The authors would like to thank Laura Anne Lowery who developed this technique in the lab. HS is supported by NIHMH70926. Jennifer Gutzman was supported by the CSBi/Merck postdoctoral fellowship.

Protocol
Part 1: Preparing for microinjection
<ol>
	<li>Prepare for the injection by pulling capillary needles using Sutter Instruments needle puller.</li>
	<li>Fill the needle with a fluorescent dye (such as Texas-Red Dextran).</li>
	<li>Mount the needle in a micromanipulator and microinjection apparatus.</li>
	<li>Cut the needle to the appropriate size at an angle to create a beveled tip.</li>
	<li>Measure the drop size and check that it is between 1-2nl per injection in oil.</li>
</ol>
Part 2: Preparing the embryos
<ol>
	<li>Inject the embryos 30 minutes earlier than the stage of interest. For example, to study the ventricles at 24 hours post fertilization (hpf), inject the embryos at 23.5 hpf. Stage the embryos according to Kimmel et al. (4).</li>
	<li>Under the stereomicroscope, carefully remove the chorion from the embryos using forceps.</li>
	<li>Using a plastic dish coated with 1% agarose, poke holes in the agarose with a plastic 1-200 &#956;l pipette tip. Carefully remove the agarose plugs from the holes using forceps.</li>
	<li>Transfer the embryos in embryo media into the agarose dish with holes.</li>
	<li>Add tricaine (made according to Westerfield (5)) to the media to anesthetize the embryos and prevent movement during the experiment.</li>
	<li>Gently place the embryo tail down into the hole, and orient it so that the injection apparatus is on the posterior side of the embryo.</li>
</ol>
Part 3: Injecting the brain ventricle
<ol>
	<li>To inject the brain ventricle, arrange the micromanipulation setup so that the needle tip is in the same field of view as the embryo on high power.</li>
	<li>Carefully put the needle through the thin roofplate of the hindbrain just posterior to the r0/r1 hingepoint without hitting the brain tissue below.</li>
	<li>Inject just enough dye to completely fill the ventricles. It may take several injections to fill the ventricle space, depending on the stage of the embryo.</li>
</ol>
Part 4: Imaging
<ol>
	<li>Using a clean 1% agarose-coated dish, poke new holes in the dish, remove the plugs, and add tricaine to anesthetize the embryos and prevent movement.</li>
	<li>Place the tail of the ventricle-injected embryo into the hole and position for a dorsal image.</li>
	<li>Take an image using transmitted light.</li>
	<li>It is very critical to NOT MOVE THE EMBRYO OR MICROSCOPE.</li>
	<li>Change the settings on the microscope and take an image with fluorescent light.</li>
	<li>Reposition the embryo to take lateral images with both transmitted light and fluorescent light.</li>
	<li>Save the images as TIFF files for image processing in Photoshop.</li>
</ol>
Part 5: Image Processing
<ol>
	<li>To process the images in Photoshop, open the transmitted light and fluorescent light images of the same embryo taken in the same position. Be sure all of the settings are the same for each image.</li>
	<li>Drag your fluorescent image onto the transmitted light image and line them up exactly.</li>
	<li>With the fluorescent image selected, select &#8220;Image,&#8221; &#8220;Adjustments,&#8221; and then &#8220;Replace color&#8221; and change the black background to white.</li>
	<li>In the &#8220;Replace color&#8221; window adjust the &#8220;fuzziness&#8221; factor to the right until the fluorescent images looks uniform in color.</li>
	<li>In the &#8220;Layers&#8221; window, select &#8220;Multiply&#8221; to view the overlay images. Your ventricle space image should match the transmitted light image exactly.</li>
	<li>Save this file and repeat for any other images.</li>
</ol>
Representative Results/Outcome
Proper ventricle injection images are shown in Figure 1 panels A-D, while an example of incorrect injection images are shown in E-H. Proper injections have sharp edges and non-diffuse dye. Incorrect injection, which occurs when the needle is inserted too far into the ventricle and hits the brain tissue below, can result in dye that is visible outside of the ventricle space and in the yolk.
<img alt="Figure 1" height="742" src="/files/ftp_upload/1218/1218fig1.jpg" />
Figure 1. Ventricle injections of 25 hpf wild type embryos. (A-D) Correct injection method and (E-H) incorrect injection method are shown as examples. (A,E) Dorsal brightfield images with corresponding fluorescent images (B,F) and the overlay of the fluorescent and brightfield images (C,G) for both correct and incorrect injection methods shown. Arrows indicate regions where dye has gone beyond the brain ventricle space due to an incorrect ventricle injection (see &#8220;Troubleshooting&#8221;). Anterior is to the left in all images.
Troubleshooting:
<table border="1">
	<tbody>
		<tr style="font-weight:bold; font-size:16px;">
			<td>Problem</td>
			<td>Cause</td>
			<td>Remedy</td>
		</tr>
		<tr>
			<td>You are squashing the embryo with the needle.</td>
			<td>Your needle is too blunt or the tip is too large.</td>
			<td>Start with a new needle and do not break it as far back. Bevel your needle to make it sharp like a syringe needle.</td>
		</tr>
		<tr>
			<td>The dye is throughout the embryo and the yolk (see Fig 1E-H).</td>
			<td>Your needle went through the tissue and didn&#8217;t stay in the ventricle space.</td>
			<td>Your needle may not be sharp enough, so you are unable to control the speed of puncture.</td>
		</tr>
		<tr>
			<td>The dye is too faint or not in the forebrain.</td>
			<td>You did not use a high enough pressure for the injection, or your needle was not far enough anterior to reach the forebrain when you injected the dye.</td>
			<td>Increase your injection pressure or place your needle slightly more anterior during the injection. You can put the needle through the midbrain-hindbrain boundary if more detail is required in the forebrain. You can also simply wait a bit for the dye to further diffuse before imaging.</td>
		</tr>
		<tr>
			<td>Your fluorescent images are dark around the edges after the color replacement in Photoshop.</td>
			<td>You forgot to change the &#8220;fuzziness&#8221; setting in the color replace window.</td>
			<td>Move the &#8220;fuzziness&#8221; all the way to the right in the color replace window.</td>
		</tr>
		<tr>
			<td>The needle will not puncture the hindbrain roofplate.</td>
			<td>Your needle is not cut properly.</td>
			<td>Make a new needle. If you are doing a lot of injections you may need several needles as they can get dull.</td>
		</tr>
	</tbody>
</table>

<ol>
	<li>Gutzman, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+of+the+zebrafish+midbrain-hindbrain+boundary+constriction+requires+laminin-dependent+basal+constriction.">Formation of the zebrafish midbrain-hindbrain boundary constriction requires laminin-dependent basal constriction.</a> Mech. Dev. 125, 974-983 (2008).</li><li>Lowery, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+and+Classification+of+Zebrafish+Brain+Morphology+Mutants.">Characterization and Classification of Zebrafish Brain Morphology Mutants.</a> Anat. Rec. (Hoboken). 292, 94-106 (2009).</li><li>Lowery, L. A., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Initial+formation+of+zebrafish+brain+ventricles+occurs+independently+of+circulation+and+requires+the+nagie+oko+and+snakehead/atp1a1a.1+gene+products.">Initial formation of zebrafish brain ventricles occurs independently of circulation and requires the nagie oko and snakehead/atp1a1a.1 gene products.</a> Development. 132, 2057-2067 (2005).</li><li>Kimmel, C. B. <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> Dev. Dyn. 203, 253-310 (1995).</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. , (1995).</li></ol>

In this video we demonstrate how to inject fluorescent dye (Texas-Red Dextran) into developing zebrafish brain ventricles. This method is used to visualize the brain ventricle space in contrast to the surrounding neuroepithelium and is extremely useful for determining the shape of the ventricle space as well as the shape of the surrounding brain tissue. It allows us to better understand the process of brain ventricle formation and brain morphogenesis over time in the live embryo. This technique is an excellent tool for...

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Forceps</td>
<td>Tool</td>
<td>Fine Science Tools</td>
<td>11252-20</td>
<td>&nbsp;</td>
<tr>
<td>Capillary Tubes (needles)</td>
<td>Tools</td>
<td>Frederick Haer and Co.</td>
<td>30-30-1</td>
<td>&nbsp;</td>
<tr>
<td>Agarose-Coated dishes</td>
<td>Tools</td>
<td>Agarose (Seakem GTG; Lonza) Dishes (Corning) </td>
<td>Agarose (50027) Dishes (430166) </td>
<td>Make agarose approximately 2-3 mm deep in the dish depending on the stage of embryo you are injecting</td>
</tr>
<tr>
<td>Dissecting Microscope</td>
<td>Microscope</td>
<td>Zeiss Stereomicroscope Stemi SV 6 </td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Microscope Camera with Fluorescence capabilities and with image capture software</td>
<td>Tools</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>1-200 &mu; pipette tips</td>
<td>Tools</td>
<td>Tip One, US Scientific</td>
<td>1111-0806</td>
<td>These are the perfect size to poke the holes in the agarose and have the embryos fit perfectly.</td>
</tr>
<tr>
<td>Photoshop</td>
<td>Image Analysis</td>
<td>Adobe</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Embryo Loop</td>
<td>Tools</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>Make these yourself with a glass pipette and hair or fishing line and seal with beeswax.</td>
</tr>
<tr>
<td>Microinjector</td>
<td>Tools</td>
<td>Medical Systems Research Products Harvard Apparatus</td>
<td>PL1-100</td>
<td>&nbsp;</td>
<tr>
<td>Needle Puller</td>
<td>Tools</td>
<td>Sutter Instruments</td>
<td>&nbsp;</td>
<td>Settings (Heat 785, Pull 70, Velocity 40, Time 70).</td>
</tr>
<tr>
<td>Micromanipulator</td>
<td>Tools</td>
<td>Narishige</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Fluorescent dye Texas-Red Dextran 70,000MW</td>
<td>Reagent</td>
<td>Molecular Probes</td>
<td>D1830</td>
<td>Other Molecular weights are available if desired.</td>
</tr>
<tr>
<td>Tricaine (3-aminobenzoic acid ethyl ester)</td>
<td>Reagent</td>
<td>Sigma</td>
<td>A-5040</td>
<td>&nbsp;</td>
<tr>
<td>Embryo Media</td>
<td>Reagent</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>According to Westerfield (5)</td>
</tr>		</tbody>
		</table>
		</div>

zebrafish brain ventricle injection

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved cavities within the brain that are filled with cerebrospinal fluid.  In zebrafish, after neural tube formation, the neuroepithelium undergoes a series of constrictions and folds while it fills with fluid resulting in brain ventricle formation.  In order to understand the process of ventricle formation, and the neuroepithelial shape changes that occur at the same time, we needed a way to visualize the ventricle space in comparison to the brain tissue.  However, the nature of transparent zebrafish embryos makes it difficult to differentiate the tissue from the ventricle space.  Therefore, we developed a brain ventricle injection technique where the ventricle space is filled with a fluorescent dye and imaged by brightfield and fluorescent microscopy.  The brightfield and the fluorescent images are then processed and superimposed in Photoshop.  This technique allows for visualization of the ventricle space with the fluorescent dye, in comparison to the shape of the neuroepithelium in the brightfield image.  Brain ventricle injection in zebrafish can be employed from 18 hours post fertilization through early larval stages.  We have used this technique extensively in our studies of brain ventricle formation and morphogenesis as well as in characterizing brain morphogenesis mutants (1-3).

Watch this Scientific Journal Video about Zebrafish Brain Ventricle Injection at JoVE.com

Zebrafish Brain Ventricle Injection

After neural tube formation, the neuroepithelium constricts and folds while the tube fills with embryonic cerebrospinal fluid (eCSF) to form the embryonic brain ventricles. We developed this ventricle injection technique to better visualize the fluid filled space in contrast to the neuroepithelial shape in a live embryo.

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved ...

Research

JoVE Journal

Biology

21.3K Views.  Whitehead Institute for Biochemical Research. After neural tube formation, the neuroepithelium constricts and folds while the tube fills with embryonic cerebrospinal fluid (eCSF) to form the embryonic brain ventricles. We developed this ventricle injection technique to better visualize the fluid filled space in contrast to the neuroepithelial shape in a live embryo.

Video: Zebrafish Brain Ventricle Injection

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 ...

Retro-orbital Injection in Adult Zebrafish

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is often the most effective and noninvasive form of delivery of an agent of interest. In the zebrafish (Danio rerio), IV injection of drugs has long been a challenge because of the small vessel diameter. This has also proved a significant hurdle for the injection of cells during hematopoeitic stem cell transplantation. Historically, injections into the bloodstream were done directly through the heart. However, this intra-cardiac procedure has a very high mortality rate as the heart is often punctured during injection leaving the fish prone to infection, massive blood loss or fatal organ damage. Drawing on our experience with the mouse, we have developed a new injection procedure in the zebrafish in which the injection site is behind the eye and into the retro-orbital venous sinus. This retro-orbital (RO) injection technique has been successfully employed in both the injection of drugs in the adult fish as well as transplantation of whole kidney marrow cells. RO injection has a much lower mortality rate than traditional intra-cardiac injection. Fish that are injected retro-orbitally tend to bleed less following injection and are at a much lower risk of injury to a major organ like the heart. Further, when performed properly, injected cells and/or drugs quickly enter the bloodstream allowing compounds to exert their effect on the whole fish and kidney cells to easily home to their niche. Thus, this new injection technique minimizes mortality while allowing efficient delivery of material into the bloodstream of adult fish. Here we exemplify this technique by retro-orbital injection of Tg(globin:GFP) cells into adult casper fish as well as injection of a red fluorescent dye (dextran, Texas Red ) into adult casper fish. We then visualize successful injections by whole animal fluorescence microscopy.

Here we show how to do retro-orbital injection in adult zebrafish.

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is ...

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

Intraperitoneal Injection into Adult Zebrafish

A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, this method makes it difficult to know how much reagent is absorbed or taken up per fish. Some experimental questions, particularly those related to metabolic studies, may be better addressed by delivering a defined quantity to each fish, based on weight. Here we present a method for intraperitoneal (IP) injection into adult zebrafish. Injection is into the abdominal cavity, posterior to the pelvic girdle. This procedure is adapted from veterinary methods used for larger fish. It is safe, as we have observed zero mortality. Additionally, we have seen bleeding at the injection site in only 5 out of 127 injections, and in each of those cases the bleeding was brief, lasting several seconds, and the quantity of blood lost was small. Success with this procedure requires gentle handling of the fish through several steps including fasting, weighing, anesthetizing, injection, and recovery. Precautions are required to minimize stress throughout the procedure. Our precautions include using a small injection volume and a 35G needle. We use Cortland salt solution as the vehicle, which is osmotically balanced for freshwater fish. Aeration of the gills is maintained during the injection procedure by first bringing the fish into a surgical plane of anesthesia, which allows slow operculum movements, and second, by holding the fish in a trough within a water-saturated sponge during the injection itself. We demonstrate the utility of IP injection by injecting glucose and monitoring the rise in blood glucose level and its subsequent return to normal. As stress is known to increase blood glucose in teleost fish, we compare blood glucose levels in vehicle-injected and non-injected adults and show that the procedure does not cause a significant rise in blood glucose.

We demonstrate intraperitoneal injection into adult zebrafish. We use a 10 &mu;l NanoFil microsyringe controlled by a Micro4 controller and UltraMicroPump III. This demonstration includes the use of cold water as an anesthetic.

A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, ...

Direct Delivery of MIF Morpholinos Into the Zebrafish Otocyst by Injection and Electroporation Affects Inner Ear Development

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the eye, brain, and somites of zebrafish. The advantage of electroporation over other methods of genetic manipulation is that specific tissues can be targeted, both spatially and temporally, for the introduction of macromolecules by the application of electrical current. Here we describe the use of electroporation for transfecting mif and mif-like morpholinos into the tissues of the developing inner ear of the zebrafish. In past studies, mif morpholino injected into embryos at the 1- to 8-cell stage resulted in widespread morphological changes in the nervous system and eye, as well as the ear. By targeting the tissues of the inner ear at later stages in development, we can determine the primary effects of MIF in the developing inner ear, as opposed to secondary effects that may result from the influence of other tissues. By using phalloidin and acetylated tubulin staining to study the morphology of neurons, neuronal processes, and hair cells associated with the posterior macula, we were able to assess the efficacy of electroporation as a method for targeted transfection in the zebrafish inner ear. The otic vesicles of 24hpf embryos were injected with morpholinos and electroporated and were then compared to embryos that had received no treatment or had been only injected or electroporated. Embryos that were injected and electroporated showed a decrease in hair cell numbers, decreased innervation by the statoacoustic ganglion (SAG) and fewer SAG neurons compared with control groups. Our results showed that direct delivery of morpholinos into otocysts at later stages avoids the non-specific nervous system and neural crest effects of morpholinos delivered at the 1-8 cell stage. It also allows examination of effects that are directed to the inner ear and not secondary effects on the ear from primary effects on the brain, neural crest or periotic mesenchyme.

A method to deliver morpholinos directly into the zebrafish otocyst at 24hpf has been developed. Using microinjection of morpholinos into the lumen of otic vesicle and electroporation to effect penetration, we were able to bypass the effect of morpholinos on the brain and obtain effects specific to the inner ear.

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the ...

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 ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

Microgavage of Zebrafish Larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

Reverse Genetic Morpholino Approach Using Cardiac Ventricular Injection to Transfect Multiple Difficult-to-target Tissues in the Zebrafish Larva

The zebrafish is an important model to understand the cell and molecular biology of organ and appendage regeneration. However, molecular strategies to employ reverse genetics have not yet been adequately developed to assess gene function in regeneration or tissue homeostasis during larval stages after zebrafish embryogenesis, and several tissues within the zebrafish larva are difficult to target. Intraventricular injections of gene-specific morpholinos offer an alternative method for the current inability to genomically target zebrafish genes in a temporally controlled manner at these stages. This method allows for complete dispersion and subsequent incorporation of the morpholino into various tissues throughout the body, including structures that were formerly impossible to reach such as those in the larval caudal fin, a structure often used to noninvasively research tissue regeneration. Several genes activated during larval finfold regeneration are also present in regenerating adult vertebrate tissues, so the larva is a useful model to understand regeneration in adults. This morpholino dispersion method allows for the quick and easy identification of genes required for the regeneration of larval tissues as well as other physiological phenomena regulating tissue homeostasis after embryogenesis. Therefore, this delivery method provides a currently needed strategy for temporal control to the evaluation of gene function after embryogenesis.&#160;

An adaptable reverse genetic method for zebrafish to assess gene function during later stages of development and physiological homeostasis such as tissue regeneration using intraventricular injections of gene-specific morpholinos.

The zebrafish is an important model to understand the cell and molecular biology of organ and appendage regeneration. However, molecular strategies to ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...