This work was supported by Canadian Institutes of Health Research (CIHR), Natural Sciences and Engineering Research Council (NSERC), Alberta Innovates Technology Futures, and Women and Children&#39;s Health Research Institute (WCHRI).

All methods described here have been approved by the University of Alberta Animal Care and Use Committee.
1. Protocol 1: Visualization of SOS using stereomicroscopy and differential interference contrast (DIC) imaging
<ol>
	<li>Embryo collection
	<ol>
		<li>In a tank of dechlorinated water, prepare crosses of gdf6a+/- zebrafish in the evening by pairing a male zebrafish with a female zebrafish. Be sure to separate the male from the female by using a divider to ensure that the embryos are born within a small range of time.</li>
		<li>The following morning, pull the divider and allow the zebrafish to breed for no longer than 30 min. Collect the embryos in Petri dishes with E3 media, described in The Zebrafish Book12, and place them in a 28.5 &#176;C incubator.</li>
		<li>Remove any unfertilized eggs or dead embryos, which will appear white and opaque.</li>
	</ol>
	</li>
	<li>Preparation and live-imaging of zebrafish embryos
	<ol>
		<li>At 20 hpf, replace the E3 media with E3 media containing 0.004% 1-phenyl 2-thiourea (PTU) to prevent pigment production. 
		NOTE: Addition of PTU at a slightly later timepoint, such as 22-24 hpf, is unlikely to interfere with the experiment due to the early age of the embryos at the time of imaging. However, it is recommended to treat the embryos early to completely prevent pigmentation as there is a band of pigmentation that appears in the dorsal eye, which can interfere with the imaging of the SOS.</li>
		<li>Ensure that all embryos are at the correct developmental stages at various points leading up to the time of observation. It is recommended that this is done at the stages at which somite number is clearly visible as outlined by Kimmel et al.13. Remove those that are developmentally immature.</li>
		<li>Place the embryos under a dissecting microscope, and dechorionate the embryos by gently pulling apart the chorion using fine forceps. Visualize the SOS in the dorsal eye. The SOS may appear as an indentation at the dorsal margin of the eye, and a line should be visible across the dorsal eye. For normal SOS closure, observe the embryos at around 20-23 hpf. For examination of delayed SOS closure phenotypes, observe the embryos at 28 hpf or later.</li>
		<li>Sort the embryos that show SOS closure delay from those that do not.</li>
		<li>To photograph these embryos using a dissecting microscope, prepare a Petri dish containing 1% agarose in E3. Lightly prick the center of the agarose to create a shallow hole in which the yolk of the embryo can sit when the embryo is placed on the agarose. This will ensure that the embryo is not at an oblique angle when being photographed.</li>
		<li>Anesthetize embryos with 0.003% tricaine in E3 and place laterally on the agarose.</li>
		<li>To image the embryos using a compound or confocal microscope, transfer the embryo into 35 mm Petri dish containing a small bolus of non-gelled 1% low-melting point agarose in E3 (w/v). Quickly position the embryo laterally using a fine fishing line or an eyelash and wait for the agarose to cool. Once the agarose is firm, pour enough E3 into the dish to cover the agarose. For more details, see Distel and K&#246;ster14. 
		NOTE: If using an inverted microscope, the embryo can be placed against the glass of a glass-coverslip-bottom dish and imaged with a standard 20X objective lens.</li>
		<li>Use a water immersion 20x objective lens to visualize the SOS with a compound microscope. Following visualization, gently pull the agarose from the embryos and fix in 4% paraformaldehyde (PFA) or allow to continue their development.</li>
	</ol>
	</li>
</ol>
2. Protocol 2: Whole-mount immunofluorescent staining of laminin
<ol>
	<li>Whole-mount immunofluorescent staining of laminin: Day 1
	<ol>
		<li>Dechorionate embryos as described in Step 1.2.3, if not already done. Fix embryos in a microcentrifuge tube at the desired timepoint in freshly made 4% PFA for 2 h on a room temperature (22-25 &#176;C) shaker. Wash in 1x PBST for 5 min, four times. 
		NOTE: Following gastrulation, embryos may fix better after dechorionation.</li>
		<li>Permeabilize embryos in 10 &#181;g/mL proteinase K at room temperature for 5 min. Incubation time will depend on the developmental stage at which the embryos are fixed (see Thisse and Thisse15).</li>
		<li>Wash in 1x PBST for 5 min, four times.</li>
		<li>Block embryos in a solution of 5% goat serum and 2 mg/mL bovine serum albumin (BSA) in 1xPBST for 1-2 h on a room temperature shaker.</li>
		<li>Prepare primary antibody solution by diluting rabbit anti-laminin antibody in block solution at a 1:200 dilution.</li>
		<li>Incubate the embryos in anti-laminin primary antibody overnight on a 4 &#176;C shaker.</li>
	</ol>
	</li>
	<li>Whole-mount immunofluorescent staining of laminin: Day 2
	<ol>
		<li>Wash in 1x PBST for 15 min, five times.</li>
		<li>Prepare secondary antibody solution by diluting goat anti-rabbit Alexa Fluor 488 antibody in 1x PBST to a dilution of 1:1000. 
		NOTE: It is possible to adapt this step to suit the resources available to the experimenter by using a different secondary antibody.</li>
		<li>Incubate the embryos in secondary antibody overnight on a 4 &#176;C shaker. Shield from light as much as possible from this step onwards.</li>
		<li>Wash in 1x PBST for 15 min, four times. The embryos can be stored at 4 &#176;C for up to a week, if necessary.</li>
	</ol>
	</li>
	<li>Dissection and mounting of embryonic eyes
	<ol>
		<li>If desired, place the embryos in a small Petri dish and deyolk the embryos in 1x PBST. Do this by gently disrupting the yolk with fine forceps and removing the yolk cells through mild scraping of the yolk sac.</li>
		<li>Prepare the following concentrations of PBS-glycerol series solutions in microcentrifuge tubes: 30%, 50%, and 70% glycerol in PBS. Transfer embryos into 30% glycerol/PBS, making sure to place the embryos on top of the solution and transferring as little of the previous solution as possible. Wait for the embryos to sink to the bottom of the tube.</li>
		<li>When embryos have sunk to the bottom, transfer them to 50% glycerol/PBS. Repeat and transfer to 70% glycerol/PBS.</li>
		<li>Once the embryos have sunk in 70% glycerol/PBS, move them to a small plastic dish for dissections.</li>
		<li>Sever the embryo posterior to the hindbrain, and use the posterior tissue for genotyping, if necessary.</li>
		<li>Move the head to a glass slide, transferring as little glycerol as possible. Use forceps or other fine dissection tools to hold onto the posterior end to keep the head stationary. Use a fine minutien pin or other fine dissection tools to gently insert into the forebrain ventricle from the anterior and push downward to separate the right and left halves of the head from each other. Repeat this while moving posteriorly through the midbrain and into the hindbrain ventricle, essentially fileting the head down the midline. This minimizes manual manipulation of the eye and surrounding tissue, thereby leaving the SOS undamaged.</li>
		<li>Mount each side of the head midline down, eye up. Position four posts of vacuum grease at the corners (an appropriate distance apart for the coverslip being used) and cover with a glass coverslip, pushing down sequentially on each post until the coverslip makes contact with the samples. Pipette 70% glycerol at the edge of the coverslip so that the glycerol is pulled underneath, filling the space between the coverslip and the slide.</li>
		<li>Image samples within a day, or seal around the coverslip with nail polish and image samples only after the nail polish has dried. Store in the dark at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
3. Protocol 3: Visualization of SOS using eGFP-CAAX mRNA
<ol>
	<li>Synthesis of eGFP-CAAX mRNA
	<ol>
		<li>Linearize 1 &#181;g of pCS2-eGFP-CAAX plasmid16 with NotI in a reaction volume of 40 &#181;L for 4 hours at 37 &#176;C.</li>
		<li>To stop the restriction digest reaction, add 10 &#181;L RNase-free water, 2.5 &#181;L 10% SDS and 2.0 &#181;L 10 mg/mL Proteinase K.</li>
		<li>Incubate 1 hour at 50 &#176;C.</li>
		<li>Add the following to the reaction (total volume 200 &#181;L) and proceed to next step: 50 &#181;L RNase-free water, 20 &#181;L 3 M sodium acetate pH 5.2 and 75.5 &#181;L RNase-free water 
		NOTE: RNase-free water is added in two separate occasions to prevent excessive dilution of the sodium acetate.</li>
	</ol>
	</li>
	<li>Purification of DNA through phenol/chloroform extraction and ethanol precipitation
	<ol>
		<li>Add 200 &#181;L phenol:chloroform:isoamyl alcohol and vortex for 20 s. Separate the aqueous and organic phases through centrifugation at 18,000 x g for 5 min.</li>
		<li>Transfer the upper aqueous layer to a new microcentrifuge tube, making sure to avoid the transfer of the bottom organic layer. Add an equal volume of chloroform to the new tube. 
		NOTE: Addition of chloroform is optional, but it is recommended to ensure complete removal of phenol from the sample.</li>
		<li>Vortex for 20 s. Separate the aqueous and organic phases through centrifugation at 18,000 x g for 5 min.</li>
		<li>As before, transfer the upper aqueous layer to a new microcentrifuge tube, making sure to avoid the transfer of the bottom organic layer.</li>
		<li>Add 1/10 volume of 3 M sodium acetate pH 5.2.</li>
		<li>Precipitate DNA by adding 3 volumes of 100% RNase-free ethanol and chill at -20 &#176;C for 15 min. Centrifuge at 18,000 x g for 20 min at 4 &#176;C. A pellet should be visible. Decant the supernatant.</li>
		<li>Wash the pellet with 100 &#181;L of cold 70% ethanol/RNase-free water. After gently mixing to break the pellet loose, centrifuge at 18,000 x g for 15 min at 4 &#176;C. A pellet should be visible. Decant the supernatant.</li>
		<li>Air-dry the pellet for 5 min and resuspend the DNA in 7 &#181;L water. 
		NOTE: The pellet may need to be dried for longer than 5 min depending on the airflow available.</li>
	</ol>
	</li>
	<li>Transcription and purification of eGFP-CAAX mRNA
	<ol>
		<li>In an RNase-free manner, prepare an in vitro transcription reaction with a commercially available Sp6 RNA polymerase kit, using about 1 &#181;g of purified linearized plasmid DNA obtained in Step 3.2. Incubate for 2 h at 37 &#176;C. 
		NOTE: Sp6 RNA polymerase must be used for the production of capped mRNA.</li>
		<li>Add 1 &#181;L of DNase (2 U/&#181;L; RNase free) and incubate for 30 min at 37 &#176;C.</li>
		<li>Purify the mRNA with any commercially available RNA purification kit. Aliquot the mRNA to avoid repeated freeze-thaw, and store at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Injection and visualization
	<ol>
		<li>Obtain embryos as outlined in Protocol 1.1.</li>
		<li>Using a microinjection apparatus, inject 300 pg of eGFP-CAAX mRNA at the 1-cell stage.</li>
		<li>Screen for embryos with the bright expression of eGFP in the eyes using a fluorescence stereoscope.</li>
		<li>Image the embryos as described in Protocol 1.2.</li>
		<li>Alternatively, dechorionate and fix the embryos at the desired timepoint in 4% PFA for 4 hours at room temperature or overnight at 4&#176;C. Wash the embryos in 1x PBST for 5 min, four times, and dechorionate, if not previously done. Dissect the eyes and mount them on slides as described in Protocol 2.3.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Chow, R. L., Lang, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+eye+development+in+vertebrates.">Early eye development in vertebrates.</a> Annual Review of Cell and Developmental Biology. 17, (2001).</li><li>Slavotinek, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eye+development+genes+and+known+syndromes.">Eye development genes and known syndromes.</a> Molecular Genetics and Metabolism. 104 (448-456), (2011).</li><li>Gregory-Evans, C. Y., Williams, M. J., Halford, S., Gregory-Evans, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ocular+coloboma:+a+reassessment+in+the+age+of+molecular+neuroscience.">Ocular coloboma: a reassessment in the age of molecular neuroscience.</a> Journal of Medical Genetics. 41 (12), (2004).</li><li>Onwochei, B. C., Simon, J. W., Bateman, J. B., Couture, K. C., Mir, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ocular+colobomata.">Ocular colobomata.</a> Survey of Ophthalmolgy. 45, 175-194 (2000).</li><li>Williamson, K. A., FitzPatrick, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+genetic+architecture+of+microphthalmia,+anophthalmia+and+coloboma.">The genetic architecture of microphthalmia, anophthalmia and coloboma.</a> European Journal of Medical Genetics. 57, 369-380 (2014).</li><li>Chang, L., Blain, D., Bertuzzi, S., Brooks, B. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uveal+coloboma:+clinical+and+basic+science+update.">Uveal coloboma: clinical and basic science update.</a> Current Opinion in Ophthalmology. 17, 447-470 (2006).</li><li>Kaufman, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+origins+of+Zebrafish+ocular+vasculature.">Development and origins of Zebrafish ocular vasculature.</a> BMC Developmental Biology. 15 (18), (2015).</li><li>Hocking, J. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphogenetic+defects+underlie+Superior+Coloboma,+a+newly+identified+closure+disorder+of+the+dorsal+eye.">Morphogenetic defects underlie Superior Coloboma, a newly identified closure disorder of the dorsal eye.</a> PLOS Genetics. 14 (3), (2018).</li><li>Lawson, N. D., Wolfe, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Forward+and+reverse+genetic+approaches+for+the+analysis+of+vertebrate+development+in+the+zebrafish.">Forward and reverse genetic approaches for the analysis of vertebrate development in the zebrafish.</a> Developmental Cell. 21 (1), (2011).</li><li>Howe, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+zebrafish+reference+genome+sequence+and+its+relationship+to+the+human+genome.">The zebrafish reference genome sequence and its relationship to the human genome.</a> Nature. 496 (7446), (2013).</li><li>Bilotta, J., Saszik, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+zebrafish+as+a+model+visual+system.">The zebrafish as a model visual system.</a> International Journal of Developmental Neuroscience. 19, 621-629 (2001).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book; A guide for the laboratory use of zebrafish (Danio rerio). , (2007).</li><li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+embryonic+development+of+the+zebrafish.">Stages of embryonic development of the zebrafish.</a> Developmental Dynamics. 203, 253-310 (1995).</li><li>Distel, M., K&#246;ster, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+time-lapse+imaging+of+zebrafish+embryonic+development.">In vivo time-lapse imaging of zebrafish embryonic development.</a> Cold Spring Harbor Protocols. , (2007).</li><li>Thisse, C., Thisse, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+in+situ+hybridization+to+whole-mount+zebrafish+embryos.">High-resolution in situ hybridization to whole-mount zebrafish embryos.</a> Nature Protocols. 3, 59-69 (2008).</li><li>Kwan, K. M., Otsuna, H., Kidokoro, H., Carney, K. R., Saijoh, Y., Chien, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+complex+choreography+of+cell+movements+shapes+the+vertebrate+eye.">A complex choreography of cell movements shapes the vertebrate eye.</a> Development. 139, 359-372 (2012).</li><li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullman, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+Embryonic+Development+of+the+Zebrafish.">Stages of Embryonic Development of the Zebrafish.</a> Developmental Dynamics. 203, 253-310 (1995).</li><li>Gfrerer, L., Dougherty, M., Liao, E. 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The authors have no conflicting interests to declare.

Here, we present a standardized series of protocols to observe the SOS in the developing zebrafish embryo. To determine closure delay phenotypes, our protocols have focused on the ability to distinguish the separation of two discrete lobes of the dorsal-nasal and dorsal-temporal sides of the eye, similar to techniques used to visualize choroid fissure closure delay phenotypes in the ventral eye.
These visualization techniques can be used in conjunction with a variety of genetic manipulation te...

The formation of the vertebrate eye is a highly conserved process in which carefully orchestrated intercellular signaling pathways establish tissue types and specify regional identity1. Perturbations to early eye morphogenesis result in profound defects to the architecture of the eye and are frequently blinding2. One such disease results from the failure to close the choroid ocular fissure in the ventral side of the optic cup3. This disorder, known as ocular coloboma, is estimated to occur in 1 out of 4-5000 live births and cause 3-11% of pediatric blindness, commonly manifesting as a keyhole-like structure that protrudes inferiorly from the pupil in the center of the eye4,5,6. The function of the choroid fissure is to provide an entry point for early vasculature growing into the optic cup, after which the sides of the fissure will fuse to enclose the vessels7.
While ocular coloboma has been known since ancient times, we have recently identified a novel subset of coloboma patients with tissue loss affecting the superior/dorsal aspect of the eye. Recent work in our lab has led to the discovery of an ocular structure in the zebrafish dorsal eye, which we refer to as the superior ocular sulcus (SOS) or superior fissure8. It is important to note that the structure has characteristics of both a sulcus and a fissure. Similar to a sulcus, it is a continual tissue layer that spans from the nasal to the temporal retina. In addition, the closure of the structure is not mediated by a fusion of the two opposing basement membrane, and it appears to require a morphogenetic process by which the structure is populated by cells. However, similar to a fissure, it forms a structure that separates the nasal and temporal sides of the dorsal eye with the basement membrane. For consistency, we will refer to it as SOS in this text.
The SOS is evolutionarily conserved across vertebrates, being visible during eye morphogenesis in fish, chick, newt, and mouse8. In contrast to the choroid fissure, which is present from 20-60 hours post-fertilization (hpf) in zebrafish, the SOS is highly transient, being easily visible from 20-23 hpf and absent by 26 hpf8. Recent research in our lab has found that, similar to the choroid fissure, the SOS plays a role in vascular guidance during eye morphogenesis8. Although the factors that control the formation and closure of the SOS are not yet fully understood, our data did highlight roles for dorsal-ventral eye patterning genes8.
Zebrafish is an excellent model organism to study the SOS. As a model system, it provides a number of advantages in studying eye development: it is a vertebrate model; each generation exhibits high fecundity (~200 embryos); its genome has been fully sequenced, which facilitates genetic manipulation; and approximately 70% of human genes have at least one zebrafish orthologue, making it an ideal genetics-based model of human disease9,10. Most importantly, its development takes place externally to the mother, and its larvae are transparent, which allows for the visualization of the developing eye with relative ease11.
In this set of protocols, we describe the techniques through which the SOS can be visualized in zebrafish larvae. The variety of visualization techniques used in this report will allow clear observation of the SOS during normal eye development, as well as the ability to detect SOS closure defects. Our example protocols will feature investigations of Gdf6, a BMP localized to the dorsal eye and known regulator of SOS closure. Further, these techniques can be combined with experimental manipulations to identify genetic factors or pharmacological agents that affect proper SOS formation and closure. In addition, we have included a protocol through which the fluorescent imaging of all cell membranes is possible, allowing the experimenter to observe morphological changes to the cells surrounding the SOS. Our goal is to establish a set of standardized protocols that can be used throughout the scientific community to offer new insights into this novel structure of the developing eye.

<table>
	<tbody>
		<tr>
			<td>1-phenyl 2-thiourea</td>
			<td>Sigma Aldrich</td>
			<td>P7629-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>100 mm Petri dish</td>
			<td>Fisher Scientific</td>
			<td>FB0875713</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Petri dish</td>
			<td>Corning</td>
			<td>CLS430588</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>BioShop Canada Inc.</td>
			<td>AGA001.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td>Sigma Aldrich</td>
			<td>A7906-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>DIC/Fluorescence microscope</td>
			<td>Zeiss</td>
			<td>AxioImager Z1</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection microscope</td>
			<td>Olympus</td>
			<td>SZX12</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection microscope camera</td>
			<td>Qimaging</td>
			<td>MicroPublisher 5.0 RTV</td>
			<td></td>
		</tr>
		<tr>
			<td>Dow Corning High-vacuum grease</td>
			<td>Fisher Scientific</td>
			<td>14-635-5D</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine)</td>
			<td>Sigma Aldrich</td>
			<td>A5040-25G</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-rabbit Alexa Fluor 488</td>
			<td>Abcam</td>
			<td>ab150077</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat serum</td>
			<td>Sigma Aldrich</td>
			<td>G9023</td>
			<td></td>
		</tr>
		<tr>
			<td>Image capture software</td>
			<td>Zeiss</td>
			<td>ZEN</td>
			<td></td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>VWR</td>
			<td>Model 1545</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Cover Glass (22 mm x 22 mm)</td>
			<td>Fisher Scientific</td>
			<td>12-542B</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slide</td>
			<td>Fisher Scientific</td>
			<td>12-544-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Minutien pin</td>
			<td>Fine Science Tools</td>
			<td>26002-10</td>
			<td></td>
		</tr>
		<tr>
			<td>mMessage mMachine Sp6 Transcription Kit</td>
			<td>Invitrogen</td>
			<td>AM1340</td>
			<td></td>
		</tr>
		<tr>
			<td>NotI</td>
			<td>New England Biolabs</td>
			<td>R0189S</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde (PFA)</td>
			<td>Sigma Aldrich</td>
			<td>P6148-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenol:Chloroform:Isoamyl Alcohol pH 6.7 +/- 0.2</td>
			<td>Fisher Scientific</td>
			<td>BP1752-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Proteinase K</td>
			<td>Sigma Aldrich</td>
			<td>P4850</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti-laminin antibody</td>
			<td>Millipore Sigma</td>
			<td>L9393</td>
			<td></td>
		</tr>
		<tr>
			<td>TURBO Dnase (2 U/&#181;L)</td>
			<td>Invitrogen</td>
			<td>AM2238</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrapure low-melting point agarose</td>
			<td>Invitrogen</td>
			<td>16520-100</td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure Sodium Dodecyl Sulfate (SDS)</td>
			<td>Invitrogen</td>
			<td>15525017</td>
			<td></td>
		</tr>
	</tbody>
</table>

visualization of the superior ocular sulcus during danio rerio embryogenesis

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid fissure closure. Recently, the identification of individuals with coloboma in the superior aspect of the iris, retina, and lens led to the discovery of a novel structure, referred to as the superior fissure or superior ocular sulcus (SOS), that is transiently present on the dorsal aspect of the optic cup during vertebrate eye development. Although this structure is conserved across mice, chick, fish, and newt, our current understanding of the SOS is limited. In order to elucidate factors that contribute to its formation and closure, it is imperative to be able to observe it and identify abnormalities, such as delay in the closure of the SOS. Here, we set out to create a standardized series of protocols that can be used to efficiently visualize the SOS by combining widely available microscopy techniques with common molecular biology techniques such as immunofluorescent staining and mRNA overexpression. While this set of protocols focuses on the ability to observe SOS closure delay, it is adaptable to the experimenter&#39;s needs and can be easily modified. Overall, we hope to create an approachable method through which our understanding of the SOS can be advanced to expand the current knowledge of vertebrate eye development.

The zebrafish SOS appears at 20 hpf in the presumptive dorsal retina8. By 23 hpf the SOS transitions from its initial narrow architecture to a wide indentation and by 26 hpf it is no longer visible8. Therefore, to examine the SOS during normal zebrafish eye development, the embryos must be observed between 20-23 hpf. During this period, the SOS is observable through the dissecting microscope and via DIC imaging as a thin line in the dorsal e...

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Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Here, we present a standardized series of protocols to observe the superior ocular sulcus, a recently-identified, evolutionarily-conserved structure in the vertebrate eye. Using zebrafish larvae, we demonstrate techniques necessary to identify factors that contribute to the formation and closure of the superior ocular sulcus.

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid ...

visualization-superior-ocular-sulcus-during-danio-rerio

Research

JoVE Journal

Developmental Biology

5.3K Views.  University of Alberta. Here, we present a standardized series of protocols to observe the superior ocular sulcus, a recently-identified, evolutionarily-conserved structure in the vertebrate eye. Using zebrafish larvae, we demonstrate techniques necessary to identify factors that contribute to the formation and closure of the superior ocular sulcus.

Video: Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

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 ...

Assessment of Maternal Vascular Remodeling During Pregnancy in the Mouse Uterus

The placenta mediates the exchange of factors such as gases and nutrients between mother and fetus and has specific demands for supply of blood from the maternal circulation. The maternal uterine vasculature needs to adapt to this temporary demand and the success of this arterial remodeling process has implications for fetal growth. Cells of the maternal immune system, especially natural killer (NK) cells, play a critical role in this process. Here we describe a method to assess the degree of remodeling of maternal spiral arteries during mouse pregnancy. Hematoxylin and eosin-stained tissue sections are scanned and the size of the vessels analysed. As a complementary validation method, we also present a qualitative assessment for the success of the remodeling process by immunohistochemical detection of smooth muscle actin (SMA), which normally disappears from within the arterial vascular media at mid-gestation. Together, these methods enable determination of an important parameter of the pregnancy phenotype. These results can be combined with other endpoints of mouse pregnancy to provide insight into the mechanisms underlying pregnancy-related complications.

This protocol describes a technique to assess changes in the maternal vasculature during pregnancy in mice. Using stereological methods, remodeling of the decidual spiral arteries is assessed quantitatively and the results confirmed qualitatively using immunohistochemistry.

The placenta mediates the exchange of factors such as gases and nutrients between mother and fetus and has specific demands for supply of blood from the ...

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth factors, and extracellular matrix. The ability of HSCs to remain quiescent, self-renew or differentiate, and acquire mutations and become malignant depends upon the complex interactions they establish with different stromal components. To observe the crosstalk between human HSCs and the human BM niche in physiological and pathological conditions, we designed a protocol to ectopically model and image a humanized BM niche in immunodeficient mice. We show that the use of different cellular components allows for the formation of humanized structures and the opportunity to sustain long-term human hematopoietic engraftment. Using two-photon microscopy, we can live-image these structures in situ at the single-cell resolution, providing a powerful new tool for the functional characterization of the human BM microenvironment and its role in regulating normal and malignant hematopoiesis.

A method to create and live-image different humanized bone-marrow niches in mice is presented. Based on the supportive niche created by human mesenchymal cells, the addition of human endothelial cells induces the formation of human vessels, while the addition of rhBMP-2 induces the formation of human-mouse chimeric mature bone tissue.

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth ...

A Method for Characterizing Embryogenesis in Arabidopsis

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, early stage plant embryos are small and contain few cells, making them difficult to observe and analyze. A method is described here for characterizing pattern formation in plant embryos under a microscope using the model organism Arabidopsis. Following the clearance of fresh ovules using Hoyer's solution, the cell number in and morphology of embryos could be observed, and their developmental stage could be determined by differential interference contrast microscopy using a 100X oil immersion lens. In addition, the expression of specific marker proteins tagged with Green Fluorescent Protein (GFP) was monitored to annotate cell identity specification during embryo patterning by confocal laser scanning microscopy. Thus, this method can be used to observe pattern formation in wild-type plant embryos at the cellular and molecular levels, and to characterize the role of specific genes in embryo patterning by comparing pattern formation in embryos from wild-type plants and embryo-lethal mutants. Therefore, the method can be used to characterize embryogenesis in Arabidopsis.

A Method for Characterizing Embryogenesis in Arabidopsis

This protocol outlines a method for observing embryogenesis in Arabidopsis via ovule clearance followed by the inspection of embryo pattern formation under a microscope.

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Visualization of Tangential Cell Migration in the Developing Chick Optic Tectum

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture can be recorded under video microscopy. For analyzing cell migration in the developing brain, slice culture is commonly used to observe cell migration parallel to the slice section, such as radial cell migration. However, limited information can be obtained from the slice culture method to analyze cell migration perpendicular to the slice section, such as tangential cell migration. Here, we present the protocols for time-lapse imaging to visualize tangential cell migration in the developing chick optic tectum. A combination of cell labeling by electroporation in ovo and a subsequent flat-mount culture on the cell culture insert enables detection of migrating cell movement in the horizontal plane. Moreover, our method facilitates detection of both individual cell behavior and the collective action of a group of cells in the long term. This method can potentially be applied to detect the sequential change of the fluorescent-labeled micro-structure, including the axonal elongation in the neural tissue or cell displacement in the non-neural tissue.

We describe the methods for fluorescent labeling of tangentially migrating cells by electroporation, and for time-lapse imaging of the labeled cell movement in a flat-mount culture in order to visualize migrating cell behavior in the developing chick optic tectum.

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture ...

Analysis of Cardiac Chamber Development During Mouse Embryogenesis Using Whole Mount Epifluorescence

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during heart development using ventricular specific fluorescent reporter knock-in mice (MLC-2v-tdTomato mice). Heart development involves a linear heart tube formation, the heart tube looping, and four chamber septation. These complex processes are highly conserved in all vertebrates. The mouse embryonic heart has been widely used for heart developmental studies. However, due to their extremely small size, dissecting mouse embryonic hearts is technically challenging. In addition, visualization of cardiac chamber formation often needs in situ hybridization, beta-galactosidase staining using LacZ reporter mice, or immunostaining of sectioned embryonic hearts. Here, we describe how to dissect mouse embryonic hearts and directly visualize ventricular chamber formation of MLC-2v-tdTomato mice using whole mount epifluorescent microscopy. With this method, it is possible to directly examine heart tube formation and looping, and four chamber formation without further experimental manipulation of mouse embryos. Although the MLC-2v-tdTomato reporter knock-in mouse line is used in this protocol as an example, this protocol can be applied to other heart-specific fluorescent reporter transgenic mouse lines.

We present the protocols to examine mouse heart development using whole mount epifluorescent microscopy on mouse embryos dissected from ventricular specific MLC-2v-tdTomato reporter knock-in mice. This method allows us to directly visualize each stage of the ventricular formation during mouse heart development without labor-intensive histochemical methods.

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during ...

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C (UV-C) Damage

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational health hazard. Here, we demonstrate the exposure of primary stem cells from the mouse ocular surface to UV-C radiation. Reactive oxygen species (ROS) formation is the readout of the extent of oxidative stress/damage. In an experimental in vitro setting, it is also essential to assess the percentage of dead cells generated due to oxidative stress. In this article, we will demonstrate the 2&#39;,7&#39;-Dichlorofluoresceindiacetate (DCFDA) staining of UV-C exposed mouse primary ocular surface stem cells and their quantification based on the fluorescent images of DCFDA staining. DCFDA staining directly corresponds to ROS generation. We also demonstrate the quantification of dead and live cells by simultaneous staining with propidium iodide (PI) and Hoechst 3332 respectively and the percentage of DCFDA (ROS positive) and PI positive cells.

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C UV-C Damage

This protocol demonstrates the simultaneous detection of reactive oxygen species (ROS), live cells, and dead cells in live primary cultures from mouse ocular surface cells. 2&#39;,7&#39;-Dichlorofluoresceindiacetate, propidium iodide, and Hoechst staining are used to assess the ROS, dead cells, and live cells, respectively, followed by imaging and analysis.

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

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Chapters in this video

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

Assessment of Maternal Vascular Remodeling During Pregnancy in the Mouse Uterus

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

A Method for Characterizing Embryogenesis in Arabidopsis

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Visualization of Tangential Cell Migration in the Developing Chick Optic Tectum

Analysis of Cardiac Chamber Development During Mouse Embryogenesis Using Whole Mount Epifluorescence

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C (UV-C) Damage

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages