The authors thank Thomas Schilling for kindly gifting the Tg(sox10:eGFP) fish and Mary Halloran for kindly gifting the Tg(foxd3:GFP) fish.

The protocol described here was performed in accordance with the guidelines for the humane treatment of laboratory animals established by the University of Michigan Committee on the Use and Care of Animals (UCUCA).
1. Embryo Collection for Time-lapse Imaging
<ol>
	<li>Between 3 and 9 pm, set up male and female adult Tg(sox10:EGFP) or Tg(foxd3:GFP) transgenic zebrafish in a divided breeding tank for pairwise mating. 
	NOTE: The Tg(sox10:EGFP) and Tg(foxd3:GFP) fish, kind gifts from Thomas Schilling and Mary Halloran, respectively, were crossed into the Casper (roy -/-, nacre -/-) background to decrease auto-fluorescence and interference with pigmentation.
	<ol>
		<li>Assemble the breeding tank (slotted inner tank + solid outer tank) and fill it with reverse osmosis (RO) system water.</li>
		<li>Transfer up to 3 females and 3 males into opposing sides of the tank separated by a divider; up to 6 fish can be bred pairwise in a single breeding tank.</li>
		<li>The next morning, remove the divider shortly after the lights turn on in the vivarium. 
		NOTE: In the present study, the lights are on a 14-hour light (on at 9 am) and 10-hour dark (off at 11 pm) cycle.</li>
		<li>Allow undisturbed mating for 20 min or until sufficient numbers of embryos are produced; the embryos will be at the bottom of the breeding tank. When eggs are observed, lift the slotted inner tank out along with the fish and quickly place them (fish and inner tank) into a clean solid outer tank filled with RO water.</li>
	</ol>
	</li>
	<li>Prepare standard 1X embryo medium by adding the following per 1 L of RO water: 0.287 g (4.9 mM) NaCl, 0.127 g (0.17 mM) KCl, 0.048 g (0.33 mM) CaCl2.2H2O, 0.04 g (0.33 mM) MgSO4 (anhydrous). Stir solution until the salts are completely dissolved.
	<ol>
		<li>Add 100 &#181;L of 0.1% methylene blue as a fungicide. Add 0.25 g of sodium bicarbonate to adjust the pH to 7.75. Store medium at room temperature.</li>
	</ol>
	</li>
	<li>Collect eggs from the outer tank using an egg-collecting screen, and transfer the eggs to Petri dishes (~50 embryos per dish) containing 30 mL of 1X embryo medium. Incubate the collected eggs at 28.5 &#176;C.</li>
	<li>Embryo preparation
	<ol>
		<li>Tg(sox10:EGFP) embryos.
		<ol>
			<li>For Tg(sox10:EGFP) embryos, at 12 h post-fertilization (hpf), remove the dead eggs and assess the developmental stage of the embryos. Screen for GFP-positive embryos using a dissecting fluorescent stereomicroscope with a standard 460-490 nm band-pass excitation filter. Using forceps, remove the chorions from 3-4 embryos that express GFP, and transfer these embryos to a separate Petri dish containing 30 mL of fresh 1X embryo medium.</li>
		</ol>
		</li>
		<li>Tg(foxd3:GFP) embryos.
		<ol>
			<li>For Tg(foxd3:GFP) embryos, at 22-24 hpf, remove the dead eggs and assess the developmental stage of the embryos. Screen for positive embryos using a dissecting fluorescent stereomicroscope with a standard 460-490 nm band-pass excitation filter. Using forceps, remove the chorions from 3-4 embryos that express GFP.</li>
			<li>Prepare 1000X Phenylthiourea (PTU) stock solution by dissolving 0.75 g (3%) of PTU in 25 mL of Dimethylsulfoxide (DMSO). Aliquot and store the stock solution at 4 &#176;C. Add 30 &#181;L of 1000X PTU (3%) stock solution to 30 mL of 1X embryo medium to generate 0.003% PTU solution.</li>
			<li>Place the screened and dechorionated GFP-positive embryos in 0.003% PTU solution to inhibit pigmentation. Do not initiate treatment of the embryos with PTU prior to 20 hpf, as early treatment (at &#60;20 hpf) can have adverse effects on neural crest and neuroepithelial development.28</li>
		</ol>
		</li>
		<li>Prior to conducting the time-lapse experiment, monitor the embryos through live imaging using a standard stereomicroscope at 12-24 h intervals in age-matched comparisons as a control for temperature drift.</li>
	</ol>
	</li>
</ol>
2. Mounting of Embryo for Time-lapse Imaging
<ol>
	<li>Preparation of time-lapse embryo media
	<ol>
		<li>Prepare standard 0.4% tricaine stock solution by adding 0.004 g of tricaine to 100 mL of RO water. Aliquot (~1 mL) the stock solution into fresh 1.5-mL centrifuge tubes and store at -20 &#176;C.</li>
		<li>Prepare time-lapse embryo medium (50 mL of 1X embryo medium, 0.016% tricaine) by adding 2 mL of 0.4% tricaine solution to 48 mL of 1X embryo medium. If using embryos &#62;22 hpf, then also add 50 &#181;L of 3% PTU stock solution (final concentration.003% PTU) to the embryo medium.</li>
	</ol>
	</li>
	<li>Preparation of 2% low-melt agarose
	<ol>
		<li>Add 0.4 g of low-melt agarose powder to 20 mL of 1X embryo medium. Heat for 1-2 min or until the solution is clear and all particles are dissolved. 
		NOTE: Increasing the percentage of the agarose gel decreases the porosity of the matrix, which may physically impede the growth and development of the embryo. Therefore, the agarose solution can be lowered to 1.5% to minimize these effects. However, when the excitation beam is held stationary during laser-scanning microscopy, greater heating can occur, increasing rapidly in a logarithmic relationship with time. In this protocol, heating due to fluorophore absorption is highly localized to the focal region. Thus, in the region of interest, the temperature could increase to a value high enough (&#8805; 30 &#176;C) to melt agarose solutions made at percentages lower than 1.5, thereby directly exposing the embryo to the laser or enabling conditions in which the embryo floats out of the focal plane. For this reason, the use of agarose solutions lower than 1.5% is not recommended.</li>
		<li>Aliquot (~1 mL) the agarose solution into fresh 1.5-mL centrifuge tubes for storage. Store excess agarose solution in liquid form on a heat block (60-70 &#176;C) for 2-3 weeks.</li>
	</ol>
	</li>
	<li>Mounting the Embryo
	<ol>
		<li>To set up the open bath chamber, place a small amount of high vacuum grease on the base of the open bath chamber. Place a circular glass coverslip onto the base and screw the top of the open bath chamber onto the base until tight ( Figure 1A, B).</li>
		<li>Pipet (~500-700 &#181;L) of 2% agarose solution (60-70 &#176;C) into the open bath chamber until the base is ~3/4 filled ( Figure 1C). Note that once the agarose solution is in the open bath chamber on the lab bench, the temperature of the solution decreases approximately 1 &#176;C/s.</li>
		<li>Wait 30 s to allow the agarose to cool slightly (~30-40 &#176;C), without completely polymerizing, and subsequently transfer a single embryo to the center of the base ( Figure 1D, E).</li>
		<li>Under a fluorescent stereomicroscope, position the embryo using a 1-10 &#181;L micropipette tip, making sure that the embryo is placed near the bottom of the agarose, as the embryo may float away when covered with embryo media if it is placed too near the surface of the agarose. 
		NOTE: In the presented videos (Videos 1-3), the embryos are oriented laterally, but depending on the area of interest, the embryos can be oriented ventrally or dorsally.</li>
		<li>Monitor and reposition the embryo until the agarose has set. Once the agarose has set, use a transfer pipet to fill the assembled open chamber entirely with time-lapse embryo media ( Figure 1F).</li>
		<li>Place the open bath chamber in the quick exchange platform, which fits onto the stage adapter ( Figure 1G). Place the entire setup (embryo, open bath chamber, quick exchange platform and stage adaptor) onto the stage of the microscope immediately above the condenser ( Figure 1H).</li>
	</ol>
	</li>
</ol>
3. Microscope Set-up for Time-lapse Imaging
<ol>
	<li>Determining laser settings
	<ol>
		<li>Determining laser wavelength to use. Use a wavelength that is twice the excitation wavelength of the fluorophore of interest (e.g. wavelength between 880 and 940 nm for GFP). 
		NOTE: In the present study, the wavelength setting for GFP was between 880 and 940 nm. The higher the wavelength, the lower the output power of the laser.</li>
		<li>Determine laser transmission. A high percent of laser transmission will kill the embryo, use the lowest level of transmission (recommended). For 24 to 48 h time-lapse studies, as presented herein, keep transmission below 5%.</li>
		<li>Determine microscope detection systems and ensure that the correct filters for the fluorophore are in place. 
		NOTE: For multi-photon microscopes, there are multiple detection systems with various sensitivities for the emitted fluorescence. In general, an internal detection system has less sensitivity than an external detection system. For transgenic lines with high levels of GFP expression, the internal detection system is adequate. For transgenic lines with low levels of GFP expression or with other fluorophores (e.g., red fluorescent protein), an external detection system may be required to maintain the percent of laser transmission at a reasonable level. Regardless the detection system used, the correct filters for the fluorophore must be in place.</li>
	</ol>
	</li>
	<li>Adjusting software settings on the multi-photon microscope
	<ol>
		<li>Using the 5X objective, locate the embryo. Manually raise the stage to the highest position, and use the fine focus to position the embryo in the middle of the microscope range.</li>
		<li>Manually lower the stage, and change the 5X objective to the 25X water immersion objective (numerical aperture NA, 0.95). Carefully raise the stage to bring the embryo back into focus.</li>
		<li>In the software, click on the &#34;xyzt&#34; mode for obtaining multiple images at time intervals (t) in the x-y plane over a depth of &#34;z&#34;. Use the epifluorescence or brightfield view to find the depth of focus in the area of interest, which will demarcate the Z-stack.
		<ol>
			<li>In the software, click on &#34;begin&#34; button; for these experiments, the lateral edge of the eye was the beginning of the Z-stack. Click on &#34;end&#34; button; the midline of the embryo was the end of the Z-stack. 
			NOTE: The step size was 0.3-0.6 &#181;m and there was a total of ~200 steps for a z-stack size of 60 to 120 &#181;m).</li>
		</ol>
		</li>
		<li>Click on the menu for adjusting acquisition time and imaging frequency. For the present system, ~200 steps requires approximately 5 min for each z-stack acquisition. For adequate recovery of the fluorophore and survival of the embryo, allow for a ratio of at least 1:3 between z-stack acquisition (laser power on) and recovery time (laser power off). 
		NOTE: For example, z-stacks are acquired every 20 min with 5 min of z-stack acquisition and 15 min of recovery. For this protocol, larger z-stacks can be obtained, but would appropriately increase the time between z-stack acquisitions, resulting in fewer images over the time-lapse course.
		<ol>
			<li>With appropriate time for embryo recovery, set the time between z-stacks in the designated window. Set the total length of time for the experiment in the appropriate window.</li>
		</ol>
		</li>
		<li>Final software, laser, and embryo adjustments.
		<ol>
			<li>Turn on the live image setting to make final adjustments to the laser settings. Adjust laser transmission (see 3.1.2), gain and offset slider bars within the software to optimize the fluorescent image. Also, adjust the orientation of the embryo, as needed, depending on the length of the experiment, anticipated growth of the embryo, etc. Make sure that the area of interest remains within the frame through the duration of the experiment.</li>
			<li>Turn off the epifluorescent light source as it is no longer needed during time-lapse acquisition. Cover the stage with the laser safety box ( Figure 1I). When using the internal detection system, the laser safety box is adequate for protection against background light. Press &#34;start&#34;. 
			NOTE: However, with more sensitive external detection systems, the laser safety box does not block enough background light, and additional covers are required to prevent the disruption of image acquisition.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. During Time-lapse Acquisition
<ol>
	<li>Refill the open bath chamber with time-lapse embryo media every 8-12 h (at least 2 times per day) during time-lapse acquisition through the sliding doors on the laser safety box ( Figure 1I).</li>
	<li>Before opening the doors of the laser safety box, ensure that the microscope is not actively acquiring an image. 
	NOTE: The use of heaters and circulating media systems is not necessary for time-lapse imaging experiments lasting 24 to 48 h. Indeed, the temperatures of both stage and in-line heaters are difficult to control, and during image acquisition the embryo exhibits an adequate development rate at a temperature range from 25-28 &#176;C. Moreover, circulating media systems tend to overflow and potentially damage the equipment. Thus, all embryos are routinely staged post-acquisition.</li>
</ol>
5. Post-acquisition processing
<ol>
	<li>In the software, click on the &#34;file&#34; menu and choose &#34;save&#34;.</li>
	<li>Open the file in image processing software (see the Table of Materials). Highlight the correct image series. In the software, choose the &#34;Process&#34; menu. Click on 3D Deconvolution and &#34;Apply&#34; to deconvolve each z-stack. 
	NOTE: The file is large; therefore, this step may take many hours.</li>
	<li>In the software, under the &#34;Process&#34; menu, click on &#34;maximum projection.&#34; Click on &#34;Apply&#34; to initiate maximum projection to generate 1 image per z-stack. Export each maximum projected file (1 image per z-stack) as a tiff.</li>
	<li>Import individual tiff files into video processing software. Select all tiff files and drag them into the video editor. Adjust length of each image within the video to 0.1s. Export video as mov or mp4 file.</li>
</ol>

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C., Woodruff, E., Broadie, K., Appel, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CNS-derived+glia+ensheath+peripheral+nerves+and+mediate+motor+root+development.">CNS-derived glia ensheath peripheral nerves and mediate motor root development.</a> Nat Neurosci. 11, 143-151 (2008).</li><li>Denk, W., Strickler, J., Webb, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+laser+scanning+fluorescence+microscopy.">Two-photon laser scanning fluorescence microscopy.</a> Science. 248 (4951), 73-76 (1990).</li><li>Helmchen, F., Denk, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deep+tissue+two-photon+microscopy.">Deep tissue two-photon microscopy.</a> Nat Methods. 2 (12), 932-940 (2005).</li><li>Denk, W., Delaney, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anatomical+and+functional+imaging+of+neurons+using+2-photon+laser+scanning+microscopy.">Anatomical and functional imaging of neurons using 2-photon laser scanning microscopy.</a> J Neurosci Methods. 54 (2), 151-162 (1994).</li><li>Bohnsack, B. L., Gallina, D., Kahana, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenothiourea+sensitizes+zebrafish+cranial+neural+crest+and+extraocular+muscle+developmnt+to+changes+in+retinoic+acid+and+insulin-like+growth+factor+signaling.">Phenothiourea sensitizes zebrafish cranial neural crest and extraocular muscle developmnt to changes in retinoic acid and insulin-like growth factor signaling.</a> PLoS ONE. 6, e22991 (2011).</li><li>Gfrerer, L., Dougherty, M., Laio, E. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualization+of+craniofacial+development+in+the+sox10:kaede+transgenic+zebrafish+line+using+time-lapse+confocal+microscopy.">Visualization of craniofacial development in the sox10:kaede transgenic zebrafish line using time-lapse confocal microscopy.</a> J Vis Exp. (79), e50525 (2013).</li><li>Lopez, A. L. R., Garcai, M. D., Dickinson, M. E., Larina, I. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+confocal+microscopy+of+the+developing+mouse+embryonic+yolk+sac+vasculature.">Live confocal microscopy of the developing mouse embryonic yolk sac vasculature.</a> Methods Mol Biol. 1214, 163-172 (2015).</li><li>McGurk, P. D., Lovely, C. B., Eberhart, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analyzing+craniofacial+morphogenesis+in+zebrafish+using+4D+confocal+microscopy.">Analyzing craniofacial morphogenesis in zebrafish using 4D confocal microscopy.</a> J Vis Exp. (83), e51190 (2014).</li><li>Nowotschin, S., Ferrer-Vaquer, A., Hadjantonakis, A. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+mouse+development+with+confocal+time-lapse+microscopy.">Imaging mouse development with confocal time-lapse microscopy.</a> Methods Enzymol. 476, 351-377 (2010).</li></ol>

This work was financially supported through grants from the National Eye Institute of the National Institutes of Health (K08EY022912-01) and Vision Research Core (P30 EY007003).

Multi-photon time-lapse imaging enables the in vivo tracking of transient and migratory cell populations. This powerful technique can be used to study embryonic processes in real time, and in the present study, the results of this method enhanced the current knowledge of neural crest cell migration and development. Previous time-lapse imaging studies typically utilize confocal laser scanning microscopy.29,30,31,32 Here, we present the use of a multi-photon technique, which has many advantages over traditional confocal microscopy. The basis of multi-photon microscopy is that two photons of longer infrared wavelengths are used to excite the fluorophore. As a result, there is less scattering and thus decreased background, enabling deeper tissue penetration, achieving imaging depths exceeding 1 mm in biological tissue with more efficient detection than confocal microscopy. These properties also decrease phototoxicity, which is an important consideration with repeated and frequent image acquisition.25,26,27 From these properties, two-photon microscopy can image whole-organ preparations and tissue in small-animal models, (e.g. mice, zebrafish embryos at 12 hpf - as in the case of the present study). Further, these small animal models enable use of genetically encoded fluorescent proteins, which can be localized in the tissue of interest. Thus, the use of multi-photon microscopy can greatly enhance image acquisition for time-lapse experiments.
There are numerous steps in this protocol, which depend on the type of multi-photon laser and detection system being used. Most problems with this protocol arise with the laser settings and acquisition of images based on the detection system used. A working knowledge of the individual system and access to technical support are critical. In addition, previous experience with confocal laser scanning microscopy is helpful for troubleshooting. Moreover, the initial system set up may be time consuming. However, once the system set up is completed, only small adjustments are typically required when using the same transgenic lines.
In the present study, zebrafish embryos between 12 and 60 hpf were used for these experiments. During this age range, the embryos are small, easily embedded in agarose and readily anesthetized with tricaine. However, the embryos can become growth and developmentally delayed depending on the length of the experiment. Therefore, care must be taken to ensure that the embryo is appropriately staged at the end of the experiment.
Multi-photon time-lapse imaging yields high-resolution, three-dimensional, real-time visualization of cell migration, which not only enhances the capability to study in vivo zebrafish embryogenesis, but can also be applied to many other systems. Although the outlined protocol focuses on zebrafish neural crest cell migration, this technique can easily be adapted for other imaging purposes.

Congenital eye diseases can cause childhood blindness and are often due to abnormalities of the cranial neural crest. Neural crest cells are transient stem cells that arise from the neural tube and form numerous tissues throughout the body.1,2,3,4,5 Neural crest cells, derived from the prosencephalon and mesencephalon, give rise to the bone and cartilage of the midface and frontal regions, and the iris, cornea, trabecular meshwork, and sclera in the anterior segment of the eye.4,6,7,8 Neural crest cells from the rhombencephalon form the pharyngeal arches, jaw, and cardiac outflow tract.1,3,4,9,10 Studies have highlighted the contributions of the neural crest to ocular and periocular development, emphasizing the importance of these cells in vertebrate eye development. Indeed, disruption of neural crest cell migration and differentiation lead to craniofacial and ocular anomalies as observed in Axenfeld-Rieger Syndrome and Peters Plus Syndrome.11,12,13,14,15,16,17 Thus, a comprehensive understanding of the migration, proliferation and differentiation of these neural crest cells will provide insight into the complexities underlying congenital eye diseases.
The zebrafish is a powerful model organism for studying ocular development, as the structures of the zebrafish eye are similar to their mammalian counterparts, and many genes are evolutionarily conserved between zebrafish and mammals.18,19,20 In addition, zebrafish embryos are transparent and oviparous, facilitating the visualization of eye development in real-time.
Expanding on previously published work,6,7,20 the migratory pattern of neural crest cells was described using multi-photon fluorescence time-lapse imaging on transgenic zebrafish lines labeled with green fluorescent protein (GFP) under the transcriptional control of the SRY (sex-determining region Y)-box 10 (sox10) or Forkhead Box D3 (foxd3) gene regulatory regions.21,22,23,24. Multi-photon fluorescence time-lapse imaging is a powerful technique that combines the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation to capture high-resolution, three-dimensional images of specimens tagged with fluorophores.25,26,27 The use of the multi-photon laser has distinct advantages over standard confocal microscopy, including increased tissue penetration and decreased fluorophore bleaching.
Using this method, two distinct populations of neural crest cells varying in timing of migration and migratory pathways were discriminated, namely foxd3-positive neural crest cells in the periocular mesenchyme and developing eye and sox10-positive neural crest cells in the craniofacial mesenchyme. With this method, an approach to visualize the migration of ocular and craniofacial neural crest migration in zebrafish is introduced, making it easy to observe regulated neural crest migration in real time during development.
This protocol provides information for generating time-lapse videos during early eye development in Tg(sox10:EGFP) and Tg(foxd3:GFP) transgenic zebrafish, as an example. This protocol can be further applied for the high-resolution, three-dimensional, real-time visualization of the early development of any ocular and craniofacial structure derived from neural crest cells in zebrafish. Moreover, this method can further be applied for the visualization of the development of other tissues and organs in zebrafish and other animal models.

<table><tbody><tr><td>Breeding Tanks with Dividers</td><td>Aquaneering</td><td>ZHCT100</td><td>Crossing Tank Set (1.0-liter) Clear Polycarbonate with Lid and Insert</td></tr><tr><td>M205 FA Combi-Scope</td><td>Leica Microsystems CMS GmbH</td><td></td><td>Stereofluorescence Microscope - FusionOptics and TripleBeam</td></tr><tr><td>Sodium Chloride</td><td>Millipore (EMD)</td><td>7760-5KG</td><td>Double PE sack. CAS No. 7647-14-5, EC Number 231-598-3</td></tr><tr><td>Potassium Chloride</td><td>Millipore (EMD)</td><td>1049380500</td><td>Potassium chloride 99.999 Suprapur. CAS No. 7447-40-7, EC Number 231-211-8.</td></tr><tr><td>Calcium Chloride Dihydrate</td><td>Fisher Scientific</td><td>C79-500</td><td>Poly bottle; 500 g. CAS No. 10035-04-8</td></tr><tr><td>Magnesium Sulfate (Anhydrous)</td><td>Millipore (EMD)</td><td>MX0075-1</td><td>Poly bottle; 500 g. CAS No. 7487-88-9, EC Number 231-298-2</td></tr><tr><td>Methylene Blue</td><td>Millipore (EMD)</td><td>284-12</td><td>Glass bottle; 25 g. Powder, Certified Biological Stain</td></tr><tr><td>Sodium Bicarbonate</td><td>Millipore (EMD)</td><td>SX0320-1</td><td>Poly bottle; 500 g. Powder, GR ACS. CAS No. 144-55-8, EC Number 205-633-8</td></tr><tr><td>N-Phenylthiourea</td><td>Sigma</td><td>P7629-25G</td><td>&gt;98%. CAS Number 103-85-5, EC Number 203-151-2</td></tr><tr><td>Dimethylsulfoxide</td><td>Sigma</td><td>D8418-500ML</td><td>Molecular Biology grade. CAS Number 67-68-5, EC Number 200-664-3</td></tr><tr><td>Tricaine Methanesulfonate</td><td>Western Chemical Inc.</td><td>MS222</td><td>Tricaine-S</td></tr><tr><td>Low-Melt Agarose</td><td>ISC Bioexpress</td><td>E-3112-25</td><td>GeneMate Sieve GQA Low Melt Agarose, 25 g</td></tr><tr><td>Open Bath Chamber</td><td>Warner Instruments</td><td>RC-40HP</td><td>High Profile</td></tr><tr><td>Glass Coverslips</td><td>Fisher Scientific</td><td>12-545-102</td><td>Circle cover glass. 25 mm diameter</td></tr><tr><td>High Vacuum Grease</td><td>Fisher Scientific</td><td>14-635-5C</td><td>2.0-lb. tube. DOW CORNING CORPORATION&lt;br/&gt; 1658832</td></tr><tr><td>Quick Exchange Platform</td><td>Warner Instruments</td><td>QE-1</td><td>35 mm</td></tr><tr><td>Stage Adapter</td><td>Warner Instruments</td><td>SA-20LZ-AL</td><td>16.5 x 10 cm</td></tr><tr><td>TC SP5 MP multi-photon microscope</td><td>Leica Microsystems CMS GmbH</td><td></td><td></td></tr><tr><td>Mai Tai DeepSee Ti-Sapphire Laser</td><td>SpectraPhysics</td><td></td><td></td></tr><tr><td>Laser Safety Box</td><td>Leica Microsystems CMS GmbH</td><td></td><td></td></tr><tr><td>Leica Application Suite X (LAS X)&amp;nbsp; Software</td><td>Leica Microsystems CMS GmbH</td><td></td><td></td></tr><tr><td>Photoshop CS 6 Version 13.0 x64 Software</td><td>Adobe</td><td></td><td></td></tr><tr><td>iMovie Version 10.1.4 Software</td><td>Apple</td><td></td><td></td></tr></tbody></table>

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.

Multi-photon fluorescence time-lapse imaging generated a series of videos that revealed the migration patterns of cranial neural crest cells that give rise to the craniofacial structures and anterior segment of the eye in the Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish lines. As an example, sox10-positive neural crest cells between 12 and 30 hpf migrate from the edge of the neural tube into the craniofacial region (Video 1, Figure 2). The cells from the prosencephalon and mesencephalon migrate dorsal and ventral to the developing eye to populate the periocular mesenchyme. In addition, these sox10-positive cells form the frontonasal process. Neural crest cells from the rhombencephalon migrate ventrally and give rise to the pharyngeal arches. Time-lapse imaging of foxd3-positive neural crest cells between 30 and 60 hpf showed that these cells migrated between the optic cup and surface ectoderm and through the ocular fissure (Video 2, Figure 3). These foxd3-positive cells coalesced around the lens, forming the iris stroma.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56214/56214fig1.jpg" /> 
Figure 1.Setup. A. Each component of the embryo mounting apparatus, including the open bath chamber, quick exchange platform and stage adapter. B. Assembly of the open bath chamber. C. Addition of 2% agarose solution (60-70 &#176;C) to the open bath chamber. D. Placement of the embryo in the open bath chamber under a fluorescent microscope. E. Embryo (at 24 hpf) positioned laterally in the center of the polymerized agarose solution in the open bath chamber. F. Addition of media to the surface of the polymerized agarose solution to completely cover the embryo in the open bath chamber. G. Open bath chamber placed in the quick exchange platform and positioned in the stage adapter. H. Placement of the embryo mounting apparatus on the stage of the multi-photon microscope. I. Laser safety box. The sliding doors (1 and 2) for refilling the open chamber are indicated. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56214/56214fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56214/56214fig2.jpg" /> 
Figure 2. Representative deconvolved and max-projected images from multi-photon time-lapse imaging of a Tg(sox10:EGFP) zebrafish embryo from 12 to 30 hpf. The sox10-positive cells migrated from the prosencephalon (P) and mesencephalon (M) to populate the periocular mesenchyme (the dotted circle denotes the eye) and frontonasal process (open arrow). Sox10-positive cells from the rhombencephalon (R) migrated ventrally to form the pharyngeal arches (closed arrow). Images were obtained every 20 min during the time frame and sewn together to create Video 1. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56214/56214fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56214/56214fig3.jpg" /> 
Figure 3. Representative deconvolved and max-projected images from multi-photon time-lapse imaging of a Tg(foxd3:GFP) zebrafish embryo from 24 to 48 hpf. Foxd3-positive cells entered into the anterior chamber of the eye (asterisk denotes lens) between the surface ectoderm and optic cup, with more cells localized to the dorsal (&#34;D&#34;)-posterior (&#34;P&#34;) quadrant compared with the anterior (&#34;A&#34;) and ventral (&#34;V&#34;) quadrants. In addition, foxd3-positive cells migrated adjacent to and through the ocular fissure. Images were obtained every 20 min during the time frame and sewn together to create Video 2. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56214/56214fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56214/56214fig4.jpg" /> 
Figure 4. Representative deconvolved and max-projected images from multi-photon time-lapse imaging of a Tg(foxd3:GFP) zebrafish embryo from 30 to 60 hpf. Foxd3-positive cells entered into the anterior chamber of the eye (outer dotted circle denotes peripheral edges of the eye, inner dotted circle denotes lens) between the surface ectoderm and optic cup, with more cells localized to the dorsal (&#34;d&#34;)-posterior (&#34;p&#34;) quadrant compared with the anterior (&#34;a&#34;) and ventral (&#34;v&#34;) quadrants. In addition, foxd3-positive cells migrated adjacent to and through the ocular fissure (white arrowhead denotes migrating neural crest, red dashed line demarcates ocular fissure). At 60 hpf, foxd3-positive cells completely encircled the lens (closed arrows), indicating the closure of the fissure. In addition, at 60 hpf, foxd3 was also expressed in photoreceptors (open arrowhead), which was not associated with the expression of this transcription factor in neural crest cells. Images were obtained every 20 min during the time frame and sewn together to create Video 3. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56214/56214fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Video 1" src="/files/ftp_upload/56214/56214vid1.jpg" /> 
Video 1. Time-lapse video of a Tg(sox10:EGFP) zebrafish embryo from 12 to 30 hpf. The asterisk denotes the eye. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56214/12h_sox10_gfp.mp4" target="_blank">Please click here to download this video.</a>
<img alt="Video 2" src="/files/ftp_upload/56214/56214vid2.jpg" /> 
Video 2. Time-lapse video of a Tg(foxd3:GFP) zebrafish embryo from 24 to 48 hpf. The asterisk denotes the lens. The asterisk denotes the ocular fissure. d, dorsal; v, ventral; p, posterior; a, anterior. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56214/24h_foxd3_gfp.mp4" target="_blank">Please click here to download this video.</a>
<img alt="Video 3" src="/files/ftp_upload/56214/56214vid3.jpg" /> 
Video 3. Time-lapse video of a Tg(foxd3:GFP) zebrafish embryo from 30 to 60 hpf. The asterisk denotes the lens. The arrow denotes the ocular fissure. d, dorsal; v, ventral; p, posterior; a, anterior. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56214/30h_foxd3_gfp.mp4" target="_blank">Please click here to download this video.</a>

Watch this Scientific Journal Video about Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos at JoVE.com

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

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 ...

multi-photon-time-lapse-imaging-to-visualize-development-real-time

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

7.6K Views.  University of Michigan. Een combinatie van de geavanceerde optische technieken van laser scanning microscopie met lange golflengte fluorescentie multi foton excitatie werd uitgevoerd om vast te leggen van real-time, hoge resolutie, driedimensionale beeldvorming van neurale crest migratie in GS (sox10:EGFP) en GS (foxd3:GFP) zebrafish embryo's.

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

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Neuroscience

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

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. ...

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Biology

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

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 ...

Multicolor Time-lapse Imaging of Transgenic Zebrafish: Visualizing Retinal Stem Cells Activated by Targeted Neuronal Cell Ablation

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound transgenic fish which express different fluorescent reporters in neighboring cell types provide a means of following cellular interactions 2 and/or tissue-level responses to experimental manipulations over time. In this video, we demonstrate methods that can be used for imaging multiple transgenically labeled cell types serially in individual fish over time courses that can span from minutes to several days. The techniques described are applicable to any study seeking to correlate the "behavior" of neighboring cells types over time, including: 1) serial 'catch and release' methods for imaging a large number of fish over successive days, 2) simplified approaches for separating fluorophores with overlapping excitation/emission profiles (e.g., GFP and YFP), 3) use of hypopigmented mutant lines to extend the time window available for high-resolution imaging into late larval stages of development, 4) use of membrane targeted fluorescent reporters to reveal fine morphological detail of individual cells as well as cellular details in larger populations of cells, and 5) a previously described method for chemically-induced ablation of transgenically targeted cell types; i.e., nitroreductase (NTR) mediated conversion of prodrug substrates, such as metronidazole (MTZ), to cytotoxic derivatives 3,5.
As an example of these approaches, we will visualize the ablation and regeneration of a subtype of retinal bipolar neuron within individual fish over several days. Simultaneously we will monitor several other retinal cell types, including neighboring non-targeted bipolar cells and potential degeneration-stimulated retinal stem cells (i.e., M&#971;ller glia). This strategy is being applied in our lab to characterize cell- and tissue-level (e.g., stem cell niche) responses to the selective loss and regeneration of targeted neuronal cell types.

In this video, techniques for multicolor confocal time-lapse imaging and targeted cell ablation are provided. Time-lapse imaging is used to monitor the behavior of multiple cell types of interest in vivo. Targeted cell ablation facilitates the study neural circuit function and cell-specific neuronal regeneration paradigms.

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound ...

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.

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 ...

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 ...

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

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