Isolation and Characterization of Single Cells from Zebrafish Embryos

The overall goal of this experiment is to isolate single cells from early stage zebrafish embryos for their capture in a microfluidic-based single-cell multiplex system to assess the gene expression of individual cells. This method can help answer key questions in the developmental biology field, such as what transcriptional changes do cells undergo during specification and differentiation. The main advantage of this technique is that it is an unbiased approach for revealing the heterogeneity that is masked in conventional population-based techniques.

Generally, individuals new to this technique will struggle because generating the viable preparations of single cells is technically challenging but essential for downstream analysis. Once mastered, processing the whole embryo samples into single lysed cells on an integrated microfluidic chip can be completed in six hours if steps are performed properly. While attempting this procedure, it's important to remember to make sure that only single cells are captured on the microfluidic device.

After breeding adult wild-type and transgenic zebrafish, collect embryos at 15-minute intervals according to facility and institution-approved standard operating procedures. At least two hours later, transfer 100 to 300 fertilized embryos from a single timepoint into a new Petri dish containing fresh egg water. And use fine forceps to manually remove the chorion from each embryo.

Next, use a wide bore glass pipette to transfer the embryos into a 2-milliliter microcentrifuge tube in a minimum volume of egg water. When all of the embryos have been transferred, replace the water with 1 milliliter of ice cold egg water and submerge the tube in ice for 20 minutes. After the euthanization, use a wide bore glass pipette to remove the supernatant.

And a P-1000 pipette to add fresh egg water to the embryos two times. After the second wash, replace the egg water with 1 milliliter of de-yolking buffer and use a P-1000 pipette to triturate the embryos eight to 12 times until the yolk is dissolved and only the bodies of the embryos are visible. Collect the tissue by centrifugation.

Then, use the pipette to gently remove the supernatant without disturbing the tissue pellet, and re-suspend the cells in 1 milliliter of fresh egg water. After washing the embryos two more times, re-suspend the pellet in 1 milliliter of room temperature cell dissociation reagent-1. Then place the tube on its side for a 10-minute incubation at room temperature.

With gentle trituration every two to three minutes to prevent clumping. At the end of the incubation, spin down the cells again and re-suspend the pellet in 1 milliliter of cell dissociation reagent-2. Incubate the cells horizontally at room temperature for another five to 15 minutes with gentle trituration every two to three minutes.

Every five minutes, dilute 2 microliters of supernatant in 18 microliters of FACS buffer, and dispense the cells as a droplet onto a cell culture dish. Place a coverslip over the sample and use a tissue culture microscope to assess the digestion progression at 10 and 20x magnifications. When the preparation appears to be a mix of primarily single cells with some small and large cell clusters and a few mostly intact embryo bodies, spin down the cells and re-suspend the pellet in 1 milliliter of cold FACS buffer.

Next, wet a 40-micron cell strainer with FACS buffer. And filter the cells through the strainer into a 35-millimeter cell culture dish. After spinning down the cells again, re-suspend the pellet in 100 microliters of FACS buffer, and count the number of viable cells by Trypan blue exclusion.

Next, dilute the sample to 5 times 10 to the 6th cells per milliliter. And reserve 10 to 20%of the cells for the unstained control. Stain the rest of the cells with a fluorescent live/dead discrimination dye.

Then wet a 35-micron cell strainer capped FACS tube with 20 microliters of FACS buffer, and add the cells to the strainer, collecting them by gravity. Using the illustrated gating strategy, sort the single live cells expressing the fluorescent marker. Verifying the gating with the double-labled cells.

Sort 2, 000 to 4, 000 cells from the population of interest into 5 microliters of cold FACS buffer in a microcentrifuge tube on ice. Then transfer 1 microliter of the sorted cells into a new FACS tube with 100 microliters of FACS sorting buffer and a 1 to 1, 000 dilution of live/dead stain. And run the cell sample using the sorting gates to assess the post sort viability.

Next, load 1 microliter of cells diluted in 9 microliters of FACS buffer onto a hemocytometer to determine the cell concentration and average diameter. Now select an integrated microfluidic circuit, or IFC chip, of the appropriate size for the average cell diameter of the sample, and prime the fluidics according to the manufacturer's instructions. Load the cells onto the plate according to the manufacturer's instructions, and load the plate into a compatible fluidic machine.

Then run an appropriate cell loading script for the IFC chip to push the cells into the capture lanes. To confirm that the cells are lodged in the capture sites, mount the plate on a microscope equipped with a plate adapter and obtain brightfield and fluorescence images of each capture site at a 10x magnification. The cells can then be processed for downstream assessment of their target gene expression by QRT-PCR.

At the 18-somite stage, fluorescence microscopy can be used to verify the ZsYellow expression of the embryos. After sorting, cells with a range of diameters are obtained. The IFC plate can be used to capture cells with a diameter of interest.

For example, in this experiment, cells five to 10 micrometers in diameter. As expected, the sorted and captured cells from 18-hour post-fertilization embryos expressed the ZsYellow fluorescent marker. Here, QRT-PCR was used to assess the relative gene expression of specific genes of interest in 40 single cells from cell capture sites on an IFC plate.

Since the range of cycle threshold values for the housekeeping gene was too broad for comparing gene expression between samples and many cycle threshold values exceeded 30, The values were visualized as a heat map. Comparing across samples, a substantial heterogeneity in gene expression was observed in this experiment. With cells classified as Types 1 through 5 based on their gene expression pattern.

Once mastered, processing the whole embryo samples into single lysed cells on an integrated microfluidic chip can be completed in six hours if the steps are performed properly. While attempting this procedure, it's important to remember that only single cells are captured on the microfluidic chip. Following this procedure, other methods like high-throughput single cell sequencing can be performed to fully characterize the transcriptomes of single cells.

After its development, this technique paved the way for researchers in the field of developmental biology for exploring cellular heterogeneity in early developing embryos. After watching this video, you should have a good understanding of how to isolate single cells from zebrafish embryos, capture single cells in a microfluidic base single-cell multiplex system and assess the gene expression of individual cells.