This method can help answer fundamental questions about the dynamics of neural progenitor cells and their progeny which underlie vertebrate brain development. The main advantage of this technique is the ability to directly visualize multiple clusters of clonally related cells within the living brain over many hours of development. The protocol can also be applied to studying clonal and progenitor dynamics in other developing systems of the zebrafish embryo.
Visualization using this method is particularly useful for directly observing fundamental processes that occur during neural development. On the afternoon prior to performing micro injections, set up wild type adult zebrafish in sex segregated mating tanks. On the next morning prepare the DNA solution according to manuscript directions and inject approximately 4.2 nano liters of this solution into one cell zebrafish embryos within 45 minutes of fertilization.
Maintain the injected embryos in E three medium and a 28 degrees Celsius incubator for 24 hours, then call the dead and deformed embryos from the group. Transfer up to 20 healthy embryos into a 50 milliliter tube, fill it with 10 milliliters of E three and place a cap on top of each tube. Place a rack with the 50 milliliter tubes upright in a 37 degrees Celsius water bath for 80 to 90 minutes, making sure that the water level in the bath is higher than the E three in the tube.
Remove the tube rack from the water bath and place it in the 28 degrees Celsius incubator. Allow up to one hour for the E three to cool and the embryos to reacclimate to the temperature. Then transfer them to Petri dishes with 0.2 millimolar PTU and E three kept warm in the 28 degrees Celsius incubator.
At two to four hours after the heat shock, examine the embryos under a standard fluorescence dissection microscope for expression of CFP or YFP which indicates successful Brainbow recombination. Select embryos with a robust FP expression throughout and transfer them to a separate dish with PTU. Image them one, two, or three days post fertilization.
Expression which appears dim under a fluorescence microscope may actually be well visualized in confocal time-lapse imaging. Prior to the day of the experiment, prepare the imaging chamber and embryo manipulation tool. To prepare the imaging chamber, carefully superglue a plastic ring to the center of a 60 millimeter Petri dish.
Construct the manipulator by super gluing a small length of nylon fishing line to the end of a four inch wooden swab stick. If necessary, dechorionate the embryos under a dissection microscope prior to mounting. To mount the embryos, transfer the fish to the center of the plastic ring in the imaging chamber and remove as much excess E three as possible with a fine tipped transfer pipette.
Use a clean transfer pipette to cover the fish with one percent low melt agarose and E three, filling the entire plastic ring with a layer of agarose. Then gently pull the embryo up into the pipette tip and back into the agarose without introducing any air bubbles. Use an embryo manipulator to quickly orient the fish and the agarose before it hardens.
If imaging with an upright microscope, position the embryo as close to the upper surface of the agarose as possible, making sure that they are parallel to the bottom of the imaging chamber with their tail straight. Wait for the agarose to harden, then fill the imaging chamber with E three, adding as much as possible to account for evaporation over the course of imaging. Place the imaging chamber with the fish and E three on the confocal microscope.
Then select the objective with a high numerical aperture and a long working distance. Find a region with appropriately dense and bright labeling of cells and set up the acquisition parameters. This is an example using Zen software, but settings will vary depending on microscope and laser lines available.
Prepare three tracks to image each FP channel sequentially. Use an argon laser to excite CFP at 458 nanometers and YFP at 514 nanometers, and use a DPSS 561 nanometer laser to excite dTomato. Collect the missions for the three at the appropriate wavelengths.
Select the Z stack range to image and a time interval between 10 and 30 minutes to track mitotic and apoptotic events. Finally, select the length the imaging session and run the experiment. After imaging is complete, save the raw data in CZI format if using Zen or another format compatible with Fiji, then import the images into Fiji using the bio formats importer.
In vivo multicolor time-lapse imaging was used to show Brainbow color coded clones of cells in the proliferative ventricular zone of the developing zebrafish hindbrain. Brainbow labeled cells arranged along a particular radial fiber shared the same color. Which can be quantified as the relative RGB channel weights.
This suggests that these radio groups were clones of dividing cells and that their similar color could be used to identify them as being clonally related. A quantitative color analysis showed that daughter cells express the same color as their mother's cell. But that neighboring radio clusters of cells can be distinguished from one another.
Numerous clones of related cells can be followed simultaneously over hours in vivo, allowing for a multiplex lineage analysis and comparison. Quantification of color expression in clones at two and then again at three days post fertilization showed that Brainbow expression remained relatively constant from two to three days. Time-lapse imaging revealed numerous cells undergoing interkinetic nuclear migration and cell division from one to two days post fertilization, making it possible to study the cell cycle.
An average cell cycle of 8.4 hours was calculated which is comparable to previous measurements in zebrafish. Furthermore, this imaging technique was used to observe individual cells undergoing the stereotypical morphological changes associated with apoptosis, such as membrane blebbing and cell fragmentation. Because this method is performed in vivo, zebrafish can be rescued from the imaging chamber and maintained for imaging or other experiments at later time points.
The use of this technique allows us to explore new questions about the roles of clonally related cells during neural development.
