cells in 4 x 4 grids of hemocytometer. Count the number of live cells and calculate live cells/ml. Live cells do not take up trypan blue.  
      NOTE: Use any standard image analysis software to measure the diameter of all live cells. Calculate the average and standard deviation of the cell size, and select an IFC plate suitable for the cell size range of interest. If cell size range straddles an IFC plate cell size range, use multiple plates to capture the full range of cells of interest.
2. Load cells on to IFC plate according to manufacturer's instructions (see Materials).
   1. Dilute cells to 1×10⁶ cells/ml and perform buoyancy optimization as per manufacturer's instructions.  
      NOTE: Buoyancy optimization ensures that cells neither sink to the bottom nor float to the top of the loading well for optimal loading onto IFC chip. The ratio of buffer to cells may vary by cell type, but typically ranges 6:4-7:3 of cells:buffer.
   2. Add cells to primed-IFC plate. Load plate into compatible fluidics machine, and run a cell loading script to push cells through microfluidics circuit and into capture lanes.
3. Confirm that cells are lodged in capture sites on IFC plate.
   1. Remove IFC plate from fluidics machine. Mount the IFC plate on a microscope equipped with a plate adaptor.
   2. Collect snapshots of bright field and fluorescence for each capture site at 10X magnification.  
      NOTE: Brightfield capture times may range from 10-50 ms, depending on lamp intensity. Fluorescence capture times may range from 250-750 ms, depending on fluorophore brightness. Avoid overexposing cells to prevent photodamage.

6. cDNA Synthesis

1. Perform cell lysis in situ according to IFC plate manufacturer's instructions.
   1. Add lysis buffers to wells on IFC plate. Load plate onto compatible fluidics machine and run cell lysis script as per manufacturer's instructions.
2. Perform reverse transcription according to IFC plate manufacturer's instructions.
   1. Add reverse transcription reagents to IFC plate. Load plate onto compatible fluidics machine. Run reverse transcription script. Sample cycling conditions: 1 cycle at 25°C for 10 min, 1 cycle at 42°C for 1 hr, then 1 cycle at 85°C for 5 min.
3. Perform pre-amplification with gene-specific probes according to IFC plate manufacturer's instructions.
   NOTE: Due to their greater specificity with low-copy numbers, use fluorogenic-labeled probes rather than probes optimized for use with intercalator dyes.
   1. Pool primers and dilute to final 180 nM each. Add pooled primers to IFC plate. Load plate onto compatible fluidics machine. Run preamplification script.
4. Perform pre-amplification with the following cycle conditions: 1 cycle at 95°C for 10 min, then 18 cycles with denaturing at 95°C for 15 sec then annealing and extension at 60°C for 4 min. Store pre-amplified cDNA at 4°C until harvesting. Harvest cDNA from microfluidic plate according to manufacturer's instructions and store at -20˚C until use.

7. Select cDNA from Single Cells for Single Target qRT-PCR

1. Dilute cDNA in 25 µl dilution reagent according to manufacturer's instructions.  
   NOTE: Approximate final yield is 28 µl of pre-amplified cDNA per cell. Reserve an aliquot of diluent for use as a no-template control in qRT-PCR.
   1. Referring to bright field and fluorescent images recorded in step 5.3.2, score each capture site. Manually count and record the number of cells. Assess and record whether each cell is fluorescent.  
      NOTE: Each individual site may contain 0, 1, or >1 cell, where >1 cell includes capture sites that contain multiple cells and/or unidentifiable debris. If the images are sufficient quality, a pixel value may also be assigned to quantify the intensity of the fluorescence signal.
2. Select samples for qRT-PCR analysis.
   1. Select cDNA samples from capture sites identified in step 7.2 that contain exact 1 cell with positive fluorescence. Pool 1 µl aliquots of cDNA from each sample.
      NOTE: This is the "Pool" positive control used to determine whether genes of interest are expressed in any of the selected cells.
   2. Select cDNA from at least one capture site that contains exactly 0 cells as a negative control. Use diluent from step 7.1.2 as a no-template control.
3. Run qRT-PCR assay.
   1. Use only probes used for pre-amplification in step 6.4.1. Load all samples in triplicate. Cycling conditions for a 10 µl reaction on a 384 well plate: 1 cycle 50°C 2 min, 1 cycle 95°C for 10 min, 40 cycles with denaturing at 95°C 15 sec then annealing and extension at 60°C for 1 min.

8. Data Analysis

1. Validate qRT-PCR products by standard gel electrophoresis. Confirm product is a single band at expected molecular weight (varies for each probe).  
   NOTE: If a probe produces multiple bands or a single band at an inappropriate size, then specificity is questionable, and this excludes the gene from analysis.
2. Examine all amplification curves and account for any abnormalities¹⁴. Calculate the average CT value for each sample/gene combination¹⁵.  
   NOTE: CT values for positive controls typically range from 10-30. CT values for negative controls typically range from 35-40 (no amplification). CT values 30-35 are considered very low expression and should be interpreted with caution¹⁴.
3. Report data
   1. If samples fall in the CT=10-30, data may be also reported as fold change in transcript abundance relative to a control sample.  
      NOTE: Fold change equals 2^^(-ΔΔCT) where ΔCT is relative to a housekeeping gene by subtracting the average CT of the sample from the housekeeping gene and ΔΔCT is the differences in ΔCT between the sample and a positive control¹⁵.