Name: imaging and 3d reconstruction of cerebrovascular structures in embryonic zebrafish
Uploaded: 2014-04-22T13:45:00
Duration: 8 min
Description: Watch this Scientific Journal Video about Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish at JoVE.com

The authors thank past and present members of our laboratories who helped develop these techniques. Partial funding provided to DE by a grant from the California Institute for Regenerative Medicine/CIRM (RN1-00538).

1. Zebrafish Husbandry, Embryo Generation, and Treatment
<ol>
	<li>Conduct the following zebrafish protocols under the guidance of an institutional animal care and use committee (IACUC) and within animal care guidelines of the NIH or other regulatory bodies/guidelines.</li>
	<li>Zebrafish strains that express fluorescent proteins in specific tissues, cells, or organs are available from the Zebrafish International Resource Center (ZIRC). For example, Tg (kdr:EGPF)s843 express EGFP in vascular endothelial cells6, which can be used to produce complete 3D vascular structures as shown in this protocol. Other lines of transgenic zebrafish are available from ZIRC.</li>
	<li>House adult zebrafish in an appropriate aquaculture system that monitors pH, salinity, temperature, dissolved oxygen, light, and other environmental factors7. The zebrafish shown here were housed in an Aquaneering Inc. system (San Diego, CA) at 28.5 &#176;C with a 14 hour light/10 hour dark cycle. Feed adult zebrafish a balanced diet of brine shrimp and NRD 4/6 Fish food (Brine Shrimp Direct, Ogden, Utah).</li>
	<li>Adult male and female zebrafish of breeding age should be housed separately to increase mating successful.</li>
	<li>Stimulate egg laying and fertilization by placing females (2-4) and males (4-6) together in a mating container that has a mesh bottom with holes large enough for the eggs to fall through, but too small for the zebrafish adults to pass. Set up matings the evening before; eggs are laid near dawn, usually while the daytime light cycle slowly increases in intensity (dawn). Check for eggs at the bottom of the mating container every 15-30 minutes.</li>
	<li>Collect eggs using a mesh strainer and clean with E3 buffer (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4). Transfer the eggs to 100 mm culture plates filled with E3 buffer and store in a 28 &#176;C incubator.</li>
	<li>To study chemicals that alter cerebrovascular branching add desired chemical concentrations. For example neovasular branching can be induced with &#947;-secretase inhibitor (GSI IX/DAPT/N-[N-(3,5-Difluorophenacetyl-L-alanyl)]-S-phenylglycinet-butyl ester) solubilized in DMSO4, beginning 24 hours post fertilization (hpf). At that time-point embryos are still within the chorion, and many chemicals can pass through it8. If a treatment condition has neural or motor effects that may impair the ability of embryos to break free of the chorion, then embryos should be de-chorionated between 24 and 48 hpf, which can be done with either forceps of pronase2. Embryos shown in the images provided were dechorionated by gently grabbing the chorion with two sharpened forceps and tearing it open.</li>
	<li>If needed/desired, pigment formation can be inhibited by adding 0.003% N-Phenylthiourea (PTU) to the E3 buffer at 24 hpf.</li>
</ol>
2. Confocal Imaging of Cerebrovascular Structures in Fixed Zebrafish Embryos
<ol>
	<li>Sacrifice embryos in 250 mg/L Tricaine methanesulfonate, and then fix them by immersion in 2 - 4% paraformaldehyde overnight at 4 &#176;C. Containers with fluorescent embryos should be wrapped in foil. Once fixed, embryos should be stored in PBS at 4 &#176;C until imaged - EGFP fluorescence intensity becomes less resolved after about a week, but morphology (as viewed in bright field) is preserved much longer.</li>
	<li>Prepare to mount the embryo by first removing one eye using a sharpened tungsten needle. Cut around the tissue connecting the eye first then cut the muscles, and finally cut the optic nerve to displace the eye. (Note: If imaging both sides is desired remove both eyes.)</li>
	<li>Once the eye is removed, mount the embryo on a coverglass using a drop of 3% methylcellulose. Orient the embryo so that the side with the removed eye is facing the coverglass and is as close to the glass as possible. Cover the entire embryo with methylcellulose to prevent desiccation during imaging.</li>
	<li>Image the mounted embryo immediately using an inverted confocal microscope equipped with a high-quality 20x Plan Apo objective (numerical aperture = 0.75 or better). This configuration is preferable to a non-inverted microscope that would require sandwiching the embryo between two glass planes, and pressing the embryo against the top glass.</li>
	<li>Collect optical slices in 1 &#181;m increments using a medium or large aperture setting. Larger steps of 2.5 &#181;m can also be used, but it may be more difficult to determine the spatial order of smaller objects. Small apertures produce sharper detail, but the longer scans required may bleach EGFP and also limit the depth of imaging into the embryo that can be achieved. A confocal microscope allows for imaging about half-way through an embryo.</li>
	<li>If imaging of the entire fish is desired, remove both eyes at the outset (see Note in step 3.2), rotate the embryo after imaging one side and repeat steps 2.3 - 2.5 for the opposite side.</li>
</ol>
3. 3D Reconstruction of Embryonic Zebrafish Cerebrovasculature
<ol>
	<li>Use the Fiji distribution3 of open source ImageJ (http://fiji.sc), which is optimized for 3D renderings, free of charge, and compatible with PC, Mac and Linux computers.</li>
	<li>Import confocal stacks to Fiji by going to Plugins&#62;LOCI&#62;BioFormats Importer 9 then select the confocal file, e.g. name.ids for Nikon stacks (Figure 2A). Select View stack with Hyperstack, Color mode: grayscale, check autoscale, check split channels.</li>
	<li>These selections will open four separate channel panels; for EGFP only one will be needed, usually the third one down. Close the other three panels (red, blue and alpha) leaving the 16-bit image with a long name followed by C=1 (Figure 2A).</li>
	<li>Adjust threshold by scanning though the image using the scroll tab along the bottom to find a slice that has the region or structure of interest, then go to Image&#62;Adjust&#62;Threshold (Figure 2C).</li>
	<li>In the panel that comes up, slide the top bar (black level) to the left so that the structure can be seen quite well over background, and leave the bottom slide (white level) where it is (Figure 2D). Select B&#38;W, select Dark background. Do not select calculate Threshold for each image unless different adjustments are required for each slice, select Black background. This process creates a new 8-bit image.</li>
	<li>WARNING: Threshold cannot be undone or saved in ImageJ, so the confocal stack will have to be reloaded every time a change is needed. Write down the numbers and try different settings until the desired result is achieved.</li>
	<li>Go to Plugins&#62;3D Viewer&#62;Threshold 0, Resampling factor 1 (best) or 2 (good), deselect the red and blue color boxes to make a green 3D output - otherwise it will produce a white rendering - select Apply (not Auto) (Figure 2E).</li>
	<li>Rotate, spin and zoom the 3D image using the mouse and keyboard controls (Figure 2F).</li>
	<li>Save still images at any point by using the capture option in the menu. The size of the image box on the screen dictates pixel dimensions of the image produced so if a high resolution image is desired make the box larger by dragging the bottom right corner.</li>
	<li>Create a spinning 3D movie by selecting View&#62;Record 360 deg rotation (Figure 2G). Rotation defaults to 2 degrees per step, but it can be changed to 5 degrees for example, which will create much smaller file sizes, but may add jerkiness to the animation.</li>
	<li>Save the file in one of several available formats for later viewing with media players, uploading to the Internet, or using in PowerPoint presentations. The open source media player, VLC (<a href="http://www.videolan.org/vlc/index.html">http://www.videolan.org/vlc/index.html</a>), is free of charge and handles these videos very well.</li>
</ol>

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Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+and+molecular+analyses+of+vascular+tube+and+lumen+formation+in+zebrafish.">Cellular and molecular analyses of vascular tube and lumen formation in zebrafish.</a> Development. 132, 5199-5209 (2005).</li><li>Lawrence, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+husbandry+of+zebrafish+(Danio+rerio):+a+review.">The husbandry of zebrafish (Danio rerio): a review.</a> Aquaculture. 269, 1-20 (2007).</li><li>Hagedorn, M., Kleinhans, F. 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There is nothing to disclose.

The methods described here provide a foundation for visual studies of the vascular system in developing zebrafish. Live specimens can be used to assess physiological parameters, such as heart rate and heart stroke volume, while fixed samples can be used for high-resolution confocal imaging. Drosophila and C. elegans allow for whole body imaging, but zebrafish are vertebrates and provide a useful model for vertebrate tissues, including an endothelial cell-lined vascular system. These studies can incorporate significant transgenic lines and genomic resources from the zebrafish research community. 3D reconstruction and rendering of confocal images from embryonic zebrafish, as described here, allow for a systems biology approach to vascular branching and blood vessel density that is not possible with larger animal models such as rats and mice. Further, as amniotes zebrafish develop in a modifiable environment (E3 buffer), where one can easily add chemicals that inhibit specific enzymes or other processes that affect vascular development. The concentration and timing of chemical delivery can be altered, enabling the researcher to fine-tune treatment conditions.
1. Modifications and troubleshooting
Modifications to this system can incorporate transgenic lines of zebrafish that express EGFP or other fluorescent proteins in a variety of tissue, organ or region specific patterns10. Further, analysis of neovascular changes in the zebrafish retina has recently been published5. Problems with pigmentation in older embryos and adult zebrafish can be compensated by crossing with transgenic lines that do not produce scale pigments or retinal pigment. Problems with diminished fluorescence typically result from inappropriate fixation conditions. Paraformaldehyde (4%) for 1 day is optimal, but stronger fixatives, such as glutaraldehyde, osmium tetroxide or alcohol, may destroy EGFP fluorescence. After fixation, embryos should be kept in PBS at 4 &#176;C and always protected from direct sunlight.
2. Limitations of this technique
The quality and resolution of 3D renderings produced with this protocol depend on the quality of images generated. Light penetration through these embryos is limited to the mid-sagittal plane using a standard confocal microscope. This aspect of imaging limits the depth of imaging in older embryos and adults, but more advanced multi-photon microscopy systems allow for imaging at greater depths.
3. Significance with respect to existing methods
This protocol provides an approach to the analysis of blood vessel networks on a systems level that can incorporate an entire animal. Earlier representations of such data often relied on series of images laid out together, but 3D rendering provides better resolution of the spatial relationships involved.
4. Future applications
New developments in imaging and tissue processing will provide many new applications for these methods that may include making older embryos or adults transparent11-14.Enhanced transparency will greatly increase tissue penetration by confocal and multi-photon lasers. Further, as the speed of cameras and photomultiplier tubes increase it may soon be possible to produce 3D renderings of fish in real-time, providing a 4th dimension of analysis.
5. Critical steps
A critical step in this protocol is preparation for imaging, which includes proper fixation. Imaging should be done as soon as possible after fixation with high quality objectives that have the best numeric apertures available. Resolution depends on the imaging system used, so high quality systems are generally better. Generating 3D renderings is memory intensive for so newer, high-end computers with a large amount of memory and good graphics processors are recommended.
The visual biology system described here has been optimized for transgenic zebrafish that express EGFP in vascular endothelial cells, though these methods can be adapted to transgenic embryos that express GFP, or other fluorescent proteins, in populations of neurons, muscles, glands or any number of other cells. The major advantage in working with this system is the ability to study what is happening in the entire embryo at any time during this developmental period, in fixed and/or live animals.

Zebrafish provide a powerful system to study developmental biology, and the relative transparency of their embryos is amenable to imaging-based studies1. The zebrafish has now been used as a model for vertebrate development for decades. Teleosts, including zebrafish, have a simplified vertebrate vascular system that has no reasonable homolog in invertebrates. Blood is pumped from the anterior chamber of a two-chambered heart through gills, where it is oxygenated. Blood from the gills converges at the dorsal aorta and passes through arteries that branch into smaller and smaller vessels, eventually reaching capillaries in organ tissues. Within capillaries oxygen is released and carbon dioxide is absorbed. On the venous side of capillaries blood flows into larger and larger veins and is finally drawn into the posterior chamber of the heart, where the cycle repeats.
An adult zebrafish can lay 200 or more eggs at a time, and once fertilized, they develop quickly2. Within one day the body axis is well developed, including muscles that contract and move the embryo around inside the chorionic membrane. From 2-7 days post-fertilization (dpf) most body systems develop, including eyes and a central nervous system that can coordinate swimming toward food or away from bright light. Up to 7 dpf embryos are small enough to allow for visualization with microscopy. Transgenic lines that express fluorescent proteins can be imaged with confocal or fluorescence microscopy. Confocal imaging can be paired with open-source software3 to create 3D renderings of complete vascular structures in zebrafish embryos that provide a systems biology perspective of vascular development. Studies concerned with changes in vascular and cerebrovascular complexity will benefit from this protocol as it allows for a systems level analysis of vascular networks4,5. A compilation of methods and resources are provided to allow for easy adoption and of these techniques for studies that require imaging of vascular structures in embryonic zebrafish. The cost efficiency of zebrafish as an animal model is combining with emerging imaging technologies to provide new platforms with which to assess angiogenic effects of molecular pathways in vertebrate development and homeostasis.

<table border="0" cellpadding="0" cellspacing="0" width="889">
	<tbody>
		<tr>
			<td>N &#8211; Phenylthiourea&#160;</td>
			<td>Alfa Aesar, catalog #41972</td>
			<td></td>
			<td>0.2 M in E3 buffer, kept at 4oC</td>
		</tr>
		<tr>
			<td>E3 buffer</td>
			<td>Sigma</td>
			<td></td>
			<td>5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4</td>
		</tr>
		<tr>
			<td>Confocal microscope&#160;</td>
			<td>Nikon</td>
			<td></td>
			<td>D-EclipseC1 on a Nikon TE-2000U</td>
		</tr>
		<tr>
			<td>Glass bottom dishes</td>
			<td>Mat-Tek</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>GSI IX/DAPT</td>
			<td></td>
			<td></td>
			<td>N-[N-(3,5-Difluorophenacetyl-L-alanyl)]-S-phenylglycinet-butyl ester EMD Biosciences</td>
		</tr>
		<tr>
			<td>24 well plates&#160;</td>
			<td>Becton-Dickinson, cat# 351147</td>
			<td></td>
			<td>BD Falcon</td>
		</tr>
		<tr>
			<td>Transfer pipettes&#160;</td>
			<td>VWR,&#160; cat #414004-001</td>
			<td></td>
			<td>VWR disposable transfer pipets</td>
		</tr>
		<tr>
			<td>Methyl-cellulose&#160;</td>
			<td>Alfa Aesar, cat#43146</td>
			<td></td>
			<td>3% in E3 buffer</td>
		</tr>
		<tr>
			<td>NRD 4/6 Fish food&#160;</td>
			<td>Brine Shrimp Direct</td>
			<td></td>
			<td>Dried</td>
		</tr>
		<tr>
			<td>Brine shrimp</td>
			<td>Brine Shrimp Direct</td>
			<td></td>
			<td>Live</td>
		</tr>
		<tr>
			<td>Tungsten wire&#160;</td>
			<td>Small Parts # TW-016-60</td>
			<td></td>
			<td>0.016&#8221; OD</td>
		</tr>
		<tr>
			<td>Tricaine</td>
			<td>VWR # 101107-950</td>
			<td></td>
			<td>Tricaine methanesulfonate 250 mg/L in E3 buffer</td>
		</tr>
	</tbody>
</table>

imaging and 3d reconstruction of cerebrovascular structures in embryonic zebrafish

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

3D reconstruction of vascular structures provides a comprehensive and visually interesting perspective of zebrafish development. Figures 1 and 2 show methods as they typically done. Figure 3 shows several angles of vascular structures in a 6 dpf zebrafish embryo that expressed EGFP in endothelial cells. With a solid green or white color it can be difficult to appreciate signal intensity; pseudo-coloring provide image intensity from a look-up-table and allows better depth perception when structures overlap. An example of a pseudo-colored 3D image of the vasculature in a 6 dpf zebrafish is provided in Figure 4. Fluorescence imaging of live embryos can be used to study physiological characteristics that include eye and body movement, and cardiac activity. Figures 3 and 4 show representative results obtained with these methods, using the transgenic zebrafish line described. Imaging resolution depends on microscope characteristics, but the brightness of the EGFP signal is sufficient for good image quality with most commercial systems. Reconstruction and rendering of 3D representations is consistent and options within this open-source software provide consistently good results.
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/50417/50417fig1highres.jpg" width="600" /> 
Figure 1. Eye removal. A) A fixed 3 dpf embryo with a tungsten need positioned next to the eye. Tissue is cut around the eye from this position. B) The eye is falls out and the underlying ocular muscles and optic nerve are cut. The empty eye socket is indicated with dashed circle. C) The same embryo is turned over and mounted with methyl-cellulose, with the intact eye facing up. <a href="https://www.jove.com/files/ftp_upload/50417/50417fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="6in" src="/files/ftp_upload/50417/50417fig2highres.jpg" width="600" /> 
Figure 2. Step-by-step 3D reconstruction of a confocal image stack. A) Open file (4104.1.ids) loaded within Fiji using Plugins&#62;LOCI&#62;Bioformat to select. B) After finding a slice with the region of interest, threshold adjustment is selected as shown. C) Threshold is adjusted to 214 using the top slider and apply is selected. D) 3D viewer is called as shown. E) The 3D reconstruction is shown of a zebrafish with the eye intact, for orientation. F) The image has been zoomed and rotated. G) A 360 degree rotation movie is made as shown. <a href="https://www.jove.com/files/ftp_upload/50417/50417fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/50417/50417fig3highres.jpg" width="600" /> 
Figure 3. Perspectives from 3D reconstruction. A) Medial perspective of 6 dpf embryo imaged with 10x objective, mouth is on the right, not gills inside mouth. B) Lateral of the same embryo, note fin is a loop in the middle. C) Same embryo imaged with 20x objective, medial perspective, note gill resolution. D) Lateral perspective of 20x objective imaging. The fin is on the right edge of the panel. E) Antero-medial view of 20x objective imaging, note gills inside mouth. F) Abdomen of the same embryo imaged with a 20x objective, head it to the right. Note vasculature on the yolk sac at the bottom right. <a href="https://www.jove.com/files/ftp_upload/50417/50417fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/50417/50417fig4highres.jpg" width="600" /> 
Figure 4. Intensity differences in 6 dpf embryo. Image of a 6 dpf reconstruction using a pseudocolor look-up-table for signal intensity. Mouth, brain, gills and yolk sac are labeled for orientation. <a href="https://www.jove.com/files/ftp_upload/50417/50417fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/50417/Zfish_still.jpg" width="500" /> 
Figure 5. Movie of reconstructed vascular system in a 4 dpf zebrafish. The fish was imaged at 2.5 &#956;m. The images were from imaging one half of the embryo. Compare vascular structures with structures in a GSI-treated zebrafish provided in Figure 6. Note the lower density of blood vessels in the head and larger gills. (See the "Zfish_spin.avi" supplemental file under Downloads)
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/50417/Zfish_still1.jpg" width="500" /> 
Figure 6. 3D Movie of vascular system in a GSI-treated embryo at 4 dpf. The fish was imaged at 2.5 &#956;m through from lateral to midline. Compare vascular structures with the control 4 dpf fish shown in Figure 5. The arched back and smaller size are typical in embryos treated with this chemical. (See the "GSI-treated_4dpf_fish.avi" supplemental file under Downloads)

Watch this Scientific Journal Video about Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish at JoVE.com

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

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

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

imaging-3d-reconstruction-cerebrovascular-structures-embryonic

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Biology

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

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

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

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

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Microdissection of Zebrafish Embryonic Eye Tissues

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post fertilization (dpf), the larvae will show the first visual response. Many genes have been identified to control a proper eye development, but we are far from a complete understanding of the underlying genetic architecture. Whole genome gene expression profiling is a useful tool to elucidate genetic regulatory network for eye development. However, the small size of the embryonic eye in zebrafish makes it challenging to obtain intact and pure eye tissues for expression analysis. For example, the anterior-posterior length of the eye between day 2 and 3 is only approximately 200-300 &mu;m, while the diameter of the lens is less 100 &mu;m. Also, the retinal pigment epithelium (RPE) underlying the retina is just a single-layer epithelium. While gene expression profiles can be obtained from the whole embryo, they do not accurately represent the expression of these tissues. Therefore pure tissue must be obtained for a successful gene expression profiling of eye development. To address this issue, we have developed an approach to microdissect intact retina and retina with RPE attached from 1-3 dpf, which cover major stages of eye morphogenesis. All procedures can be done with fine forceps and general laboratory supplies under standard stereomicroscopes. For retinal dissection, the single-layer RPE is removed and peeled off by brushing action and the preferential adherence of the RPE remnants to the surface of the culture plate for dissection. For RPE-attached retinal dissection, the adherence of RPE to the dissection plate is removed before the dissection so that the RPE can be completely preserved with the retina. A careful lifting action of this tissue can efficiently separate the presumptive choroid and sclera. The lens can be removed in both cases by a chemically etched tungsten needle. In short, our approach can obtain intact eye tissues and has been successfully utilized to study tissue-specific expression profiles of zebrafish retina1, 2 and retinal pigment epithelium3.

This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos.

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post ...

Immunostaining of Dissected Zebrafish Embryonic Heart

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology and genetics 1,2. About 100-200 embryos are readily available every week from a single pair of adult fish. The transparent embryos that develop ex utero make them ideal for assessing cardiac defects 3. The expression of any gene can be manipulated via morpholino technology or RNA injection 4. Moreover, forward genetic screens have already generated a list of mutants that affect different perspectives of cardiogenesis 5. 
Whole mount immunostaining is an important technique in this animal model to reveal the expression pattern of the targeted protein to a particular tissue 6. However, high resolution images that can reveal cellular or subcellular structures have been difficult, mainly due to the physical location of the heart and the poor penetration of the antibodies. 
 Here, we present a method to address these bottlenecks by dissecting heart first and then conducting the staining process on the surface of a microscope slide. To prevent the loss of small heart samples and to facilitate solution handling, we restricted the heart samples within a circle on the surface of the microscope slides drawn by an immEdge pen. After the staining, the fluorescence signals can be directly observed by a compound microscope. 
Our new method significantly improves the penetration for antibodies, since a heart from an embryonic fish only consists of few cell layers. High quality images from intact hearts can be obtained within a much reduced procession time for zebrafish embryos aged from day 2 to day 6. Our method can be potentially extended to stain other organs dissected from either zebrafish or other small animals.

A rapid way to conduct immunostaining of zebrafish embryonic heart is described. Compared to the whole mount immunostaining approach, this method dramatically increases the penetration of the antibodies, which allows obtaining high resolution images that reveal cellular/subcellular structures in the heart within a much reduced processing time.

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

A System for ex vivo Culturing of Embryonic Pancreas

The pancreas controls vital functions of our body, including the production of digestive enzymes and regulation of blood sugar levels1. Although in the past decade many studies have contributed to a solid foundation for understanding pancreatic organogenesis, important gaps persist in our knowledge of early pancreas formation2. A complete understanding of these early events will provide insight into the development of this organ, but also into incurable diseases that target the pancreas, such as diabetes or pancreatic cancer. Finally, this information will generate a blueprint for developing cell-replacement therapies in the context of diabetes.
 During embryogenesis, the pancreas originates from distinct embryonic outgrowths of the dorsal and ventral foregut endoderm at embryonic day (E) 9.5 in the mouse embryo3,4. Both outgrowths evaginate into the surrounding mesenchyme as solid epithelial buds, which undergo proliferation, branching and differentiation to generate a fully mature organ2,5,6. Recent evidences have suggested that growth and differentiation of pancreatic cell lineages, including the insulin-producing &beta;-cells, depends on proper tissue-architecture, epithelial remodeling and cell positioning within the branching pancreatic epithelium7,8. However, how branching morphogenesis occurs and is coordinated with proliferation and differentiation in the pancreas is largely unknown. This is in part due to the fact that current knowledge about these developmental processes has relied almost exclusively on analysis of fixed specimens, while morphogenetic events are highly dynamic.
 Here, we report a method for dissecting and culturing mouse embryonic pancreatic buds ex vivo on glass bottom dishes, which allow direct visualization of the developing pancreas (Figure 1). This culture system is ideally devised for confocal laser scanning microscopy and, in particular, live-cell imaging. Pancreatic explants can be prepared not only from wild-type mouse embryos, but also from genetically engineered mouse strains (e.g. transgenic or knockout), allowing real-time studies of mutant phenotypes. Moreover, this ex vivo culture system is valuable to study the effects of chemical compounds on pancreatic development, enabling to obtain quantitative data about proliferation and growth, elongation, branching, tubulogenesis and differentiation. In conclusion, the development of an ex vivo pancreatic explant culture method combined with high-resolution imaging provides a strong platform for observing morphogenetic and differentiation events as they occur within the developing mouse embryo.

A System for ex vivo Culturing of Embryonic Pancreas

Here, we describe a method for isolation, culture and manipulation of mouse embryonic pancreas. This represents an excellent ex vivo system for studying various aspects of pancreatic development, including morphogenesis, differentiation and growth. Pancreatic bud explants can be cultured for several days and used in a range of different applications, including whole-mount immunofluorescence and live imaging.

The pancreas controls vital functions of our body,  including the production of digestive enzymes and regulation of blood sugar  levels1. Although in the ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx mouse (for a review of these approaches see 1). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2 or sapje3) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4. Besides, chemical approaches are also possible, e.g. with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7.
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11.

The method presented here comprises the precise injury of live zebrafish embryos with high-energy laser pulses and the subsequent analysis of these injuries and their recovery with time. We also show how genetically labeled single or groups of skeletal muscle cells can be tracked during and after laser light induced damage.

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Live Cell Imaging and 3D Analysis of Angiotensin Receptor Type 1a Trafficking in Transfected Human Embryonic Kidney Cells Using Confocal Microscopy

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in transfected human embryonic kidney-293 (HEK) cells following stimulation with angiotensin II (Ang II). HEK cells are transiently transfected with plasmid DNA containing AT1aR tagged with enhanced green fluorescent protein (EGFP). Lysosomes are identified with a red fluorescent dye. Live-cell images are captured on a laser scanning confocal microscope after Ang II stimulation and analyzed by software in three dimensions (3D, voxels) over time. Live-cell imaging enables investigations into receptor trafficking and avoids confounds associated with fixation, and in particular, the loss or artefactual displacement of EGFP-tagged membrane receptors. Thus, as individual cells are tracked through time, the subcellular localization of receptors can be imaged and measured. Images must be acquired sufficiently rapidly to capture rapid vesicle movement. Yet, at faster imaging speeds, the number of photons collected is reduced. Compromises must also be made in the selection of imaging parameters like voxel size in order to gain imaging speed. Significant applications of live-cell imaging are to study protein trafficking, migration, proliferation, cell cycle, apoptosis, autophagy and protein-protein interaction and dynamics, to name but a few.

Here we present a protocol to image cells expressing green fluorescent protein-tagged angiotensin type 1a receptors during endocytosis initiated by angiotensin II treatment. This technique includes labeling lysosomes with a second fluorescent marker, and then utilizing software to analyze the co-localization of receptor and lysosome in three dimensions over time.