Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

The overall goal of this procedure is to observe and characterize pharyngeal arch movements in the developing zebra fish head. This is accomplished by first embedding a transgenic zebra fish embryo in anesthetic acrost gel. The second step is to mount the agarose embedded embryo in a drop of Methylcellulose on a microscope slide.

Next, the embryo is sealed between two microscope slides with vacuum grease. Ultimately time-lapse confocal microscopy is used to show differences between wild type and mutant zebrafish and the morphogenetic movements of the pharyngeal arches. The main advantage of this technique over static imaging methods is that it captures pharyngeal arch development in real time, allowing an investigator to directly observe the tissue movements that are critical to morphogenesis.

To make a bridged cover slip, super glue, A 22 by 22 cover glass onto each end of a 24 by 60 cover glass. Leaving the middle free. Make singles for embryos less than a day old.

Doubles for embryos one to four days old or triples for embryos, five days or older. And for each specimen, use two bridged cover slips. Now fill a 10 milliliter syringe with high vacuum grease.

Prepare fresh anesthetic in 0.2%liquid aros and maintain in a 42 degree Celsius heating block. To manually dec coate the unhatched transgenic embryos slowly tear the Corian open until the embryo is freed. Now transfer the coated embryos into clean media containing MS 2 22 and aesthetic.

To prepare a bridged cover slip slowly push the vacuum grease out of the syringe, making a smooth continuous bead that is adjacent to and inside of the bridges and the edge of the cover slip. Spread a circle of 4%Methylcellulose in the middle of the bridge to cover slip. When the embryo is anesthetized, draw it up into a glass pipette.

Hold the pipette up vertically to allow the embryo to sink to the bottom of the liquid in the pipette. Then move the pipette into the agro solution and allow the embryo to drop into the aros without expelling the liquid. Draw some agro solution and the embryo into the pipette.

Then expel the embryo in a drop of aro solution. On top of the methylcellulose. Push the embryo downward onto the methylcellulose and orient the embryo as desired for imaging to seal the specimen.

Place one bridged cover slip on top of the mounted one facing downward. Apply even pressure to both sides of the sandwiched cover slips until the bridges make direct contact and the agros drop seals to both cover slips. Verify that the embryo is properly oriented.

Rub a wooden applicator along the glass to smooth out cracks and gaps in the vacuum grease speed. Wipe away any excess grease from the edges. Activate the confocal microscope and any external laser devices.

Also turn on the computer connected to the confocal microscope and select start system and the confocal imaging software. Next, lower the staging platform of the confocal microscope and move the objective lenses out of position. Replace the default stage with an electronic heated stage.

Turn on the controller for the heated stage and set the temperature for 29.5 degrees Celsius. Place the mounted specimen on the heated stage. Move the 20 x objective lens into position and raise the staging platform.

Then activate the visible light emitter and center the embryo's head in the field of view. To activate the fluorescence channels, select the fluorescent wavelength to be detected and choose color lookup tables for each channel. For each fluorescence channel, set the laser intensity to 10.0 and the photo multiplier voltage between 600 and 700.

To reduce background and increase the fluorescent information collected, set the averaging to two or four and the bit depth to 16 bit. Activate the ZS stack and time series modules. Adjust the pinhole if necessary.

Set the time interval and the length of the experiment. Turn the camera to live mode. Adjust the focus and increase laser intensity if necessary.

To see fluorescence. Set the digital zoom to 0.6 for a wider view. If desired.

Position the stage such that all structures of interest are visible and there is room in the field of view for growth. Dorsally and anteriorly focus through the embryo to the upper and lower limits of fluorescence. Set the first and last Zack slice positions at least 100 micrometers or more beyond these limits.

Next, set the display lookup table to range indicators so the fluorescent structures appear a shade of gray and the rest of the field should be blue or black. If needed, increase the photo multiplier voltage to a level just below where saturated red pixels appear. If fluorescent structures appear red, reduce the laser intensity and or the gain until they become a shade of gray.

If necessary, adjust offset. So areas outside the sample are blue. Set the for the smallest diameter that allows images to be collected in a timely fashion.

The Z interval should always be set to the suggested optimal distance for the pinhole diameter, perform short scans to test the settings, always using the lowest laser intensity possible. Now run the experiment for the desired length of time. Afterward, remove the embryo from the heated stage and slowly separate the cover slips, submerge the cover, slip and embryo in em in a 30 millimeter Petri dish.

Using a glass pipette, gently wash the embryo off of the cover slip and remove the cover slip from the Petri dish. Place the embryo into a 28.5 degree Celsius incubator to continue to develop. At the end of the experiments, take individual Zack images of agros mounted sibling embryos that were similarly staged to the specimen at the beginning of the experiments, but were raised an EM and a 28.5 degree Celsius incubator.

Measure the pharyngeal arches as needed using confocal software or other imaging processing software in wild type embryos. Following neuro crest population, the pharyngeal archs elongate along the anterior posterior and dorsal ventral AEs while moving in a rostral direction. At its anterior end, the first pharyngeal arch forms maxillary and mandibular domains separated by the oral ectoderm key morphogenetic movements are greatly disrupted in smoothened mutants.

Here the seventh pharyngeal arch fails to migrate roly to overlap with more anterior arches by 48 hours post fertilization. Zebrafish SM five mutants have numerous cranial facial defects. In these static confocal images.

Fusions in the ventral domains of the second and third arches are apparent by 32 hours post fertilization. Using time-lapse confocal microscopy, we analyze the morphogenesis of the pharyngeal arches. In Smad five mutants fusions between the second and third arches appear first at 32 hours post fertilization.

Consistent with the static analysis. While the expression of PDGF family members suggest that it may be broadly involved in pharyngeal arch morphogenesis movement of the posterior arches appears normal. In P-D-G-F-R-A mutants, it is morphogenesis of the first pharyngeal arch that appears specifically disrupted.

This procedure is amenable to other commonly used zebrafish techniques such as morpho injection or the generation of genetic mosaics. Time lapse analyses allow for the direct analysis of the consequences of these manipulations. After watching this video, you should have a good understanding of how to perform time lapse in vivo imaging of transgenic zebrafish embryos by properly anesthetizing and mounting specimens and setting ideal parameters for confocal microscopy.