Research reported in this publication was supported by the Office of The Director of the National Institutes of Health (NIH) under Award Number S10OD020067 and by NIH award R01EY021769 (to A.C.M.). We are grateful for the assistance of Doug Harrison and Jim Begley in the Arts &#38; Sciences Imaging Center at the University of Kentucky, and to Lucas Vieira Francisco and Evelyn M. Turnbaugh for expert zebrafish care.

All experiments involving the use of zebrafish were carried out in accordance with protocols established by the University of Kentucky Institutional Animal Care and Use Committee (IACUC).
1. Sample preparation
<ol>
	<li>Set up a mating pair of Tg(Rx3:GFP) zebrafish in a gated cross tank the night before imaging is planned. The following morning pull the gate as the lights come on in the fish facility34,35. 
	NOTE: The mating pair of fish selected should be between 4 months to 2 years in age and are easily distinguished as male or female based on body shape and coloration35.</li>
	<li>Check the crosses every half hour for embryos and note what time embryos are first visualized in the bottom of the cross tank. Transfer the embryos to a Petri dish and maintain the embryos at 28.5 &#176;C for approximately 10 h to develop to the 1-2 somite stage (ss). Begin to image at the 1-2 somite stage (ss).</li>
	<li>To screen for the presence of somites, observe the embryos under a stereoscope at 10 hpf and count the number of somites1. 
	NOTE: Any embryos that are beyond the single somite stage should not be used for this experiment.</li>
	<li>Out of the embryos that are at the single somite stage, screen for GFP expression using a fluorescence adapter in combination with a stereomicroscope to confirm the presence of the Rx3:GFP transgene. Once 3-5 GFP positive individuals have been identified, use fine forceps to dechorionate the embryos and transfer them into a small Petri dish containing E3 embryo buffer with a glass pipette35.</li>
	<li>Anesthetize the embryos by transferring them into 0.5 mL E3 embryo buffer (pH 7) containing 0.168 mg/mL Tricaine (MS222) in a microcentrifuge tube35,36. Spontaneous muscle movements begin as early as 17 hpf37. Ensure that embryos are anesthetized to image beyond this timepoint.</li>
	<li>Embed the embryos in 1% low melting temperature agarose in 0.168 mg/mL Tricaine and E3. 
	NOTE: This is the optimal concentration of low melting temperature agarose to allow for the embryo to be held in place but remain pliable enough to permit the growth of the embryo over the imaging time-course30,31.</li>
	<li>Prepare a 10 mL solution of 2% low melting temperature agarose in E3 buffer. Heat it in a microwave oven to dissolve agarose using 15 s intervals, to prevent the solution from boiling over. Allow the solution to cool enough to hold without discomfort but not so much that it solidifies, and not cause harm to the embryos. Once the agarose is sufficiently cool, add 0.5 mL to the embryos in the tube of Tricaine in E3 and gently pipet the solution and embryos to mix.</li>
	<li>Using a 1 mm glass capillary and polytetrafluoroethylene (PTFE) plunger, pull up the embryos in the agarose solution into the capillary. Make sure to pull a total of 3-5 embryos (multiple embryos) into the capillary to increase the likelihood of having a well-positioned embryo. 
	NOTE: Due to the round nature of the embryos at this timepoint, it is challenging to guarantee a specific orientation in the capillary. Ideally, the body of the embryo will be positioned laterally in the capillary, allowing for the greatest ease of positioning within the microscope.</li>
	<li>Let the agarose solidify over a period of 30-60 s at room temperature. Place the capillary in a beaker of 0.168 mg/mL Tricaine in E3 buffer until ready to image (Figure 1A). 
	NOTE: The excess 2% low melting temperature agarose solution can be allowed to solidify and subsequently reheated in future experiments.</li>
	<li>Place the capillary into the sample holder (Figure 1B-F) as described in step 2.5.</li>
</ol>
2. Zeiss Lightsheet Z.1 setup
<ol>
	<li>Switch on each component of the microscope and the computer in the following order: 1) System, 2) PC, then 3) Incubation (Figure 2A).</li>
	<li>Place the 20x imaging objective and the 10x illumination objective into the microscope chamber. Match the objective settings in the Zen Software interface under the Maintain tab.</li>
	<li>Slide the chamber (Figure 2C) into the housing on the track (Figure 2B) with the tubing facing out. The tubing connects to the appropriate ports on the right as shown in Figure 2E.</li>
	<li>Attach the extension line to the syringe with the luer-lock mechanism (Figure 2D); fill it with Tricaine in E3 and place it in the holder attached to the right of the microscope35. Connect the extension attached to the syringe filled with Tricaine in E3 to the bottom right of the chamber with the luer-lock mechanism. Push the plunger to fill the chamber with the Tricaine/E3 buffer. Close the door to the chamber.</li>
	<li>Place the capillary with the sample into the capillary sample holder. The capillary sample holder comprises two rubber sleeves, a metal sample holder disc, a metal stem, and a metal cap (Figure 1B). Click the metal sheath into the center of the metal disc (Figure 1C). Place the two rubber stoppers into the sheath, with the slits facing the ends of the sheath followed by the capillary. Slide the capillary through the middle of the rubber stoppers. Fasten it in place with the metal cap once the marker is at the base of the metal sheath (Figure 1D).</li>
	<li>Place the capillary sample holder onto the top of the microscope, with the white marks aligning (Figure 1E,F). Close the lid.</li>
	<li>Click on the Locate Capillary button on the software interface (Figure 3A). Use the ErgoDrive control panel, a manual device that controls the capillary orientation (Figure 3E), to move the capillary and position it just above the objective (Figure 3B).</li>
	<li>Open the lid, and gently push on the plunger until the section of agarose containing the embryo is hanging below the capillary bottom and is in front of the objective (Figure 3C).</li>
	<li>Turn off Locate Capillary and click on the Locate Sample button (Figure 3A). This switches the view from the sample chamber&#39;s webcam to the microscope objective (Figure 3D). Use this view to adjust the position of the sample more precisely. Turn off Locate Sample (Figure 3A).</li>
	<li>Switch over to the Acquisition tab. Check the boxes for Z-Stack and Time Series.</li>
	<li>In the Acquisition Mode parameters window, choose the Dual Side Lightsheet setting and check the boxes for Online Dual Side Fusion and Pivot Scanning.</li>
	<li>In the Channels window, choose 488 channel, set the laser power to 1, and the exposure time to 7.5 ms.</li>
	<li>Next, click on the Continuous button to get a live view of the embryo. Use the ErgoDrive control panel to adjust the position of the embryo until the eye field is directly facing the camera. Continue adjusting the left and right lightsheets in the Channels parameters until the eye field is sufficiently in focus.</li>
	<li>Set the Z-Stack parameters using the ErgoDrive control panel to move through the Z-plane. Set the first and the last Z-Positions around 500 &#181;m beyond the last detectable fluorescent signal. This leaves room for the eye field to remain in frame as the embryo grows throughout the time-lapse imaging session. After setting the range of the Z-Stack, click on the Optimal button to set the step size to 0.477 &#181;m, the optimal setting.</li>
	<li>Set the Incubation parameters by checking the box for the Peltier Unit to keep the temperature at 28 &#176;C. In the Time Series window, choose the frequency and time interval to acquire images. In this protocol, the parameters were set to image every 5 min, for a total of 166 intervals.</li>
	<li>Click on the Start Experiment button. Choose the folder to save the image set. Set the image prefix and hit Save to start the imaging. 
	NOTE: The microscope will now run through each image set at the interval specified.</li>
	<li>After the time-lapse imaging session has been completed, send the stage to the load position, and remove the capillary sample holder. Take apart the capillary sample holder in the reverse order that it was put together and use the plunger to remove the sample and excess agarose from the capillary. Open the chamber door. Use the syringe to remove the chamber liquid from the chamber, disconnect and remove the chamber, then rinse with water and air dry.</li>
</ol>
3. Image analysis
<ol>
	<li>Open the arivis Vision4D software program.</li>
	<li>Click on File, then choose Import file. Select all .czi files from the time-lapse imaging session and open. The next window opens with the options on how to import the files; select Z-stacks as Frames to order the z-stacks in the order obtained. Once the files have been imported, the software saves as a single .sis file. To specify the location for the file to be saved, select the folder prior to clicking on Import.</li>
	<li>After the file is imported, arivis is now ready to render a video of the time-lapse images. Click on the 4D Viewer Cube in the bottom-left corner and the Scale Bar icon (Figure 4D-E). Click on the video icon to bring the Storyboard taskbar to the bottom of the screen (Figure 4C).</li>
	<li>In the Storyboard taskbar, choose Add Keyframe Sequence (Figure 4F). Specify the duration of the video in seconds, uncheck the Create Rotation box, and check the Use Time Progression to include the specific timepoints in the video. The software displays these parameters to the right of the Storyboard taskbar for any adjustments at any time. Save the Storyboard to apply the same parameters to multiple image sets (Figure 4F).</li>
	<li>Click on Export Movie to save a video of the time-lapse imaging (Figure 4F). Specify the movie export settings, including the file name and location, video format (.mp4), video resolution (1,080 p), framerate (60 FPS), and data resolution (1,297 x 1,297 x 784). Add timestamps here, if desired. Once these parameters are set, choose Record (Figure 4F).</li>
	<li>Steps 3.4 and 3.5 can be repeated with modification to Step 3.4 to create rotation videos at specific timepoints. When adding a keyframe sequence to the storyboard, check the Create Rotation box, and uncheck Use Time Progression. Then, follow the same instructions as in step 3.5.</li>
	<li>To render a high-resolution image at any orientation at any individual time point, select the Camera icon in the 4D Viewer to obtain a high-resolution image (Figure 4C).</li>
	<li>To build a pipeline for analysis, choose the Flask icon to access the Analysis panel (Figure 4B). In the Analysis panel, click on the Analysis Operations dropdown menu. As an example, the protocol demonstrates below the sequence of operations for a volume analysis pipeline. Run or undo each step of the pipeline individually to fine tune each parameter (Table 1).</li>
	<li>Click on the blue triangle at the top of the pipeline to allow the pipeline to run. 
	NOTE: This will take some time depending on the speed of the computer. Once the pipeline has been optimized, the pipeline can be exported and imported to arivis with ease and applied to multiple datasets in a batch analysis.</li>
	<li>To access batch analysis, click on Batch Analysis under the Analysis tab.</li>
	<li>After the pipeline runs, a window pulls up with all of the objects found. Access this manually through the Table icon (Figure 4B). At the top of this window, click on the box labeled Feature Columns to pull up a list of features that can provide information about the object of interest. For the eye field volume analysis, these features include Surface Area, Volume (Volume, VoxelCount), Intensities #1 (Mean), Attributes (Id, Type), and Time Point (First). Click on Export to export the data to a spreadsheet.</li>
</ol>

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A Guide for the laboratory use of zebrafish (Danio rerio)., 5th Edition. , (2007).</li><li>Hirsinger, E., Steventon, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+versatile+mounting+method+for+long+term+imaging+of+zebrafish+development.">A versatile mounting method for long term imaging of zebrafish development.</a> Journal of Visualized Experiments: JoVE. (1199), e55210 (2017).</li><li>Saint-Amant, L., Drapeau, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time+course+of+the+development+of+motor+behaviors+in+the+zebrafish+embryo.">Time course of the development of motor behaviors in the zebrafish embryo.</a> Journal of Neurobiology. 37 (4), 622-632 (1998).</li><li>Cavodeassi, F., Ivanovitch, K., Wilson, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eph/Ephrin+signalling+maintains+eye+field+segregation+from+adjacent+neural+plate+territories+during+forebrain+morphogenesis.">Eph/Ephrin signalling maintains eye field segregation from adjacent neural plate territories during forebrain morphogenesis.</a> Development (Cambridge). 140 (20), 4193-4202 (2013).</li><li>Muthu, V., Eachus, H., Ellis, P., Brown, S., Placzek, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rx3+and+Shh+direct+anisotropic+growth+and+specification+in+the+zebrafish+tuberal/anterior+hypothalamus.">Rx3 and Shh direct anisotropic growth and specification in the zebrafish tuberal/anterior hypothalamus.</a> Development (Cambridge). 143 (14), 2651-2663 (2016).</li><li>Rojas-Mu&#241;oz, A., Dahm, R., N&#252;sslein-Volhard, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=chokh/rx3+specifies+the+retinal+pigment+epithelium+fate+independently+of+eye+morphogenesis.">chokh/rx3 specifies the retinal pigment epithelium fate independently of eye morphogenesis.</a> Developmental Biology. 288 (2), 348-362 (2005).</li></ol>

The authors declare no competing financial interests.

In this protocol, the Lightsheet microscope was used to perform time-lapse imaging of eye development and the resulting data were analyzed. The resulting dataset can provide valuable insights into the process of ocular morphogenesis, as well as perturbations to this process as a result of genetic mutation, exposure to pharmacological agents, or other experimental parameters. Here the protocol demonstrated how this dataset can be obtained and provided an example of how to analyze the volume of the eye field through early development. This data was found to be reproducible and consistent (less than 10% variation in volume) across biological replicates, bearing in mind that slight differences in embryo staging prior to the start of the run can lead to some variation in final volume measurements.
Care should be taken in the initial positioning of the embryo in the capillary and in positioning the embedded embryo in front of the objective. Orientation plays an important role in preventing the embryo from growing and moving out of the view of the objective. The embryos have a round shape at 10 hpf, which makes it challenging to guarantee a specific orientation in the capillary. Ideally, the body of the embryo will be positioned laterally in the capillary. Loading multiple embryos in the capillary will increase the likelihood of having a well-positioned embryo.
In this procedure, the embryo is embedded in agarose in order to suspend it in front of the imaging and illumination objectives. Choosing the correct concentration of the low melting temperature agarose is critical. Too high of a concentration will constrict the embryo and prevent it from properly developing; too low of a concentration will result in the agarose falling apart and not holding the embryo. The concentration optimal for this protocol is a final concentration of 1% low melting temperature agarose30,31.
Another element that should be taken into consideration is the level of saturation. As the eyefield grows and differentiates, the strength of the Rx3:GFP signal intensifies. Therefore, when setting the initial imaging parameters, the exposure and laser power should be reduced to undersaturate the image. This will prevent the image from becoming oversaturated as the Rx3:GFP gets brighter over time. Modifications can be made to correct for undersaturation in image processing but oversaturation cannot be corrected after the images have been acquired.
There are a few additional modifications that can be made to this protocol that may be advantageous to some projects that are not described in this paper. For example, it is possible to set up Multiview imaging in the image acquisition set up. This parameter would allow multiple embryos at different positions along the y-axis to be sequentially imaged at each time interval. While adding complexity to the data set, it would increase the rate of data collection. Additionally, in terms of image processing, it is possible to quantify the eye field by other parameters. Here, we described how to quantify the data in terms of the eye field volume. Alternatively, a pipeline could be made to quantify and track individual cells or determine the rate of optic vesicle evagination.
As previously mentioned, both confocal and Lightsheet microscopy have been used to perform time-lapse imaging studies of zebrafish. Lightsheet was specifically chosen for this project due to its superior ability to image through a thick (&#62;1 mm) sample, because it is equipped with an incubation unit to maintain an ideal temperature environment for the zebrafish embryo, and because its ability to image at a faster rate than confocal microscopy allows for image acquisition at the numerous time intervals required for this protocol no accompanying damage or photobleaching of the embryo21,22,23,24,25,26,27,28,29,30,31. It is also important to note that the Lightsheet microscope is equipped to image the signal from multiple fluorophores. The Lightsheet microscope used in this study has solid state laser excitation lines at 405, 445, 488, 515, 561, and 638 nm, which could be useful for imaging transgenic embryos expressing more than one fluorescent reporter transgene.
While this protocol details instructions for image acquisition analysis specifically using the Lightsheet Z.1 Dual Illumination Microscope System and arivis Vision4D analysis software, there are other commercially available Lightsheet microscopes made by Leica, Olympus, and Luxendo, as well as image analysis software by Imaris, that could be used to achieve similar results. The selection of equipment and software for this protocol was determined by the availability at our institution.
In summary, it is anticipated this protocol will provide a solid starting point for conducting time-lapse imaging using Lightsheet microscopy, and for image quantification of early eye development in zebrafish.

Embryonic development is a complex process that requires the precise coordination of many different events. The formation of the vertebrate eye begins in the developing forebrain, where a portion of the cells are specified as the eye field. These cells will evaginate toward the surface ectoderm, giving rise to two bilateral optic vesicles1,2,3,4,5,6,7,8,9,10. Contact with the surface ectoderm then induces an invagination of the optic vesicle into an optic cup. The surface ectoderm will give rise to the anterior structures of the eye, such as the lens and cornea, while the optic cup will give rise to the neural retina and retinal pigmented epithelium1,2,3,5,6,7,8,9,10,11,12. Disruptions in this process can lead to congenital defects such as microphthalmia, anophthalmia, and coloboma (MAC). At this time, there are no options to correct these defects3,5,6,7,9,10,11,12. Further studies of the mechanisms of ocular morphogenesis and the problems that can lead to MAC will provide a foundation of knowledge that will potentially lead to treatments. One powerful tool to investigate the dynamic behaviors of cells during eye development is time-lapse imagining, which allows this process to be visualized and characterized in vivo and in real time.
Zebrafish (Danio rerio) are an excellent model to visualize early ocular development using time-lapse imaging. They have a highly conserved vertebrate eye and possess the ability to develop rapidly and externally while remaining optically transparent1. Zebrafish provide a great resource for time-lapse imaging due to these characteristics that mammalian models lack. Time-lapse imaging studies of zebrafish eye development are greatly facilitated by use of the transgenic zebrafish line Tg(rx3:GFP)13. Rx3 (Retinal homeobox protein 3) is a transcription factor essential for eye development14. Rx3 is the first of the three rx genes in the zebrafish to be expressed, starting its expression mid-gastrulation, approximately 8 h post fertilization (hpf)15,16. The rx3:GFP transgene can be visualized in the developing forebrain starting at the 1 somite stage (ss), approximately 10 hpf15,17,18,19,20. In the developing forebrain, rx3:GFP expression marks the cells of the single eye field, and GFP (green fluorescent protein) continues to be expressed through the remainder of ocular morphogenesis. High resolution time lapse imaging of rx3:GFP expression, therefore, allows us to track the single eye field through time as it develops into the retina17,18,20.
Time-lapse imaging studies of zebrafish development have primarily been performed using confocal or Lightsheet microscopy. Confocal microscopy is advantageous in that it allows for the precise imaging of samples along the z-axis and reduces fluorescent background signal. However, it is limited by the amount of time it takes to acquire an image, sample position rigidity, and its propensity toward photobleaching and phototoxicity of live samples. Lightsheet microscopy is an ideal method to image ocular morphogenesis over time due to its ability to penetrate thicker samples for fluorescent imaging, increased flexibility in sample orientation, minimized photobleaching and phototoxicity, and imaging at a high speed21,22,23,24,25,26,27,28,29,30,31. The spatial resolution achievable with current light sheet microcopy systems is approximately 250-500 nanometers (nm). Although this is not significantly different from what can be obtained with confocal microscopy, the sample can be freely rotated and imaged from multiple angles, improving both imaging depth and resolution, and offering much greater flexibility for in vivo time lapse imaging experiments than the confocal platform27,32. For these reasons, Lightsheet microscopy is quickly becoming the favored method for time-lapse imaging studies of zebrafish development. This protocol describes the steps of quantifying oculogenesis through the imaging of Rx3:GFP transgenic zebrafish using a commercially available Lightsheet microscope33&#160;and details a pipeline for image analysis using the arivis software platform.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>60mL Syringe Luer-Lok Tip</td>
			<td>BD</td>
			<td>309653</td>
		</tr>
		<tr>
			<td>Agarose, Low Melting Temperature</td>
			<td>Promega</td>
			<td>V2111</td>
		</tr>
		<tr>
			<td>arivis Vision4D Software</td>
			<td>arivis</td>
			<td>https://www.arivis.com/en/imaging-science/arivis-vision4d</td>
		</tr>
		<tr>
			<td>Dumoxel N3C Forceps</td>
			<td>Dumont</td>
			<td>0203-N3C-PO</td>
		</tr>
		<tr>
			<td>E3 fish buffer (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Capillary &#8211; Size 2 (~1mm)</td>
			<td>Zeiss</td>
			<td>Black/701904</td>
		</tr>
		<tr>
			<td>Heidelberger Extension Line 100cm</td>
			<td>B. Braun</td>
			<td>4097262</td>
		</tr>
		<tr>
			<td>Light &#38; Filter Set &#8211; Royal Blue</td>
			<td>Night Sea</td>
			<td>SFA-LFS-RB</td>
		</tr>
		<tr>
			<td>Petri Dish (100 x 15 mm)</td>
			<td>VWR</td>
			<td>25384-302</td>
		</tr>
		<tr>
			<td>Stereomicroscope Fluorescence Adapter</td>
			<td>NightSea</td>
			<td></td>
		</tr>
		<tr>
			<td>Teflon Tipped Plunger &#8211; Size 2</td>
			<td>Zeiss</td>
			<td>#701997</td>
		</tr>
		<tr>
			<td>Tg(rx3:GFP) zebrafish</td>
			<td></td>
			<td>This line was established by Rembold et al.13</td>
		</tr>
		<tr>
			<td>Tricaine (MS222)</td>
			<td>Sigma</td>
			<td>A-5040</td>
		</tr>
		<tr>
			<td>Z.1 Lightsheet</td>
			<td>Zeiss</td>
			<td></td>
		</tr>
		<tr>
			<td>Zen Software</td>
			<td>Zeiss</td>
			<td>https://www.zeiss.com/microscopy/us/products/microscope-software/zen.html</td>
		</tr>
	</tbody>
</table>

visualizing ocular morphogenesis by lightsheet microscopy using rx3:gfp transgenic zebrafish

Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, proliferation, and differentiation. Time-lapse imagining offers unique insight to the behavior of cells during eye development because it allows us to visualize oculogenesis in vivo. Zebrafish are an excellent model to visualize this process due to their highly conserved vertebrate eye and their ability to develop rapidly and externally while remaining optically transparent. Time-lapse imaging studies of zebrafish eye development are greatly facilitated by use of the transgenic zebrafish line Tg(rx3:GFP). In the developing forebrain, rx3:GFP expression marks the cells of the single eye field, and GFP continues to be expressed as the eye field evaginates to form an optic vesicle, which then invaginates to form an optic cup. High resolution time lapse imaging of rx3:GFP expression, therefore, allows us to track the eye primordium through time as it develops into the retina. Lightsheet microscopy is an ideal method to image ocular morphogenesis over time due to its ability to penetrate thicker samples for fluorescent imaging, minimize photobleaching and phototoxicity, and image at a high speed. Here, a&#160;protocol is&#160;provided&#160;for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software. The resulting dataset can provide valuable insights into the process of ocular morphogenesis, as well as perturbations to this process as a result of genetic mutation, exposure to pharmacological agents, or other experimental manipulations.

The dataset displayed here was imaged using the protocol described above. A Tg(Rx3:GFP) embryo was imaged starting at the 1 somite stage (ss) through 24 hpf, a total time period of 14 h, with the images acquired at 5 min intervals. Time-lapse imaging allows for easy selection and comparison of any time-point that shows a phenotype of interest. Figure 5 demonstrates a set of high-resolution images that were rendered from the dorsal vantage point at select developmental time points. The pipeline run in arivis Vision4D builds a mask that represents the developing eye as identified by fluorescent signal. In Figure 5 and Videos 1-6, the mask can be visualized in comparison to the fluorescent rendering of the developing eye. Additionally, Table&#160;2 displays the volume data from the developing eye at every imaging point. This dataset includes the segment name, id, volume in both &#181;m3 and voxel count, the mean fluorescence intensity of the object, the time point the object was identified, and the object&#39;s surface area in &#181;m2. It is important to note that when the eye field separates into two optic vesicles (starting at Timepoint 64), there remains a third region that is Rx3:GFP positive in the forebrain, which will contribute to the hypothalamus38,39,40&#160;(Figure 5D-M). This shows up in the volume data represented in Table 2 (highlighted in yellow starting at Timepoint 71) and can easily be separated out from the optic vesicles, since it is much smaller in volume than either optic vesicle.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62296/62296fig01.jpg" /> 
Figure 1: Sample preparation. (A) Positioning of embryos in a glass capillary. The arrow points to an embryo in the capillary. (B) Glass capillary and capillary holder parts. (C) Partially assembled capillary holder. (D) Fully assembled capillary holder. (E) Lightsheet mounting chamber. The arrow indicated the white line used to orient the capillary holder. (F) Capillary holder properly mounted in the Lightsheet. The arrow shows the matching white lines, indicating proper orientation of the capillary holder. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/62296fig01large.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/62296/62296fig02.jpg" /> 
Figure 2: Lightsheet imaging set-up. (A) Switchboard to turn on the Lightsheet, computer, and incubation unit. The numbers indicate the order of operations. (B) Lightsheet objective chamber. (C) Imaging chamber. (D) Syringe and tubing that will be connected to the imaging chamber. (E) The imaging chamber properly positioned within the objective chamber with all of the tubes connected to the appropriate ports to the right. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/62296fig02large.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/62296/62296fig03.jpg" /> 
Figure 3: Sample positioning. (A) Locate Capillary and Locate Sample buttons in Zen Software as indicated by the arrows. (B) The positioning on the glass capillary. The arrow indicates the edge of the glass capillary positioned just above the lens of the objective. (C) The embryo suspension beyond the glass capillary. The arrow indicated the embryo suspended in agarose beneath the glass capillary in front of the objective&#39;s lens. (D) View of the embryo through the objective. (E) The ErgoDrive control panel. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/62296fig03large.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/62296/62296fig04.jpg" /> 
Figure 4: Important icons for navigating arivis Vison4D. Each panel has the icon function identified from left to right. (A) Open, Save, Close. (B) Analysis Panel, Show Objects Table, Open Track Editor. (C) Copy current viewer content as an image into clipboard, Toggle Bookmarks, Create a high-resolution image for the current view, Toggle Storyboard. (D) Show Measure Box, Show Orientation Cross, Show Legend, Show Scale Bar. (E) Show as 2D Viewer, Show as Gallery Viewer, Show as 4D Viewer, Show as Info Viewer, Show as Projection Viewer. F) Refresh all Keyframes, Add Keyframe, Add Keyframe sequence, Insert Keyframe, Remove all Keyframes, Export Movie, Load Storyboard, Save Storyboard, Adjust the target time of the entire movie, First Keyframe, Play, Pause, Stop, Last Keyframe. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/62296fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62296/62296fig05.jpg" /> 
Figure 5: High-resolution images and eye field masks. (A-M) A set of high-resolution images that were rendered from the dorsal vantage points; (A&#39;-M&#39;) the eye field masks for each corresponding timepoint. Each image set is notated by the time it was acquired from the start of imaging and the corresponding developmental stage in either somite stage (ss) or hours post fertilization (hpf). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/62296fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Time-lapse video of Tg(rx3:GFP) zebrafish embryo from 1 ss-24 hpf.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Video1_timelapse_rx3-GFP_1ss-24hpf MED RES.mov" target="_blank">Please right click here to download the video (save link as).</a>
Video 2: Time-lapse video of Tg(rx3:GFP) zebrafish embryo from 1 ss-24 hpf with the eyefield as identified by the arivis Vision4D pipeline.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Video2_timelapse_rx3-GFP_eyefieldmask_1ss-24hpf MED RES.mov" target="_blank">Please right click here to download the video (save link as).</a>
Video 3: 360&#176; rotation of a Tg(rx3:GFP) zebrafish embryo at 1 ss.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Video3_360-rotation_rx3-GFP_1ss(1).mp4" target="_blank">Please right click here to download the video (save link as).</a>
Video 4: 360&#176; rotation of a Tg(rx3:GFP) zebrafish embryo eye field mask at 1 ss.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Video4_360-rotation_rx3-GFP_eyefieldmask_1ss.mp4" target="_blank">Please right click here to download the video (save link as).</a>
Video 5: 360&#176; rotation of a Tg(rx3:GFP) zebrafish embryo at 24 hpf.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Video5_360-rotation_rx3-GFP_24hpf.mp4" target="_blank">Please right click here to download the video (save link as).</a>
Video 6: 360&#176; rotation of a Tg(rx3:GFP) zebrafish embryo eye field mask at 24 hpf.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Video6_360-rotation_rx3-GFP_eyefieldmask_24hpf.mp4" target="_blank">Please right click here to download the video (save link as).</a>
Table 1: arivis Vision4D pipeline for volume analysis of the developing eye field.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Table1 revised.xlsx" target="_blank">Please click here to download this Table.</a>
Table 2: Volume and surface area of the developing eye field acquired by arivis Vision4D. The rows pertaining to the presumptive hypothalamus are highlighted in yellow to distinguish them from the optic vesicles.&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62296/Table2(2).xlsx" target="_blank">Please click here to download this&#160;Table.</a>

Watch this Scientific Journal Video about Visualizing Ocular Morphogenesis by Lightsheet Microscopy Using rx3:GFP Transgenic Zebrafish at JoVE.com

Visualizing Ocular Morphogenesis by Lightsheet Microscopy Using rx3:GFP Transgenic Zebrafish

Here, a&#160;protocol is&#160;provided&#160;for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software.&#160;

Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, ...

visualizing-ocular-morphogenesis-lightsheet-microscopy-using-rx3gfp

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

2.7K Views.  University of Kentucky. Here, a protocol is provided for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software. 

Video: Visualizing Ocular Morphogenesis by Lightsheet Microscopy Using rx3:GFP Transgenic Zebrafish

Deriving Retinal Pigment Epithelium (RPE) from Induced Pluripotent Stem (iPS) Cells by Different Sizes of Embryoid Bodies

Pluripotent stem cells possess the ability to proliferate indefinitely and to differentiate into almost any cell type. Additionally, the development of techniques to reprogram somatic cells into induced pluripotent stem (iPS) cells has generated interest and excitement towards the possibility of customized personal regenerative medicine. However, the efficiency of stem cell differentiation towards a desired lineage remains low. The purpose of this study is to describe a protocol to derive retinal pigment epithelium (RPE) from iPS cells (iPS-RPE) by applying a tissue engineering approach to generate homogenous populations of embryoid bodies (EBs), a common intermediate during in vitro differentiation. The protocol applies the formation of specific size of EBs using microwell plate technology. The methods for identifying protein and gene markers of RPE by immunocytochemistry and reverse-transcription polymerase chain reaction (RT-PCR) are also explained. Finally, the efficiency of differentiation in different sizes of EBs monitored by fluorescence-activated cell sorting (FACS) analysis of RPE markers is described. These techniques will facilitate the differentiation of iPS cells into RPE for future applications.

Deriving Retinal Pigment Epithelium RPE from Induced Pluripotent Stem iPS Cells by Different Sizes of Embryoid Bodies

The objective of this report is to describe the protocols to derive the retinal pigment epithelium (RPE) from induced pluripotent stem (iPS) cells using different sizes of embryoid bodies.

Pluripotent stem cells possess the ability to proliferate indefinitely and to differentiate into almost any cell type. Additionally, the development of ...

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

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 ...

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 ...

Visualizing the Interrenal Steroidogenic Tissue and Its Vascular Microenvironment in Zebrafish

This protocol introduces how to detect differentiated interrenal steroidogenic cells through a simple whole-mount enzymatic activity assay. Identifying differentiated steroidogenic tissues through chromogenic histochemical staining of 3-&#946;-Hydroxysteroid dehydrogenase /&#916;5-4 isomerase (3&#946;-Hsd) activity-positive cells is critical for monitoring the morphology and differentiation of adrenocortical and interrenal tissues in mammals and teleosts, respectively. In the zebrafish model, the optical transparency and tissue permeability of the developing embryos and larvae allow for whole-mount staining of 3&#946;-Hsd activity. This staining protocol, as performed on transgenic fluorescent reporter lines marking the developing pronephric and endothelial cells, enables the detection of the steroidogenic interrenal tissue in addition to the kidney and neighboring vasculature. In combination with vibratome sectioning, immunohistochemistry, and confocal microscopy, we can visualize and assay the vascular microenvironment of interrenal steroidogenic tissues. The 3&#946;-Hsd activity assay is essential for studying the cell biology of the zebrafish interrenal gland because to date, no suitable antibody is available for labeling zebrafish steroidogenic cells. Furthermore, this assay is rapid and simple, thus providing a powerful tool for mutant screens targeting adrenal (interrenal) genetic disorders as well as for determining disruption effects of chemicals on steroidogenesis in pharmaceutical or toxicological studies.

The interrenal gland in zebrafish is the teleostean counterpart of the mammalian adrenal gland. This protocol introduces how to perform the 3-&#946;-hydroxysteroid dehydrogenase (&#916;5-4 isomerase; 3&#946;-Hsd) enzymatic activity assay, which detects differentiated steroidogenic cells in the developing zebrafish.

This protocol introduces how to detect differentiated interrenal steroidogenic cells through a simple whole-mount enzymatic activity assay. Identifying ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Generation of Genetically Modified Mice through the Microinjection of Oocytes

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear injection of DNA expression constructs into fertilized oocytes remains the most commonly used technique to generate transgenic mice for overexpression. With the introduction of CRISPR technology for gene targeting, pronuclear injection into fertilized oocytes has been extended to the generation of both knockout and knockin mice. This work describes the preparation of DNA for injection and the generation of CRISPR guides for gene targeting, with a particular emphasis on quality control. The genotyping procedures required for the identification of potential founders are critical. Innovative genotyping strategies that take advantage of the "multiplexing" capabilities of CRISPR are presented herein. Surgical procedures are also outlined. Together, the steps of the protocol will allow for the generation of genetically modified mice and for the subsequent establishment of mouse colonies for a plethora of research fields, including immunology, neuroscience, cancer, physiology, development, and others.

The microinjection of mouse oocytes is commonly used for both classic transgenesis (i.e., the random integration of transgenes) and CRISPR-mediated gene targeting. This protocol reviews the latest developments in microinjection, with a particular emphasis on quality control and genotyping strategies.

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear ...

High-resolution Episcopic Microscopy (HREM) - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging organic materials with volumes up to 5 x 5 x 7 mm3 in typical numeric resolutions between 1 x 1 x 1 and 5 x 5 x 5 &#181;m3. Specimens are embedded in methacrylate resin and sectioned on a microtome. After each section an image of the block surface is captured with a digital video camera that sits on the phototube connected to the compound microscope head. The optical axis passes through a green fluorescent protein (GFP) filter cube and is aligned with a position, at which the bock holder arm comes to rest after each section. In this way, a series of inherently aligned digital images, displaying subsequent block surfaces are produced. Loading such an image series in three-dimensional (3D) visualization software facilitates the immediate conversion to digital volume data, which permit virtual sectioning in various orthogonal and oblique planes and the creation of volume and surface rendered computer models. We present three simple, tissue specific protocols for processing various groups of organic specimens, including mouse, chick, quail, frog and zebra fish embryos, human biopsy material, uncoated paper and skin replacement material.

High-resolution Episcopic Microscopy HREM - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple and robust protocols for processing biopsy material of various species, embryos of biomedical model organisms and samples of other organic tissues in order to permit digital volume data generation with the high-resolution episcopic microscopy method.

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging ...

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...