The overall goal of this procedure is to generate dispersed cultures for long-term imaging of oscillations in fluorescence reporters of cyclic gene expression in single isolated cells from the zebrafish segmentation clock. This is accomplished by first dissecting presi mesoderm or PSM tissue from transgenic zebrafish embryos. Next, the most posterior piece of the PSM, the tail bud is dissected from multiple embryos and the pieces are pooled in trypsin.
Then the tail bud pieces are manually dispersed with a pipette generating a cell suspension. Finally, the cell suspension is plated onto fibronectin coated culture dishes for long-term imaging. Ultimately, results can be obtained that show multiple oscillations in intensity over time from single PSM cells through fluorescence time-lapse microscopy.
The main advantage of this technique over existing methods, such as in C two hybridization of gene expression in PSM tissue, is that our fluorescence reporter allows for real time monitoring of the dynamics of oscillations in the segmentation clock at the level of single cells. This method can provide unique information to the segmentation clock, but it can also be used in other systems like organogenesis or disease models where it can give us information about behavior at single cell level, where the appropriate transgenic lines are available On the day before dissection. Obtain embryos from an in cross of zebrafish pears heterozygous for the transgenic allele.
Raise the embryos in E three medium without methylene blue at 28 degrees Celsius until shield stage about six hours post fertilization or HPF transfer the embryos in E three medium without methylene blue to 20 degrees celsius overnight at 20 degrees Celsius. The embryos will form one so mite per hour once they reach the tail bud stage at the beginning of the dissection protocol the following morning, the embryos should be at the five to eight, so might stage after assembling the tools for dissection. Cover a glass bottom imaging dish with fibronectin one substrate.
Leave the dish on the bench to coat during the dissection under a fluorescent stereo microscope. Identify transgenic embryos by examining the presi mesoderm or PSM. For YFP expression.
Fluorescence should be visible in the region from the last formed so mite to the tail bud. Select embryos with the brightest signal. 25%of offspring should be homozygous embryos carrying two copies of the transgene using transmitted light as a positional reference.
To distinguish any autofluorescence transfer positive embryos to a separate plastic Petri dish containing E three medium without methylene blue. To prepare embryos for dissection under a dissecting scope using fine forceps, carefully remove the corion from each embryo in the E three medium. Be sure not to damage the embryo or disrupt the oak cell.
Fill a cyl guard coated dissecting dish with L 15 medium with serum using forceps or another flat tool. Remove any air bubbles from the surface of the S guard using a fire polished glass pipette. Transfer the coated embryos to the dissection dish and with the wire tools, move all embryos to one side.
Next, in the small well made in the sard layer within the dissecting dish, orient a single embryo on its lateral side. Then using the micro scalpel slice through the embryo and yolk cell just anterior to the hind brain and through the ventral pole of the embryo. Remove the anterior piece of the embryo from the well moving it away to the side of the dish.
And then use the wire tools to scrape away remaining yolk cell granules from the posterior section, including the PSM. Next, flatten and orient the PSM with the anterior end pointing away and the posterior pointing forward. If the thin layer of ectoderm has not pulled off by this point, use the wire tools to peel it away from the top of the PSM tissue using the micro scalpel, cut the most posterior tip of the PSM past the end of the no cord or the tail bud away from the rest of the tissue.
Move the tail bud piece to one corner of the dish away from the dissecting area and use the wire tools to clear any debris and unwanted embryo tissue from the dissecting field pool. Tail bud Pieces from multiple embryos depending on how many cells will be required for the experiment. On average, a single piece of tail bud tissue yields 1000 cells.
Fill an empty 35 millimeter plastic dish with a small volume of tripsin EDTA and using the fire polished glass pipette. Transfer the tail bud pieces into the dish. Incubate for 20 minutes at room temperature while the tissue is incubating in trypsin, add milli Q water to the fibronectin one solution in the glass bottomed imaging dish.
Remove the solution and wash from the glass three times with Milli Q water. Use suction to remove each wash and ensure the dish is completely dry. To image the tail bud pieces pipette 100 microliters of L 15 with serum into an imaging dish.
Using a coated gel tip, transfer the tail bud pieces in as small a volume of trypsin EDTA as possible into the imaging dish. Pipette the pieces up and down multiple times to break them apart and suspend the cells in medium. Be careful not to introduce air bubbles.
Allow the cells to settle onto the fibronectin one coated glass for 20 minutes at room temperature. After the incubation, add a small volume of additional L 15 medium with serum to the cells. Before beginning imaging.
Use care not to disrupt settled cells. To image the cells. Begin by placing the dish in the temperature chamber on the time-lapse microscope.
Set the desired temperature and allow the dish to equilibrate for at least 30 minutes before beginning image acquisition. A stable temperature is essential for accurate measurements of period. In single PSM cells acquire test fluorescent images to check that exposure.
Time and gain. Provide a large dynamic range of intensities without saturation to ensure good signal to noise levels. A typical fluorescence image acquired from dispersed cells generated from our lines.
Using this protocol requires 400 milliseconds and 40 milliseconds for a transmitted light image with an E-M-C-C-D camera operating at an EM gain of 85. In addition, preamp gain and readout speed from the camera are also essential to maximize signal over noise using the transmitted light channel, choose a field of cells for the time-lapse acquisition. Run a time-lapse acquisition protocol by acquiring one transmitted light and one fluorescence image per field with an interval such as two minutes that will capture temporal dynamics without photo bleaching or inducing toxicity in the cells.
To process time-lapse movies, open a file and split the transmitted light and fluorescent frames to create two stacks of images. To track a single cell in the transmitted light channel, choose a circular region of interest or ROI around a cell that was still healthy at the end of recording. That does not move outside the field and that does not come in contact with other cells.
Save an ROI every few frames to the ROI manager and track the cell until the last frame. Select the fluorescent stack and with the saved ROIs, use the custom circle INTERPOLATOR plugin and macro to measure the intensity of the tracked cell over time. Check the output trace for the macro if it qualitatively captures features of the fluorescence time-lapse.
Export the values to an Excel worksheet. Save the ROI list for the cell. Shown here are representative images from a single PSM cell in a 10 hour time lapse recording.
The transgene used here generates a reporter whose cycle of production and degradation occurs with similar dynamics and expression levels to the endogenous gene end protein in the embryo using standard imaging conditions with L 15 medium containing 10%fetal bovine serum. We find that PSM cells can produce between two and seven peaks with the mean and standard deviation of three plus or minus one peaks. The median peak number as well as the 25%percentile is two peaks, and the 75%percentile is three peaks.
Using our semi-automated tracking and analysis, we can quickly generate fluorescence intensity over time traces for individual PSM cells. These raw traces can then be used to make quantitative measurements of properties of the oscillating PSM cells, such as frequency amplitude, number of cycles, and timing of peaks Following this procedure, other methods such as the addition of signaling molecules or inhibitors can be performed to answer the question how the single cell oscillator responds to factors found outside in the embryo. We hope that this technique will enable researchers in the field of cementogenesis to study single cell behavior of the segmentation clock in a simple way that gets the cells away from the complexity of cell cell interactions and tissue level factors that make this a very difficult job indeed in the intact embryo.
