Our understanding of eye morphogenesis comes from previous fixed tissue studies. However, these studies lacked cell and tissue dynamics. Using time-lapse imaging, we're able to capture this entire process for visualization and analysis.
Our method that takes advantage of the external development in optical clarity of zebrafish embryos, which facilitate live imaging, and uses laser scanning confocal microscopy which is readily available on most research campuses. Researchers new to this method should expect some trial and error before obtaining a usable dataset. Many steps, especially the injections and embedding, must go well in order for success, so have patience.
Once the zebrafish begin to breed, while waiting 15 to 20 minutes to ensure that the eggs become fertilized, use a P10 pipette and P10 microliter tips to back load a pulled glass capillary micro injection needle with 2.5 to 5 microliters of the RNA dilution of interest. When the eggs are ready, use a transfer pipette and a dissecting microscope to carefully load the eggs into the injection mold, using forceps to roll the embryos such that the single cell is visible as necessary. Then inject all of the embryos at the one-cell stage, targeting the cell and not the yolk to ensure a uniform labeling of the developing embryo.
At about 11 hours post-fertilization, place a one to five milliliter aliquot of 1.6%low melt agarose in E3 medium in a 42 degree Celsius heat block and use a fluorescence microscope to screen for successfully injected embryos with an overall brightness of florescence. Count the somites to properly stage the embryos. At 11 hours post-fertilization, three somites should be observed.
An ideal sample will have strong EGFP and mCherry signals and be at the correct developmental stage. Before mounting, place the embryos in an agar-coated dish and use forceps to remove the chorion from each embryo. When all of the embryos have been denuded, use a glass roller pipette to aspirate one embryo.
Eject as much E3 as possible so that the embryo sits at the tip of the glass Pasteur pipette and drop the embryo into the tube of agarose from the heat block. Let the embryos sink into the agarose for a few seconds before aspirating a small volume of agarose along with the embryo, taking care that the embryo remains at the tip of the pipette. Eject the embryo and agarose into an agarose droplet in a glass bottom dish and quickly but carefully use forceps to orient the embryo dorsal side down before the agarose droplet hardens.
After mounting 10 to 12 embryos in the same manner, add enough agarose to completely cover the bottom of the dish encasing all of the droplets in a single agarose disc. When the agarose has hardened, layer E3 over the agarose to keep the samples hydrated. After setting the laser scanning confocal microscope to the appropriate parameters for time-lapse imaging, liberally coat the underside of the glass bottom dish with immersion medium matching the refractive index of water, taking care to avoid air bubbles, and apply a small drop of immersion medium to the 40X water objective.
Secure the glass bottom dish in the stage insert and add additional E3 medium to keep the embryos moist overnight. Use modeling clay to seal the plastic lid over the dish and raise the objective to make contact with the dish. Under the positions heading of the microscope software, click add to save the XYZ information of the first sample to be time-lapsed and select a sample with a strong fluorescence and optimal mounting.
Under the locate tab, search for the next sample and click position and add to select the sample as demonstrated. When all of the samples have been selected, highlight the first position and click move to. While continuously scanning, line up the optic vesicle within the frame, leaving ample space in the anterior and distal regions relative to the optic vesicle and brain.
To assign the first and last Z-slices, select set first and set last while moving through the Z-direction with the fine focus knob, maintaining a total slice number of about 63 to accommodate the growth of the optic cup. Include extra room on the ventral side of the optic vesicle to allow room for growth. Once the first and last Z-slices have been set, click C to move to the center of the Z-stack and adjust the laser power and gain for both lasers.
When the laser power and gain have been set, stop the scanning and click update to update the position information. To move to the next position, click position two. Once both the first and last Z-slices have been defined as demonstrated, open the time series heading and set the number of cycles to 300 and the time interval to 2.5 minutes.
After reviewing the settings to make sure everything is finalized, click start to begin the time-lapse and confirm that the first round of imaging is captured correctly before allowing the time-lapse to run overnight. Direct injection into the single cell is necessary for a uniform labeling and a robust fluorescence. Upon their immersion in agarose, the embryos should be spaced evenly throughout the dish.
If the embryos are staged correctly, sufficiently fluorescent and adequately mounted, they will remain in the imaging frame during the time-lapse, allowing the entire organ to be imaged. Samples that are not oriented along the dorsal axis, such as those that are horizontal or diagonal, will grow out of the imaging frame as the time-lapse proceeds and cannot be used for further analysis. An embryo that is over-rotated will result in a more posterior time-lapse.
An embryo that is under-rotated will allow only the most anterior tissue to be observed. In addition, samples that are mounted with respect to the orientation of the frame are more likely to yield a successful time-lapse. This bias toward the ventral side provides extra space for tissue growth in the ventral direction resulting in an optimal time-lapse.
When these criteria are not met, the time-lapse will no longer capture the entire 3D volume of the tissue as it develops and cannot be used for further analysis. These information-rich datasets can be analyzed in many ways, including 4D cell tracking with cell speed and trajectory quantifications, cell and tissue volume measurements, and 3 and 4D visualizations.
