We thank Dr. C. Geoffrey Burns for fish stock. The authors are grateful to UNC Flow Cytometry Core Facility, UNC-CGIBD AAC core for resources enabling this project, and the ZAC facility for animal care. L.S. is supported by NIH T32 grant HL069768-13 (PI, Nobuyo Maeda). N.F. is supported by NSF Graduate Research Fellowship NSF-DGE-1144081. This study was supported by NIH P30DK034987 grant (to UNC Advanced Analytics Core), American Heart Association Scientist Development Grant 13SDG17060010 and Ellison Medical Foundation New Scholar Grant AG-NS-1064-13 (to Dr. Qian), and NIH R00 HL109079 grant (to Dr. Liu).

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
<ol>
	<li>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.</li>
	<li>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. &#160; 
	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 &#60;1.0% of the total cells at 24 hpf (hours post-fertilization).</li>
	<li>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.</li>
	<li>Collect staged embryos.
	<ol>
		<li>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.</li>
		<li>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 &#176;C with air circulation.</li>
	</ol>
	</li>
	<li>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&#176;C. &#160; 
	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.</li>
</ol>
2. Set Up for Single Cell Dissociation
<ol>
	<li>Approximately 30 min prior to the time point of interest (18 hpf) remove embryos from their chorion manually with fine forceps.</li>
	<li>Collect and label the following for each condition: two 2 ml microcentrifuge tubes, one 40 &#181;m cell strainer, one 35 mm cell culture dish, and two FACS tubes topped with a 35 &#181;m cell strainer.</li>
	<li>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&#39;s L-15 media supplemented with 5% heat-inactivated fetal bovine serum.</li>
	<li>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.</li>
</ol>
3. Single Cell Dissociation
<ol>
	<li>Using a wide bore glass pipette, transfer 100-300 embryos to a 2 ml micro centrifuge tube in a minimum volume of Egg Water.</li>
	<li>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 &#60;3 days post-fertilization are bleached after they are chilled, which is not appropriate for obtaining viable cells.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Collect tissue by centrifugation at 300 x g for 3 min. Remove supernatant and resuspend in 1 ml Cell Dissociation Reagent 2.</li>
	<li>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.
	<ol>
		<li>To assess digestion progress, dilute 2 &#181;l supernatant into 18 &#181;l FACS Buffer and pipette as a droplet onto a cell culture dish.</li>
		<li>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.</li>
		<li>Visually assess the proportion of the preparation that is single cells and small clusters. &#160; 
		NOTE: Over-digestion will reduce viability; under-digestion will reduce single cell yield.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Moisten a 40 &#181;m cell strainer with FACS Buffer. Pass the cell suspension through the 40 &#181;m cell strainer onto a 35 mm cell culture dish. Wash the cell strainer one time with 1 ml FACS sorting buffer.</li>
	<li>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 &#181;l FACS Buffer.</li>
	<li>Count cell yield using a hemocytometer. Further dilute samples to 5x106 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. &#160; 
	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.</li>
	<li>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. &#160; 
	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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Moisten the 35 &#181;m cell strainer capped FACS tube with 20 &#181;l FACS buffer. Add cells to the strainer and collect by gravity. Store on ice.</li>
</ol>
4. FACS Enrichment
<ol>
	<li>Use FACS to enrich for single, live cells expressing the fluorescent marker.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Set any compensation controls, if necessary, to account for interference between the L/D stain and fluorescent protein spectra.</li>
	</ol>
	</li>
	<li>Set sort logic to cells that fall into all of the gates set in step 4.1.
	<ol>
		<li>Using double labeled cells, verify gating for cell sorting. &#160; 
		NOTE: Cells for sorting will be cells rather than debris, single cells rather than multimers, L/D negative and fluorescence positive cells.</li>
	</ol>
	</li>
	<li>Sort 2,000-4,000 cells from the population of interest into 5 &#181;l 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. &#160; 
	NOTE: The 5 &#181;l 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.</li>
	<li>Assess the post-sort viability analysis.
	<ol>
		<li>Transfer 1 &#181;l sorted cells to a fresh FACS tube with 100 &#181;l 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.</li>
	</ol>
	</li>
</ol>
5. Load Cells onto Microfluidics Chip
<ol>
	<li>Using a hemocytometer, measure both the concentration and size of sorted cells.
	<ol>
		<li>Dilute sorted cells to at least 10 &#181;l. Dilute an aliquot of 5 &#181;l cells with 5 &#181;l trypan blue. Load onto hemocytometer and apply coverslip.</li>
		<li>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. &#160; 
		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.</li>
	</ol>
	</li>
	<li>Load cells on to IFC plate according to manufacturer&#39;s instructions (see Materials).
	<ol>
		<li>Dilute cells to 1x106 cells/ml and perform buoyancy optimization as per manufacturer&#39;s instructions. &#160; 
		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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Confirm that cells are lodged in capture sites on IFC plate.
	<ol>
		<li>Remove IFC plate from fluidics machine. Mount the IFC plate on a microscope equipped with a plate adaptor.</li>
		<li>Collect snapshots of bright field and fluorescence for each capture site at 10X magnification. &#160; 
		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.</li>
	</ol>
	</li>
</ol>
6. cDNA Synthesis
<ol>
	<li>Perform cell lysis in situ according to IFC plate manufacturer&#39;s instructions.
	<ol>
		<li>Add lysis buffers to wells on IFC plate. Load plate onto compatible fluidics machine and run cell lysis script as per manufacturer&#39;s instructions.</li>
	</ol>
	</li>
	<li>Perform reverse transcription according to IFC plate manufacturer&#39;s instructions.
	<ol>
		<li>Add reverse transcription reagents to IFC plate. Load plate onto compatible fluidics machine. Run reverse transcription script. Sample cycling conditions: 1 cycle at 25&#176;C for 10 min, 1 cycle at 42&#176;C for 1 hr, then 1 cycle at 85&#176;C for 5 min.</li>
	</ol>
	</li>
	<li>Perform pre-amplification with gene-specific probes according to IFC plate manufacturer&#39;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.
	<ol>
		<li>Pool primers and dilute to final 180 nM each. Add pooled primers to IFC plate. Load plate onto compatible fluidics machine. Run preamplification script.</li>
	</ol>
	</li>
	<li>Perform pre-amplification with the following cycle conditions: 1 cycle at 95&#176;C for 10 min, then 18 cycles with denaturing at 95&#176;C for 15 sec then annealing and extension at 60&#176;C for 4 min. Store pre-amplified cDNA at 4&#176;C until harvesting. Harvest cDNA from microfluidic plate according to manufacturer&#39;s instructions and store at -20&#730;C until use.</li>
</ol>
7. Select cDNA from Single Cells for Single Target qRT-PCR
<ol>
	<li>Dilute cDNA in 25 &#181;l dilution reagent according to manufacturer&#39;s instructions. &#160; 
	NOTE: Approximate final yield is 28 &#181;l of pre-amplified cDNA per cell. Reserve an aliquot of diluent for use as a no-template control in qRT-PCR.
	<ol>
		<li>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. &#160; 
		NOTE: Each individual site may contain 0, 1, or &#62;1 cell, where &#62;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.</li>
	</ol>
	</li>
	<li>Select samples for qRT-PCR analysis.
	<ol>
		<li>Select cDNA samples from capture sites identified in step 7.2 that contain exact 1 cell with positive fluorescence. Pool 1 &#181;l aliquots of cDNA from each sample. 
		NOTE: This is the &#34;Pool&#34; positive control used to determine whether genes of interest are expressed in any of the selected cells.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Run qRT-PCR assay.
	<ol>
		<li>Use only probes used for pre-amplification in step 6.4.1. Load all samples in triplicate. Cycling conditions for a 10 &#181;l reaction on a 384 well plate: 1 cycle 50&#176;C 2 min, 1 cycle 95&#176;C for 10 min, 40 cycles with denaturing at 95&#176;C 15 sec then annealing and extension at 60&#176;C for 1 min.</li>
	</ol>
	</li>
</ol>
8. Data Analysis
<ol>
	<li>Validate qRT-PCR products by standard gel electrophoresis. Confirm product is a single band at expected molecular weight (varies for each probe). &#160; 
	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.</li>
	<li>Examine all amplification curves and account for any abnormalities14. Calculate the average CT value for each sample/gene combination15. &#160; 
	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.</li>
	<li>Report data
	<ol>
		<li>If samples fall in the CT=10-30, data may be also reported as fold change in transcript abundance relative to a control sample.&#160;&#160; 
		NOTE: Fold change equals 2^(-&#916;&#916;CT) where &#916;CT is relative to a housekeeping gene by subtracting the average CT of the sample from the housekeeping gene and &#916;&#916;CT is the differences in &#916;CT between the sample and a positive control15.</li>
	</ol>
	</li>
</ol>

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</ol>

The authors have nothing to disclose.

The method described herein uses expression of a fluorescent protein under control of a cell-type specific promoter to enrich a population of cardiac progenitor cells from zebrafish embryos for use in microfluidic assisted single cell capture system to assess expression of a subset of cardiac genes in single cells. Provided that FACS laser excitation and emission capabilities are compatible with the fluorophore(s) of choice, this method can be used for any fluorescent reporter line. Many zebrafish reporter lines are alre...

Most current studies of cell and molecular biology are based on population averages. However, important biological events may be masked by these traditional population-based analyses since minor populations can play major roles in biological processes and disease outcome. Understanding gene expression in heterogeneous populations at the single cell level can (and has) lead to relevant biological and clinical insights1,2. Of concern to embryonic development studies, in a larger population of cells, progenitor cells are often underrepresented, making it challenging to detect subtle changes in gene expression that ultimately initiate cell fate decisions3. Similarly, a single cell type may have different expression profiles in response to the microenvironment4. For example, resident endothelial cells in the same organ or in different organs (e.g., aorta or kidney) exhibit significant heterogeneity despite sharing common morphological and functional features5. In addition, cancer cells populating the same tumor can also have varying molecular profiles or mutations at the single cell level6.
In model systems, transcriptomics in single cells has successfully identified new cell populations, characterized intermediate states that occur during cell differentiation, and revealed differential cellular responses to stimuli7,8,9. Such insights would have been masked in conventional population-based studies. Zebrafish embryos are a tremendously under-utilized source of stem, progenitor, and differentiating cells for exploring questions of single cell heterogeneity and molecular regulation of cellular identities during development. Their highly stereotyped, ex vivo development and ease of genetic manipulation make them an excellent model system for this approach10,11. Specifically, a major limitation to interpretation of single cell gene expression data is that reliable identification of novel intermediate cell states during development requires very careful timing of tissue collection9. This is necessary to ensure that heterogeneity between captured cells represents heterogeneity within a tissue at a single time point rather than heterogeneity in gene expression presented by age-dependent cell differentiation. Compared to mice, zebrafish embryo development may be precisely synchronized across a large number of embryos12. Additionally, with large clutch sizes, zebrafish embryos can be used as an abundant source of stem and progenitor cells.
This protocol describes a method to isolate cells from zebrafish embryos and capture single cells using a commercially available integrated microfluidics circuit (IFC) chip and autoprep system for qRT-PCR gene expression analysis. This protocol can be rapidly transferrable to any high throughput multiplexing assays including whole transcriptome sequencing that allows more comprehensive analysis of cellular heterogeneity13. It also offers several advantages to traditional gene expression assays. The single cell isolation protocol yields high viability after FACS, which decreases the proportion of compromised cells that are included in downstream applications. By using an IFC, captured cells may be directly observed to evaluate capture rates and assess cell health morphologically. In addition, this protocol is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidic cell capture technologies.
As proof of principle, single cells derived from cardiac progenitors were isolated and captured on an IFC chip, and then the relative abundance of cardiac differentiation markers was measured by qRT-PCR. Gene expression analysis at the single cell level demonstrates that cardiac progenitors coexist with their differentiating progeny. The insight gained from single-cell profiling of cardiac progenitors may shed light on the heterogeneity in gene expression patterns among cardiac progenitor cells during vertebrate development, which may have been masked in traditional population-based analyses.

<table><tbody><tr><td>&lt;strong&gt;Supplies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Dumont #5 forceps</td><td>Fine Science Tools</td><td>11254-20</td><td>For removing the chorion from embryos</td></tr><tr><td>Microcentrifuge tube 2 ml</td><td>GeneMate</td><td>C-3261-1</td><td></td></tr><tr><td>40 um cell strainer</td><td>Biobasic</td><td>SP104151</td><td></td></tr><tr><td>35 mm culture dish</td><td>Falcon</td><td>351008</td><td></td></tr><tr><td>FACS tubes topped with 35 um cell strainer</td><td>Falcon</td><td>352235</td><td></td></tr><tr><td>P1000 and tips</td><td>Rainin</td><td>17005089</td><td></td></tr><tr><td>P20 and tips</td><td>Rainin</td><td>17005091</td><td></td></tr><tr><td>IFC chip manufacturer&#039;s protocol</td><td>Fluidigm</td><td>100-6117</td><td>Version 100-6117 E1 was used in representative experiment</td></tr><tr><td>Wide bore pastuer glass pippette</td><td>VWR</td><td>14673-010</td><td>For transferring embryos</td></tr><tr><td>Adult wild type zebrafish</td><td>N/A</td><td></td><td>We used AB line</td></tr><tr><td>Adult transgenic zebrafish</td><td>N/A</td><td></td><td>We used Tg(nkx2.5:ZsYellow)</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Reagents for cell dissociation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Double distilled water</td><td>N/A</td><td></td><td></td></tr><tr><td>Instant Ocean Sea Salt</td><td>Instant Ocean</td><td>SS15-10</td><td></td></tr><tr><td>NaCl</td><td>FisherScientific</td><td>S271-3</td><td>make stock in water and use for de-yolking buffer</td></tr><tr><td>KCl</td><td>Sigma Aldrich</td><td>P5405</td><td>make stock in water and use for de-yolking buffer</td></tr><tr><td>NaHCO3</td><td>Sigma Aldrich</td><td>S6014</td><td>make stock in water and use for de-yolking buffer</td></tr><tr><td>Leibovitz&#039;s L-15</td><td>Gibco</td><td>21083-027</td><td></td></tr><tr><td>FBS</td><td>FisherScientific</td><td>03-600-511</td><td>Heat inactivate; any brand of FBS should be fine</td></tr><tr><td>Cell Dissociation Reagent 1 -TrypLE</td><td>Life Technologies</td><td>12605-010</td><td>Store at room temperature.</td></tr><tr><td>Cell Dissociation Reagent 2 - FACSmax</td><td>Genlantis</td><td>T200100</td><td>Store -20; thaw on ice; bring to room temp before use</td></tr><tr><td>pronase (optional)</td><td>Sigma</td><td>P5147</td><td></td></tr><tr><td>L/D Dye - Sytox Blue</td><td>Life Technologies</td><td>S34857</td><td>Any live/dead stain suitable for flow cytometry will work</td></tr><tr><td>Trypan Blue</td><td>Gibco</td><td>15250-061</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Reagents for IFC plate use and qRT-PCR&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Gene-specific probes</td><td></td><td></td><td>Probes will vary by experiment</td></tr><tr><td>TaqMan Probe ef1a</td><td>Life Technologies</td><td>Dr03432748_m1</td><td></td></tr><tr><td>TaqMan Probe gata4</td><td>Life Technologies</td><td>Dr03443262_g1</td><td></td></tr><tr><td>TaqMan Probe nk2-5</td><td>Life Technologies</td><td>Dr03074126_m1</td><td></td></tr><tr><td>TaqMan Probe myl7</td><td>Life Technologies</td><td>Dr03105700_m1</td><td></td></tr><tr><td>TaqMan Probe vmhc</td><td>Life Technologies</td><td>Dr03431136_m1</td><td></td></tr><tr><td>TaqMan Probe isl1</td><td>Life Technologies</td><td>Dr03425734_m1</td><td></td></tr><tr><td>TaqMan Gene Expression Master Mix</td><td>Life Technologies</td><td>4369016</td><td></td></tr><tr><td>Reagents listed in IFC manufacturer&#039;s protocol</td><td></td><td></td><td></td></tr><tr><td>C1 Reagent Kit</td><td>Fluidigm</td><td>100-5319</td><td>Reagents for loading cells onto IFC plate</td></tr><tr><td>Ambion Single Cell-to-CT&lt;sup&gt;TM&lt;/sup&gt;</td><td>Life Technologies</td><td>4458237</td><td>Reagents for reverse transcription and pre-amplification steps</td></tr><tr><td>Molecular Biology Quality Water</td><td>Corning</td><td>46-000CM</td><td></td></tr><tr><td>C1 IFC for PreAmp (5-10 um)</td><td>Fluidigm</td><td>100-5757</td><td>IFC plate for small cells</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>C1 AutoPrep machine</td><td>Fluidigm</td><td>100-5477</td><td>For IFC plate use</td></tr><tr><td>Hemocytometer&amp;nbsp;</td><td>Sigma Aldrich</td><td>Z359629</td><td>For counting cells and assessing cell size</td></tr><tr><td>Dissecting microscope</td><td></td><td></td><td>For removing embryos from chorion</td></tr><tr><td>Tissue culture microscope</td><td></td><td></td><td>For assessing single cell digestion</td></tr><tr><td>FACS machine</td><td></td><td></td><td>For isolating cells of interest</td></tr></tbody></table>

isolation and characterization of single cells from zebrafish embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

As proof of principle, gene expression was assessed to explore differentiation dynamics during cardiac development. In zebrafish, cardiac progenitors arise from a mesodermal population of cells that migrate to the anterior lateral plate mesoderm where they fuse to form the linear heart tube. Prior to fusion, cardiac progenitors begin to express the transcription factor nkx2.5 (NK2 homeobox 5), which is thought to be the earliest specific marker of cardiac progenitors16,17</s...

Watch this Scientific Journal Video about Isolation and Characterization of Single Cells from Zebrafish Embryos at JoVE.com

Isolation and Characterization of Single Cells from Zebrafish Embryos

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

isolation-and-characterization-of-single-cells-from-zebrafish-embryos

Research

JoVE Journal

Developmental Biology

14.5K Views.  University of North Carolina at Chapel Hill. This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells. 

Video: Isolation and Characterization of Single Cells from Zebrafish Embryos

Labeling and Imaging Cells in the Zebrafish Hindbrain

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish embryos, injection of plasmid DNA results in mosaic expression, allowing for the visualization of single cells or small clusters of cells 1 . We describe how injection of plasmid DNA encoding membrane-targeted Green Fluorescent Protein (mGFP) under the control of a ubiquitous promoter can be used for imaging cells undergoing neurulation. Central to this protocol is the methodology for imaging labeled cells at high resolution in sections and also in real time. This protocol entails the injection of mGFP DNA into young zebrafish embryos. Embryos are then processed for vibratome sectioning, antibody labeling and imaging with a confocal microscope. Alternatively, live embryos expressing mGFP can be imaged using time-lapse confocal microscopy. We have previously used this straightforward approach to analyze the cellular behaviors that drive neural tube formation in the hindbrain region of zebrafish embryos 2. The fixed preparations allowed for unprecedented visualization of cell shapes and organization in the neural tube while live imaging complemented this approach enabling a better understanding of the cellular dynamics that take place during neurulation.

Key to understanding the morphogenetic processes that shape the early embryo is the ability to image cells at high resolution. We describe here a technique for labeling single cells or small clusters of cells in whole zebrafish embryos with membrane-targeted Green Fluorescent Protein.

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish ...

Neuroscience

Sample Preparation and Analysis of RNASeq-based Gene Expression Data from Zebrafish

The analysis of global gene expression changes is a valuable tool for identifying novel pathways underlying observed phenotypes. The zebrafish is an excellent model for rapid assessment of whole transcriptome from whole animal or individual cell populations due to the ease of isolation of RNA from large numbers of animals. Here a protocol for global gene expression analysis in zebrafish embryos using RNA sequencing (RNASeq) is presented. We describe preparation of RNA from whole embryos or from cell populations obtained using cell sorting in transgenic animals. We also describe an approach for analysis of RNASeq data to identify enriched pathways and Gene Ontology (GO) terms in global gene expression data sets. Finally, we provide a protocol for validation of gene expression changes using quantitative reverse transcriptase PCR (qRT-PCR). These protocols can be used for comparative analysis of control and experimental sets of zebrafish to identify novel gene expression changes, and provide molecular insight into phenotypes of interest.

This protocol presents an approach for whole transcriptome analysis from zebrafish embryos, larvae, or sorted cells. We include isolation of RNA, pathway analysis of RNASeq data, and qRT-PCR-based validation of gene expression changes.

The analysis of global gene expression changes is a valuable tool for identifying novel pathways underlying observed phenotypes. The zebrafish is an ...

Genetics

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Biology

Culture and Transfection of Zebrafish Primary Cells

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in an intact and developing vertebrate. However, the detailed imaging of the morphologies of distinct cell types and subcellular structures is limited in whole mounts. Therefore, we established an efficient and easy-to-use protocol to culture live primary cells from zebrafish embryos and adult tissue.
In brief, 2 dpf zebrafish embryos are dechorionated, deyolked, sterilized, and dissociated to single cells with collagenase. After a filtration step, primary cells are plated onto glass bottom dishes and cultivated for several days. Fresh cultures, as much as long term differenciated ones, can be used for high resolution confocal imaging studies. The culture contains different cell types, with striated myocytes and neurons being prominent on poly-L-lysine coating. To specifically label subcellular structures by fluorescent marker proteins, we also established an electroporation protocol which allows the transfection of plasmid DNA into different cell types, including neurons. Thus, in the presence of operator defined stimuli, complex cell behavior, and intracellular dynamics of primary zebrafish cells can be assessed with high spatial and temporal resolution. In addition, by using adult zebrafish brain, we demonstrate that the described dissociation technique, as well as the basic culturing conditions, also work for adult zebrafish tissue.

We present an efficient and easy-to-use protocol for preparing primary cell cultures of zebrafish embryos for transfection and live cell imaging as well as a protocol to prepare primary cells from adult zebrafish brain.

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in ...

Preparation of Single-Somite Explants from Zebrafish Embryos

The body axis of vertebrate embryos is periodically subdivided into 3D multicellular units called somites. While genetic oscillations and molecular prepatterns determine the initial length-scale of somites, mechanical processes have been implicated in setting their final size and shape. To better understand the intrinsic material properties of somites, a method is developed to culture single-somite explant from zebrafish embryos. Single somites are isolated by first removing the skin of embryos, followed by yolk removal and sequential excision of neighboring tissues. Using transgenic embryos, the distribution of various sub-cellular structures can be observed by fluorescent time-lapse microscopy. Dynamics of explanted somites can be followed for several hours, thus providing an experimental framework for studying tissue-scale shape changes at single-cell resolution. This approach enables direct mechanical manipulation of somites, allowing for dissection of the material properties of the tissue. Finally, the technique outlined here can be readily extended for explanting other tissues such as the notochord, neural plate, and lateral plate mesoderm.

We present a protocol for isolating single somites from zebrafish embryos, the dynamics of which can be followed in culture for several hours by fluorescence time-lapse microscopy, thus providing a methodology to quantify tissue-scale shape changes at single-cell resolution.

The body axis of vertebrate embryos is periodically subdivided into 3D multicellular units called somites. While genetic oscillations and molecular ...

Processing Gut Cells from Zebrafish Larvae for Single-Cell RNA Sequencing

Gut Isolation from Zebrafish Larvae for Single-cell RNA Sequencing

The gastrointestinal (GI) tract performs a range of functions essential for life. Congenital defects affecting its development can lead to enteric neuromuscular disorders, highlighting the importance to understand the molecular mechanisms underlying GI development and dysfunction. In this study, we present a method for gut isolation from zebrafish larvae at 5 days post fertilization to obtain live,&#160;viable cells which can be used for single-cell RNA sequencing (scRNA-seq) analysis. This protocol is based on the manual dissection of the zebrafish intestine, followed by enzymatic dissociation with papain. Subsequently, cells are submitted to fluorescence-activated cell sorting, and viable cells are collected for scRNA-seq. With this method, we were able to successfully identify different intestinal cell types, including epithelial, stromal, blood, muscle, and immune cells, as well as enteric neurons and glia. Therefore, we consider it to be a valuable resource for studying the composition of the GI tract in health and disease, using the zebrafish.

Author Spotlight: Isolating and Analyzing Intestinal Cells of Zebrafish Larvae for Investigating Transcriptomic Aspects of Gastrointestinal Development 

Here, we describe a method for gut isolation from zebrafish larvae at 5 days post fertilization, for single-cell RNA sequencing analysis.

The gastrointestinal (GI) tract performs a range of functions essential for life. Congenital defects affecting its development can lead to enteric ...

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression both in terms of tissue distribution and developmental stage. Protocols are described for the use of WISH of zebrafish embryos using antisense RNA probes labeled with digoxigenin. Probes are generated by incorporating digoxigenin-linked nucleotides through in vitro transcription of gene templates that have been cloned and linearized. The chorions of embryos harvested at defined developmental stages are removed before incubation with specific probes. Following a washing procedure to remove excess probe, embryos are incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase. By employing a chromogenic substrate for alkaline phosphatase, specific gene expression can be assessed. Depending on the level of gene expression the entire procedure can be completed within 2-3 days.

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

The zebrafish, a small tropical fish, has become a popular model for studying gene function during vertebrate development and disease. The temporal and spatial expression of target genes can be determined by in situ hybridization. Our improved protocol allows for the detection of low abundant transcripts with low non-specific background signal.

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression ...

Sequential Immunofluorescence and Immunohistochemistry on Cryosectioned Zebrafish Embryos

Investigation of intercellular interactions often requires discrete labeling of specific cell populations and precise protein localization. The zebrafish embryo is an excellent tool for examining such interactions with an in vivo model. Whole-mount immunohistochemical and immunofluorescence assays are frequently applied in zebrafish embryos to assess protein expression. However, it can be difficult to achieve accurate mapping of co-localized proteins in three-dimensional space. In addition, some studies may require the use of two antibodies that are not compatible with the same technique (e.g., antibody 1 is only suitable for immunohistochemistry and antibody 2 is only suitable for immunofluorescence). The purpose of the method described herein is to perform sequential immunofluorescence and/or immunohistochemistry on individual cryosections derived from early-stage zebrafish embryos. Here we describe the use of sequential rounds of immunofluorescence, imaging, immunohistochemistry, imaging for a single cryosection in order to achieve precise identification of protein expression at the single-cell level. This methodology is suitable for any study in early-stage zebrafish embryos that requires accurate identification of multiple protein targets in individual cells.

This protocol demonstrates sequential immunofluorescence and immunohistochemistry on cryosections from early-stage zebrafish embryos enabling precise colocalization analyses in specific cell populations.

Investigation of intercellular interactions often requires discrete labeling of specific cell populations and precise protein localization. The zebrafish ...

Isolation of Cardiac and Vascular Smooth Muscle Cells from Adult, Juvenile, Larval and Embryonic Zebrafish for Electrophysiological Studies

Zebrafish have long been used as a model vertebrate organism in cardiovascular research. The technical difficulties of isolating individual cells from the zebrafish cardiovascular tissues have been limiting in studying their electrophysiological properties. Previous methods have been described for dissection of zebrafish hearts and isolation of ventricular cardiac myocytes. However, the isolation of zebrafish atrial and vascular myocytes for electrophysiological characterization was not detailed. This work describes new and modified enzymatic protocols that routinely provide isolated juvenile and adult zebrafish ventricular and atrial cardiomyocytes, as well as vascular smooth muscle (VSM) cells from the bulbous arteriosus, suitable for patch-clamp experiments. There has been no literary evidence of electrophysiological studies on zebrafish cardiovascular tissues isolated at embryonic and larval stages of development. Partial dissociation techniques that allow patch-clamp experiments on individual cells from larval and embryonic hearts are demonstrated.

The present protocol describes the acute isolation of viable cardiac and vascular smooth muscle cells from adult, juvenile, larval, and embryonic zebrafish (Danio rerio), suitable for electrophysiological studies.

Zebrafish have long been used as a model vertebrate organism in cardiovascular research. The technical difficulties of isolating individual cells from the ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

Podsumowanie

Przeglądaj więcej filmów

Labeling and Imaging Cells in the Zebrafish Hindbrain

Sample Preparation and Analysis of RNASeq-based Gene Expression Data from Zebrafish

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

Culture and Transfection of Zebrafish Primary Cells

Preparation of Single-Somite Explants from Zebrafish Embryos

Gut Isolation from Zebrafish Larvae for Single-cell RNA Sequencing

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

Sequential Immunofluorescence and Immunohistochemistry on Cryosectioned Zebrafish Embryos

Isolation of Cardiac and Vascular Smooth Muscle Cells from Adult, Juvenile, Larval and Embryonic Zebrafish for Electrophysiological Studies