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>

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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. 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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/&amp;micro;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. Deze set standaardiseert protocollen kan worden gebruikt om nieuw inzicht te bieden in de superieure oculaire sulcus, of SOS, een onlangs ontdekte structuur van het ontwikkelende oog. Deze protocollen zullen elke experimentator over de hele wereld in staat stellen om de SOS duidelijk te visualiseren tijdens de ontwikkeling van zebravissenoogontwikkeling. Begin met het plaatsen van heterozygootlijke groei differentiatie factor zes Een mannelijke en vrouwelijke zebravis op afzonderlijke zijden ...

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

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Neuroscience

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

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

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

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

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of morphogenetic events, such as gastrulation2, neurulation3-5, somitogenesis6, heart bending7,8, and brain formation9-13, during early embryogenesis. Key to understanding morphogenetic processes is to follow them dynamically by time-lapse imaging. The acquisition of time-lapse movies of chick embryogenesis ex ovo has been limited either to short time windows or to the need for an incubator to control temperature and humidity around the embryo14. Here, we present a new technique to culture chick embryos ex ovo for high-resolution time-lapse imaging using transmitted light microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique (EC-culture)1 and allows for the culturing of chick embryos without the need for a climate chamber. The embryo is sandwiched between two identical filter paper carriers and is kept fully submerged in a simple, temperature-controlled medium covered by a layer of light mineral oil. Starting from the primitive streak stage (Hamburger-Hamilton stage 5, HH5)15 up to at least the 28-somite stage (HH16)15, embryos can be cultured with either their ventral or dorsal side up. This allows the acquisition of time-lapse movies covering about 30 hr of embryonic development. Representative time-lapse frames and movies are shown. Embryos are compared morphologically to an embryo cultured in the standard EC-culture.
The submerged filter paper sandwich provides a stable environment to study early dorsal and ventral morphogenetic processes. It also allows for live fluorescence imaging and micromanipulations, such as microsurgery, bead implantation, microinjection, gene silencing, and electroporation, and has a strong potential to be combined with immersion objectives for laser-based imaging (including light-sheet microscopy).

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Here, we present a new technique to culture chick embryos ex ovo for time-lapse imaging using transmitted light and fluorescence microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique1 that allows for the culturing of chick embryos fully-submerged in a liquid culture medium.

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of ...

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

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

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

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

Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization

The Drosophila optic lobe, comprised of four neuropils: the lamina, medulla, lobula and lobula plate, is an excellent model system for exploring the developmental mechanisms that generate neural diversity and drive circuit assembly. Given its complex three-dimensional organization, analysis of the optic lobe requires that one understand how its adult neuropils and larval progenitors are positioned relative to each other and the central brain. Here, we describe a protocol for the dissection, immunostaining and mounting of larval and adult brains for optic lobe imaging. Special emphasis is placed on the relationship between mounting orientation and the spatial organization of the optic lobe. We describe three mounting strategies in the larva (anterior, posterior and lateral) and three in the adult (anterior, posterior and horizontal), each of which provide an ideal imaging angle for a distinct optic lobe structure.

Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization

This protocol describes three steps to prepare larval and adult Drosophila optic lobes for imaging: 1) brain dissections, 2) immunohistochemistry and 3) mounting. Emphasis is placed on step 3, as distinct mounting orientations are required to visualize specific optic lobe structures.&#160;

The Drosophila optic lobe, comprised of four neuropils: the lamina, medulla, lobula and lobula plate, is an excellent model system for exploring the ...

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.

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, ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Biology

Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis support continuous growth of the entire visual system, both in the eye and the brain. Coordinated growth between the retinae and the optic tectum ensures accurate retinotopic mapping as new neurons are added in the eyes and brain. To address whether retinal axons provide crucial information for regulating tectal stem and progenitor cell behaviors such as survival, proliferation, and/or differentiation, it is necessary to be able to compare innervated and denervated tectal lobes within the same animal and across animals. 
Surgical removal of one eye from living larval zebrafish followed by observation of the optic tectum achieves this goal. The accompanying video demonstrates how to anesthetize larvae, electrolytically sharpen tungsten needles, and use them to remove one eye. It next shows how to dissect brains from fixed zebrafish larvae. Finally, the video provides an overview of the protocol for immunohistochemistry and a demonstration of how to mount stained embryos in low-melting-point agarose for microscopy.

The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis ...

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

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