We thank Melissa Griffin and Jenna Rozacky for their expert fish care. PDM thanks EGN for writing assistance, generosity, and patience. This work was supported by NIH/NIDCR R01DE020884 to JKE.

1. Animal Husbandry and Mutant Alleles
<ol>
	<li>Raise and breed zebrafish as described21.</li>
	<li>Zebrafish mutant alleles used in this study were pdgfra b1059 16, smad5 b1100 22, and smob577 23. Sources for these zebrafish strains include ZIRC.</li>
</ol>
2. Preparation of Solutions and Implements
Note: All solutions and implements can be made in advance and stored for future use.
<ol>
	<li>Make embryo media (EM) as previously described21.</li>
	<li>Make 4 g/L MS-222 (Tricaine). Dissolve 4 g disodium phosphate (Na2HPO4) in 450 ml sterile water. Add 2 g MS-222 to the solution. Adjust pH to within 7.0-7.2. Add sterile water to a total volume of 500 ml.</li>
	<li>Optional: Make 4 g/L clove oil. Weigh 0.2 g pure clove oil in a 50 ml conical tube. Add EM to a total volume of 50 ml. Store at 4 &#176;C.</li>
	<li>Make 3% methylcellulose. Bring 50 ml of EM + 0.1 M HEPES to a boil. Remove from heat. Stir in 1.5 g methylcellulose until the solution is homogeneous. Store at 4 &#176;C.</li>
	<li>Make 0.2% agarose. Add 0.1 g agarose to 50 ml EM. Bring EM to a boil in the microwave or on a hotplate and verify that the agarose has dissolved. Aliquot into 500 &#956;l volumes in microcentrifuge tubes and store at 4 &#176;C. Note: Volumes can be adjusted as needed.</li>
	<li>Make capillary pokers. Place a drop of super glue inside a 0.9 mm ID capillary tube. Insert a 4-5 cm length of 6 lb test monofilament fishing line inside the capillary tube such that approximately 3-4 mm of line extends beyond the end of the tube.</li>
	<li>Make two bridged coverslips per specimen. Super-glue 22 x 22-1 cover glass onto each end of a 24 x 60-1 cover glass leaving the middle free. Make singles (1 small cover glass per end) for embryos less than a day old, doubles (2 small cover glasses on top of each other per end) for embryos one to four days old, or triples (3 small cover glasses on top of each other per end) for embryos five days or older.</li>
	<li>Fill a 10 ml syringe with high vacuum grease. Note: Vacuum grease serves as a sealant to prevent dehydration of specimens over long periods of time, and has low potential for toxicity compared to more volatile sealants.</li>
</ol>
3. Mounting Zebrafish Embryos for Confocal Microscopy (See Figure 1)
<ol>
	<li>Prepare anesthetic medium. Melt an aliquot of 0.2% agarose by microwaving in 20&#160;sec intervals until completely liquid. Mix 8 &#181;l of MS-222 into 192 &#181;l of 0.2% agarose or 5 &#181;l of 4 g/L clove oil into 195 &#181;l of 0.2% agarose. Maintain in a 42 &#176;C heating block. Note: Long exposure to MS-222 causes stronger developmental delays than clove oil does in embryonic zebrafish24.</li>
	<li>If necessary: Manually dechorionate unhatched transgenic embryos to be analyzed just prior to analysis. Using forceps, slowly tear the chorion open until the embryo is freed.</li>
	<li>Anesthetize the zebrafish embryos. Add 1 ml of a 4 g/L stock of MS-222 to 24 ml of embryo media. Alternatively, add 390 &#181;l of 4 g/L clove oil to 24 ml of fish water24. Note: Clove oil acts slowly so perform this step at least 15-20 min before time-lapse microscopy.</li>
	<li>Place a bead of vacuum grease around the center space of an upward-facing bridged coverslip. 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 coverslip.</li>
	<li>Spread a circle of 4% methylcellulose in the middle of the bridged coverslip using a wooden applicator.</li>
	<li>Transfer the embryo to be imaged. 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. Move the pipette into the agarose solution and allow the embryo to drop into the agarose. Do not expel the liquid.</li>
	<li>Place the specimen. Draw some agarose solution and the embryo into the pipette. Expel the embryo in a drop of agarose solution on top of the methylcellulose. Use a capillary poker to push the embryo downward onto the methylcellulose and orient the embryo as desired for imaging.</li>
	<li>Seal the specimen.
	<ol>
		<li>Place one bridged coverslip on top of the mounted one, facing downward. Apply even pressure to both sides of the sandwiched coverslips until the bridges make direct contact and the agarose drop seals to both coverslips. Verify that the embryo is properly oriented.</li>
		<li>Rub a wooden applicator along the glass to smooth out cracks and gaps in the vacuum grease bead. Wipe away excess grease from the edges.</li>
	</ol>
	</li>
</ol>
4. Time-lapse Confocal Imaging of Transgenic Zebrafish Embryos
(This protocol has been optimized for a Zeiss LSM 710 confocal microscope using ZEN imaging software, and can be modified for use with other systems.)
<ol>
	<li>Activate equipment. Turn on the confocal microscope and any external laser devices. Turn on the computer connected to the confocal microscope. Open confocal imaging software and select &#34;Start System&#34; to activate image acquisition controls.</li>
	<li>Prepare heated stage. 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 &#176;C.</li>
	<li>Position the specimen in the field of view. Place the mounted specimen on the stage. Move the 20X&#160;objective lens into position and raise the staging platform. Activate the visible light emitter and look through the eyepieces. Center the embryo&#39;s head in the field of view.</li>
	<li>Define experiment settings.
	<ol>
		<li>Activate fluorescence channels by selecting the fluorescent wavelength to be detected and choose color lookup tables (LUT) for each channel. If necessary, use manual controls in the software to turn on the requisite lasers (e.g.&#160;561 nm for RFP). For each fluorescence channel, set the laser intensity to 10.0 and the photomultiplier voltage (HV or gain) between 600-700. Set the averaging to 4 and the bit depth to 16 bit.</li>
		<li>Activate the Z-stack and time series modules. Set the pinhole for a 5 &#181;m or smaller Z-resolution, and set the Z-interval to the suggested optimal distance. 
		Note: Generally, 5 &#181;m Z-resolution allows each Z-stack image to be collected quickly enough for a &#34;snapshot&#34; of the embryo at a particular time point while also resolving individual cells. Lowering the averaging also reduces the amount of time per image.</li>
		<li>Define the desired time interval between images (usually 10-20 min) and the total length of the experiment.</li>
	</ol>
	</li>
	<li>Set positional information.
	<ol>
		<li>Turn the camera to live mode. Adjust focus and increase laser intensity if necessary to see fluorescence. Set the digital zoom to 0.6 for a wider view, if desired.</li>
		<li>Position the stage such that all structures of interest are visible and there is room in the field of view for outgrowth dorsally and anteriorly. Focus through the embryo to the upper and lower limits of fluorescence. Set the first and last Z-stack slice positions at least 100 &#181;m or more beyond these limits.</li>
	</ol>
	</li>
	<li>Optimize fluorescence intensity.
	<ol>
		<li>Set the display LUT to range indicator. Fluorescent structures should now appear white, and the rest of the field of view should be blue (no detection) or black.</li>
		<li>Increase HV/gain to a level just below where saturated (red) pixels appear. If necessary, adjust offset so a majority of the background is not detected (blue). Note: Smaller pinhole diameter and increased laser intensity both improve image definition, but laser intensity must be kept low to preserve living tissues. Increased HV/gain and pinhole diameter (&#60;5 &#181;m) are generally preferable to increased laser intensity.</li>
		<li>Keep the pinhole diameter wide enough to collect images in a timely fashion, averaging sets to 4. The Z-interval should always be set to the suggested optimal distance for the pinhole diameter. Perform short scans (averaging = 1) with different settings, and use the minimum laser intensity that achieves desired resolution.</li>
	</ol>
	</li>
	<li>Run the experiment for the desired length of time. Afterward, remove embryo from the heated stage and slowly separate the coverslips. Submerge the coverslip and embryo in EM in a 30 mm Petri dish. Using a glass pipette gently wash the embryo off of the coverslip and remove the coverslip from the Petri dish. Place the embryo into a 28.5 &#176;C incubator to continue to develop.</li>
	<li>Compare growth. At the end of the experiment, take individual Z-stack images of agarose-mounted sibling embryos that were similarly staged to the specimen at the beginning of the experiment, but were raised in EM in a 28.5 &#176;C incubator.</li>
	<li>Measure pharyngeal arches, as needed. Measurements can be performed in some confocal software suites. Measurements found here were performed using Fiji25 by importing and Z-projecting .lsm files of individual frames.</li>
</ol>

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The authors declare that they have no competing financial interests.

Time-lapse confocal microscopy is a powerful tool for the analysis of development. Here, we demonstrate the method&#39;s usefulness in studying pharyngeal arch morphogenesis in zebrafish that are mutant for important signaling pathways using a transgenic that labels neural crest cells. In addition to tissue-level analyses, time lapse analyses are also applicable to analyses at a cellular scale28. Many widely used zebrafish methods can also be incorporated into time-lapse microscopy experiments, including microinjection of morpholinos, mRNAs, or transgenic constructs, as well as transplants between embryos. Analysis of multiple developing tissues is limited only by the ability to fluorescently label tissues and by a confocal microscope&#39;s capability to detect the desired number of different fluorophores without crosstalk. The number of individuals that can be imaged from a clutch of zebrafish embryos is severely constrained, but advanced techniques allow automated imaging of several embryos in the same time period. This is especially useful when seeking to perform time-lapse analysis of a mutant that is not easily identified at early time points.
When performing measurements from time-lapse confocal images, it is important to consider that some developmental delay is caused by the process. Developmental delays are not uncommon in mutant embryos as well, so imaging of sibling controls is necessary for both mutant/manipulated embryos as well as wild-type controls. Even once controlled for staging, comparative measurement of fluorescence-labeled structures is difficult. Fluorescence intensity can be highly variable between individuals and between separate parts of the same structure. Thus, nondiscrete quantitative measurements have a degree of subjectivity, and we recommend that more than one individual perform measurements to ensure consistency. Investigators must also be disciplined in orienting embryos for comparative measurement consistently, as failure to do so will cause the appearance of morphological differences where there may be none. Movements of specimen embryos can outright ruin an experiment, and we find that adding anesthetic after the microwaved agarose has been allowed to thicken reduces its efficacy in sedating embryos.
Though difficult to acquire, time-lapse microscopy data have obvious advantages over static images in comparing morphogenesis qualitatively. Static images will often allow easy measurement of structures at a certain time point, but give no indication of tissue behaviors leading up to or following that stage. Such information is crucial when studying the interactions of two or more separate tissues undergoing morphogenesis. In order to obtain the best overall characterization of morphogenesis, it is beneficial to perform both time-lapsed and static analyses to best utilize the advantages of each method. In the time-lapsed analyses, the dynamic cell and tissue behaviors can be analyzed and, in static analyses, large numbers of embryos can be examined to gain the best understanding of variability. The growing number of zebrafish transgenic lines and advancement of other techniques will make time-lapse confocal microscopy of multiple tissues commonplace, as real-time in vivo data becomes the gold standard among morphogenesis researchers.

Craniofacial morphogenesis is a complex multi-step process that requires coordinated interactions between multiple cell types. The majority of the craniofacial skeleton is derived from neural crest cells, many of which must migrate from the dorsal neural tube into transient structures called pharyngeal arches1. As with many tissues, morphogenesis of the craniofacial skeleton is more complicated than can be understood by static images of embryos at specific developmental time points. Although it is time-consuming to perform, in vivo time-lapse microscopy provides a continuous look at a developing embryo&#39;s cells and tissues. Each image in a time-lapse series lends context to the others, and helps an investigator move toward deducing why a phenomenon occurs rather than deducing what is occurring at that time.
In vivo imaging is thus a powerful descriptive tool for experimental approaches to deconstruct the pathways that guide morphogenesis. The zebrafish Danio rerio is a popular genetic model of vertebrate embryonic development, and is particularly well suited for in vivo imaging of morphogenesis. Modern, convenient methods for transgenesis and genomic modification are rapidly advancing the number of tools available to zebrafish researchers. These tools enhance already robust methods for genetic manipulation and microscopy. In vivo imaging of almost any tissue in almost any desired genetic context is closer to reality than fantasy.
Morphogenetic movements of the pharyngeal arches are guided by signaling interactions between the neural crest and the adjacent epithelia, both ectoderm and endoderm. There are numerous signaling molecules expressed by the epithelia that are necessary to drive the morphogenesis of craniofacial skeletal elements. Among these signaling molecules, Sonic Hedgehog (Shh) is critically important for craniofacial development2-8. Shh is expressed by both the oral ectoderm and pharyngeal endoderm2,6,9,10. The expression of Shh in the endoderm regulates morphogenetic movements of the arches10, patterning of neural crest within the arches10, and growth of the craniofacial skeleton11.
Bmp signaling is also critically important for craniofacial development12 and may alter morphogenesis of the pharyngeal arches. Bmp signaling regulates dorsal/ventral patterning of crest within the pharyngeal arches13,14. Disruption of smad5 in zebrafish causes severe palatal defects and a failure of the Meckel&#39;s cartilages to fuse appropriately at the midline15. In addition, the mutants also display reductions and fusion in the ventral cartilage elements, with the 2nd, 3rd, and sometimes 4th pharyngeal arch elements fused at the midline15. These fusions strongly suggest that Bmp signaling directs the morphogenesis of these pharyngeal elements.
Pdgf signaling is necessary for craniofacial development, but has unknown roles in pharyngeal arch morphogenesis. Both mouse and zebrafish Pdgfra mutants have profound midfacial clefting16-18. At least in zebrafish this midfacial clefting is due to a failure of proper neural crest cell migration16. Neural crest cells continue to express pdgfra after they have entered the pharyngeal arches. Additionally, Pdgf ligands are expressed by facial epithelia and within the pharyngeal arches16,19,20, thus Pdgf signaling could also play a role in morphogenesis of the pharyngeal arches following migration. However, analyses of the morphogenesis of the pharyngeal arches in pdgfra mutants have not been performed.
Here, we demonstrate in vivo confocal microscopy of pharyngula-stage transgenic zebrafish and describe the morphogenesis of the pharyngeal arches within this period. We further demonstrate tissue behaviors that are affected by mutations that disrupt the Bmp, Pdgf, and Shh signaling pathways.

<table><tbody><tr><td>6 lb Test monofilament line</td><td>Cortland Line Company</td><td>SLB16</td><td></td></tr><tr><td>Agarose I</td><td>Amresco</td><td>0710</td><td></td></tr><tr><td>Argon laser</td><td>LASOS Lasertechnik GmbH</td><td>LGN 3001</td><td></td></tr><tr><td>Calcium chloride</td><td>Sigma-Aldrich</td><td>C8106</td><td></td></tr><tr><td>Capillary tubing, 100 mm, 0.9 mm ID</td><td>FHC</td><td>30-31-0</td><td></td></tr><tr><td>Clove oil</td><td>Hilltech Canada, Inc.</td><td>HB-102</td><td></td></tr><tr><td>High vacuum grease</td><td>Dow Corning</td><td>2021846-0807</td><td></td></tr><tr><td>Isotemp dry-bath incubator</td><td>Fisher Scientific</td><td>2050FS</td><td></td></tr><tr><td>Laser scanning microscope</td><td>Carl Zeiss AG</td><td>LSM 710</td><td></td></tr><tr><td>Magnesium sulfate hexahydrate</td><td>Sigma-Aldrich</td><td>230391</td><td></td></tr><tr><td>Microscope cover glass, 22 x 22-1</td><td>Fisher Scientific</td><td>12-542-B</td><td></td></tr><tr><td>Microscope cover glass, 24 x 60-1</td><td>Fisher Scientific</td><td>12-545-M</td><td></td></tr><tr><td>Potassium chloride</td><td>Fisher Scientific</td><td>M-11321</td><td></td></tr><tr><td>Potassium phosphate dibasic</td><td>Sigma-Aldrich</td><td>P3786</td><td></td></tr><tr><td>Sodium chloride</td><td>Fisher Scientific</td><td>M-11624</td><td></td></tr><tr><td>Sodium phosphate dibasic</td><td>Sigma-Aldrich</td><td>S7907</td><td></td></tr><tr><td>TempController 2000-2</td><td>PeCon GmbH</td><td></td><td></td></tr><tr><td>Tricaine-S</td><td>Western Chemical, Inc.</td><td></td><td></td></tr></tbody></table>

analyzing craniofacial morphogenesis in zebrafish using 4d confocal microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

In wild-type embryos, following neural crest population, the pharyngeal arches elongate along the anterior/posterior and dorsal/ventral axes while moving in a rostral direction (Movie 1). At 30 hours post-fertilization (hpf), the anterior/posterior length of the first pharyngeal arch is between 1.8-1.9 times its dorsal/ventral height. Dorsal/ventral elongation proceeds steadily, faster than anterior/posterior extension until 36.5 hpf. From here, dorsal/ventral height plateaus around 104 &#181;m through 48 hpf. Anterior/posterior extension continues through 48 hpf, with overall anterior/posterior length increasing from under 1.5 times the dorsal/ventral height at 36.5 hpf to 1.95 times the dorsal/ventral height at 48 hpf.
At its anterior end, the first pharyngeal arch forms maxillary (dorsal) and mandibular (ventral) domains separated by the oral ectoderm. The maxillary domain of the first pharyngeal arch is measured from its distal tip to the posterior apex of the oral ectoderm. It grows at a rate approximated by linear regression at 4.6 &#181;m/hr (r2 = 0.9374) between 30-48 hpf and is about 104 &#181;m long at 48 hpf (Movie 1).
Pharyngeal arches 3-7 all form from the segmentation of a single mass of neural crest cells via the pharyngeal endoderm26. Between 30-38 hpf, the 5th through 7th pharyngeal arches separate from the mass, the 3rd and 4th arches already having separated. Around 37 hpf, the 3rd pharyngeal arch begins to move medially to the 2nd arch. Laterally, the 7th pharyngeal arch overtakes the 5th and 6th arches in rostral movement by 48 hpf, leaving the 3rd through 6th arches medial to the 2nd and 7th arches (Movie 1).
It is important to note that although the convention is to describe the morphology in hpf, microscopy slightly delays zebrafish growth compared to similarly staged embryos grown in a 28.5 &#176;C incubator. For example, after 19 hr of microscopy, the distance between the eye and ear of an imaged wild-type embryo was approximately 81.7 &#181;m, and the average eye-ear distance for two sibling embryos imaged immediately afterward was 71 &#181;m. Therefore, embryos are likely to be slightly younger than the staging series27.
These morphogenetic movements are greatly disrupted in smo mutants. As previously described, zebrafish smo mutants fail to form a maxillary condensation of the first arch, and by 48 hpf the first pharyngeal arch is completely caudal to the eye2. Posterior arch movements are also disrupted10. The 3rd pharyngeal arch fails to move medial to the 2nd pharyngeal arch, and Movie 2 shows that the 7th pharyngeal arch fails to migrate rostrally to overlap with more anterior arches by 48 hpf.
Zebrafish smad5b1100 mutants have numerous craniofacial defects, including a near-complete loss of the palatal skeleton and ventral fusions between the 2nd, 3rd, and sometimes 4th pharyngeal arch skeletal elements15, suggesting that there may be defects to pharyngeal arch morphogenesis. Indeed, static confocal imaging demonstrated fusions in the ventral domains of the 2nd and 3rd arches are apparent by 32 hpf (Figure 2). Using time-lapse confocal microscopy, we analyzed the morphogenesis of the pharyngeal arches in smad5 mutants. In time-lapse analyses, fusions between the 2nd and 3rd arches appear first at 32 hpf, consistent with the static analysis (Movie 3, Figure 3). Additionally, smad5 mutants have disrupted outgrowth of the first pharyngeal arch (Movie 3). At 30 hpf, the anterior/posterior length of the first pharyngeal arch (186.8 &#181;m) is over 35% longer than in wild-type, and is 2.5 times the dorsal/ventral height of the first pharyngeal arch in a smad5 mutant. However, as the posterior edge moves rostrally, the anterior/posterior length of the first pharyngeal arch decreases almost 40 &#181;m between 30-39 hpf and then regains this length by 48 hpf. The dorsal/ventral height of the first pharyngeal arch decreases steadily to 62.3 &#181;m from 30-48 hpf. Failure of anterior extension during this time is reflected in the maxillary domain, which is exceptionally long (85 &#181;m) at 30 hpf compared to wild-type. Maxillary length decreases at a rate approximated by linear regression at 4.4 &#181;m/hr (r2 = 0.9032) from 30-36 hpf, and stays fairly consistent thereafter. Here, time-lapse imaging demonstrates unique tissue movements that may account for the loss of the palatal skeleton observed in smad5 mutants.
While the expression of Pdgf family members suggests that it may be broadly involved in pharyngeal arch morphogenesis, movement of the posterior arches appears normal in pdgfra mutants. It is morphogenesis of the first pharyngeal arch that appears specifically disrupted (Movie 4). The overall size of the first pharyngeal arch is reduced relative to wild-type, and the maxillary domain appears highly plastic, with cells condensing and withdrawing over time. This results in a failure of maxillary elongation, which peaks around 39 hpf at 56.5 &#181;m. At 48 hpf, the maxillary domain measures only 44.1 &#181;m. Collectively, these findings show that careful time lapse analyses can provide details of morphogenesis potentially lost in static analyses.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51190/51190fig1highres.jpg" src="/files/ftp_upload/51190/51190fig1.jpg" /> 
Figure 1. Schematic of bridged coverslip construction and embryo mounting for confocal microscopy.
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51190/51190fig2highres.jpg" src="/files/ftp_upload/51190/51190fig2.jpg" /> 
Figure 2. The second and third arches fuse in smad5 mutants. (A,&#160;B) Single Z slices from a (A) fli1:egfp and a (B) fli1:egfp;smad5b1100 mutant embryo. (A) In embryos wild-type for smad5, neural crest cells in the second and third arch are separated by the second pharyngeal pouch (arrowhead). (B) Neural crest cells at the ventral limit of the second and third arches intermingle in smad5 mutants (arrowhead). Pharyngeal arches are numbered. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51190/51190fig2highres.jpg" target="_blank">Click here to view larger image.</a>
 
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/51190/51190fig3highres.jpg" src="/files/ftp_upload/51190/51190fig3.jpg" /> 
Figure 3. Loss of smad5 function results in disrupted pharyngeal arch morphogenesis. (A,&#160;B, upper panels) Projections and (lower panels) orthogonal views of arch fusion in Movie 3. Fusion of arches two and three is evident, arrowhead. Pharyngeal arches are numbered. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51190/51190fig3highres.jpg" target="_blank">Click here to view larger image.</a>
Movie 1. Development of the pharyngeal arches in a wild-type (fli1:egfp) embryo from 30-48 hpf. Anterior to the left, dorsal to the top. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51190/Movie_1.mov" target="_blank">Click here to view movie.</a>
Movie 2. Medial movements of the pharyngeal arches fail in smo mutants (fli1:egfp;smob577). Anterior to the left, dorsal to the top. 30-48 hpf. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51190/Movie_2.mov" target="_blank">Click here to view movie.</a>
Movie 3. The 2nd and 3rd arches fuse in smad5 mutants (fli1:egfp;smad5b1100). Anterior to the left, dorsal to the top. Series of (A) projections, 30-48 hpf, and (B) single Z slices, 30-40 hpf. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51190/Movie_3.mov" target="_blank">Click here to view movie.</a>
Movie 4. Elongation of the maxillary region fails in pdgfra mutants (fli1:egfp;pdgfrab1059). Anterior to the left, dorsal to the top. 30-48 hpf. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51190/Movie_4.mov" target="_blank">Click here to view movie.</a>

Watch this Scientific Journal Video about Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy at JoVE.com

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.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

analyzing-craniofacial-morphogenesis-zebrafish-using-4d-confocal

Nanyang Technological University

Research

JoVE Journal

Biology

11.0K Views.  The University of Texas at Austin. 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.

Video: Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Visualization of Chondrocyte Intercalation and Directional Proliferation via Zebrabow Clonal Cell Analysis in the Embryonic Meckel&#8217;s Cartilage

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. Patterning of the Meckel&#39;s cartilage, a first pharyngeal arch derivative, involves the migration of cranial neural crest (CNC) cells and the progressive partitioning, proliferation and organization of differentiated chondrocytes. Several studies have described CNC migration during lower jaw&#160;morphogenesis, but the details of how the chondrocytes achieve organization in the growth and extension of Meckel&#8217;s cartilage remains unclear. The sox10 restricted and chemically induced Cre recombinase-mediated recombination generates permutations of distinct fluorescent proteins (RFP, YFP and CFP), thereby creating a multi-spectral labeling of progenitor cells and their progeny, reflecting distinct clonal populations. Using confocal time-lapse photography, it is possible to observe the chondrocytes behavior during the development of the zebrafish Meckel&#8217;s cartilage.
Multispectral cell labeling enables scientists to demonstrate extension of the Meckel&#8217;s chondrocytes.&#160;During extension phase of the Meckel&#8217;s cartilage, which prefigures the mandible, chondrocytes intercalate to effect extension as they stack in an organized single-cell layered row. Failure of this organized intercalating process to mediate cell extension provides the cellular mechanistic explanation for hypoplastic mandible that we observe in mandibular malformations.

Cell organization of craniofacial bones has long been hypothesized but never directly visualized. Multi-spectral cell labeling and in vivo live imaging allows visualization of dynamic cell behavior in zebrafish lower jaw. Here, we detail the protocol to manipulate Zebrabow transgenic fish and directly observe cell intercalation and morphological changes of chondrocytes in the Meckel&#8217;s cartilage.

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. ...

Developmental Biology

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics

Skeletal morphogenesis occurs through tightly regulated cell behaviors during development; many cell types alter their behavior in response to mechanical strain. Skeletal joints are subjected to dynamic mechanical loading. Finite element analysis (FEA) is a computational method, frequently used in engineering that can predict how a material or structure will respond to mechanical input. By dividing a whole system (in this case the zebrafish jaw skeleton) into a mesh of smaller 'finite elements', FEA can be used to calculate the mechanical response of the structure to external loads. The results can be visualized in many ways including as a 'heat map' showing the position of maximum and minimum principal strains (a positive principal strain indicates tension while a negative indicates compression. The maximum and minimum refer the largest and smallest strain). These can be used to identify which regions of the jaw and therefore which cells are likely to be under particularly high tensional or compressional loads during jaw movement and can therefore be used to identify relationships between mechanical strain and cell behavior. This protocol describes the steps to generate Finite Element models from confocal image data on the musculoskeletal system, using the zebrafish lower jaw as a practical example. The protocol leads the reader through a series of steps: 1) staining of the musculoskeletal components, 2) imaging the musculoskeletal components, 3) building a 3 dimensional (3D) surface, 4) generating a mesh of Finite Elements, 5) solving the FEA and finally 6) validating the results by comparison to real displacements seen in movements of the fish jaw.

Finite Element Analysis is a frequently used tool to investigate the mechanical performance of structures under load. Here we apply its use to modeling the biomechanics of the zebrafish jaw.

Skeletal morphogenesis occurs through tightly regulated cell behaviors during development; many cell types alter their behavior in response to mechanical ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

Visualization of Craniofacial Development in the sox10: kaede Transgenic Zebrafish Line Using Time-lapse Confocal Microscopy

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, convergence and extension of facial prominences, and maturation of the craniofacial skeleton. To study the contribution of the cranial neural crest to specific regions of the zebrafish palate a sox10: kaede transgenic zebrafish line was generated. Sox10 provides lineage restriction of the kaede reporter protein to the neural crest, thereby making the cell labeling a more precise process than traditional dye or reporter mRNA injection. Kaede is a photo-convertible protein that turns from green to red after photo activation and makes it possible to follow cells precisely. The sox10: kaede transgenic line was used to perform lineage analysis to delineate CNC cell populations that give rise to maxillary versus mandibular elements and illustrate homology of facial prominences to amniotes. This protocol describes the steps to generate a live time-lapse video of a sox10: kaede zebrafish embryo. Development of the ethmoid plate will serve as a practical example. This protocol can be applied to making a time-lapse confocal recording of any kaede or similar photoconvertible reporter protein in transgenic zebrafish. Furthermore, it can be used to capture not only normal, but also abnormal development of craniofacial structures in the zebrafish mutants.

Visualization of experimental data has become a key element in presenting results to the scientific community. Generation of live time-lapse recording of growing embryos contributes to better presentation and understanding of complex developmental processes. This protocol is a step-by-step guide to cell labeling via photoconversion of kaede protein in zebrafish.

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, ...

4-Dimensional Imaging of Zebrafish Optic Cup Morphogenesis

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of visual impairment, yet mechanisms of eye morphogenesis are still poorly understood. The basic organization of the embryonic eye is conserved throughout vertebrates, thus live imaging of zebrafish embryos has become a powerful approach to directly observe eye development at real time under normal and pathological conditions. Dynamic cell processes including movements, morphologies, interactions, division, and death can be visualized in the embryo. We have developed methods for uniform labeling of subcellular structures and timelapse confocal microscopy of early eye development in zebrafish. This protocol outlines the method of generating capped mRNA for injection into the 1-cell zebrafish embryo, mounting embryos at optic vesicle stage (~12 hours post fertilization, hpf), and performing multi-dimensional timelapse imaging of optic cup morphogenesis on a laser scanning confocal microscope, such that multiple datasets are acquired sequentially in the same imaging session. Such an approach yields data that can be used for a variety of purposes, including cell tracking, volume measurements, three-dimensional (3D) rendering, and visualization. Our approaches allow us to pinpoint the cellular and molecular mechanisms driving optic cup development, in both wild type and genetic mutant conditions. These methods can be employed directly by other groups or adapted to visualize many additional aspects of zebrafish eye development.

This protocol describes an approach for in toto labeling and multidimensional imaging of zebrafish early eye development. We describe labeling, embedding, and four dimensional (4D) imaging using laser scanning confocal microscopy, and considerations for optimizing acquisition of datasets for dissecting mechanisms of optic cup morphogenesis.

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of ...