Name: in vivo electroporation of morpholinos into the adult zebrafish retina
Uploaded: 2011-12-27T12:00:00
Duration: 6 min 55 s
Description: Watch this Scientific Journal Video about In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina at JoVE.com

The authors would like to thank the Freimann Life Science Center and Center for Zebrafish Research staff for their care and maintenance of the zebrafish.

1. Morpholino preparation:
<ol>
	<li>All morpholino oligos should be custom-designed and ordered through GeneTools (Philomath, OR). We use the positively-charged lissamine-tagged morpholinos to drive the electroporated morpholinos towards the negative electrode. We tried, unsuccessfully, to target negatively-charged fluorescein-tagged morpholinos to the positive electrode using this technique. Dilute 300 nmol of morpholino into 100 &mu;l of nuclease-free water to yield a 3 mM solution. Aliquot the morpholino into multiple paraffin-sealed microcentrifuge tubes. Store away from light at room temperature.</li>
	<li>Determine the morpholino concentration by diluting 5 &mu;l of morpholino (or water used to dilute the morpholino as a blank) in 95 &mu;l of 0.1 N HCl. Set the spectrophotometer wavelength (&lambda;) to 265 nm. Calculate the constant for your morpholino using the following equation: 
	 
	Constant = molecular weight particular to your morpholino X 1000/molar absorbance. 
	 </li>
	<li>You will find the molecular weight and molar absorbance data on the "Oligo Properties" sheet provided with each morpholino. Analyze the blank to verify baseline readings and take readings of each diluted morpholino. Multiply the absorbance at 265 nm of each morpholino by its determined constant and by the dilution factor to get the concentration in ng/&mu;l. Divide this number by the molecular weight to determine the concentration in mM. Dilute the morpholino, if necessary, to a working concentration.</li>
</ol>
2. Remove outer corneal layer:
<ol>
	<li>Anesthetize 6-12 month old adult zebrafish in Tricaine or 2-phenoxyethanol at 1.0 mg/ml in zebrafish tank water.</li>
	<li>Wrap zebrafish in a moistened piece of paper towel, covering the gills, but leaving the eye exposed. The wrapped fish is placed under a stereoscope and the magnification is increased until the diameter of the eye fills approximately one-third of the visual field (Figure 1).</li>
	<li>Using Dumont #5 tweezers, grab the outer cornea near the optic fissure. This tissue is colored in red and is labeled "OC" in Figure 1. There is a slight protrusion or lip indicated by the arrow labeled "CL". Pull at a low angle (i.e. 10 degrees) across the eye to remove the outer cornea. With practice, this can be completed in one motion. If the tweezers slip, re-grab the tissue and again pull at a low angle until removed. A well-attached cornea can cause the eye to be dislodged from the socket when pulled. You can keep the eye from becoming dislodged by steadying with another set of tweezers.</li>
</ol>
3. Corneal incision and morpholino injection:
<ol>
	<li>Immediately following the removal of the outer cornea, make a small incision in the cornea where the pupil meets the iris (Figure 2). This is done using a sapphire blade scalpel (See Materials).</li>
	<li>Load 0.5 &mu;l of the 3.0 mM morpholino solution into a Hamilton syringe equipped with a 33 gauge removable, blunt-end needle (See Materials). Pipette up and down to remove air bubbles in the line. Carefully insert the needle into the vitreous through the incision (Figure 2). Do not insert the needle too far or you will puncture the retina or displace the lens.</li>
	<li>Slowly inject the morpholino solution into the vitreal space. Depending on the age of the fish, the vitreal space will hold ~ 0.5 &mu;l. You should visualize uncolored vitreous leaking out of the incision as it is displaced by the morpholino solution. Inject until a small amount of morpholino solution leaks out of the incision. You should clearly visualize the lissamine-tagged morpholino inside the eye.</li>
	<li>Following the injection, return the fish to the tank to recover. We usually only inject the left eye, using the right eye as an uninjected control, but electroporation of both eyes is possible.</li>
</ol>
4. Electroporation:
<ol>
	<li>Following the injection of approximately 10 fish, re-anesthetize an injected fish and wrap it in a moistened piece of paper towel, just covering the gills, but leaving the eye exposed. Transfer the fish to a Petri dish. Put on latex or similar gloves to avoid exposure to the anesthesia. While gently holding the fish to the bottom of the dish, fill the dish with anesthesia (Figure 3, left panel).</li>
	<li>A 3-mm-diameter platinum plate electrode (See Materials) localizes the electroporation pulses to approximately one-half of the retina. We typically target the dorsal retina. Gently press the positive electrode downward on the ventral half of the eye. With the outer cornea removed, the eye is easily rotated, exposing the dorsal portion of the eyeball.</li>
	<li>While still pressing down on the ventral side of the eye, place the negative electrode near (~1 - 2 mm) the exposed dorsal half of the eye. Be careful not to touch the negative electrode directly to the eye, which will result in damaging the dorsal retina. Once you determine the appropriate distance between the electrodes, use the adjustment screw on the handle of the electrodes to maintain that spread when electroporating. This keeps the negative electrode an optimal distance from the eyeball when the positive electrode is pressed down in position.</li>
	<li>Electroporate the eye using a CUY21 Square Wave Electroporator (See Materials). The electroporation parameters should be set to two consecutive 50-msec pulses, at 75 V with a 1-sec pause between pulses.</li>
	<li>Return fish to the tank. We typically start our protocol for constant light-induced retinal degeneration immediately following electroporation.</li>
</ol>
5. Representative results:
<ol>
	<li>For 3-5 days following the electroporation (depending on the stability of the morpholino), the lissamine label can be visualized in all of the retinal layers in a sectioned retina (Figure 4). This serves as a confirmation for the presence of the labeled morpholino.</li>
	<li>We can achieve full protein knockdown up to 3-5 days following electroporation, depending of the efficacy of the morpholino (Figure 5). Typically, during light treatment studies, we harvest eyes at 3 or 4 days after morpholino electroporation. This is done by removing the entire eye with Dumont #5B forceps with curved ends (Figure 6; Materials) and processing the tissue for immunohistochemistry 15. As with any knockdown technique, a demonstration of efficient knockdown must be shown in order to validate the phenotype. Therefore, we suggest obtaining antisera for your protein of interest. Alternatively, if and antibody is not available and the morpholino is directed to either the splice acceptor or donor site in the pre-mRNA, it will block efficient splicing of the mRNA. In this case, it is possible to use reverse transcriptase-PCR to amplify the flanking sequences in the morphant RNA and determine the efficiency of blocking the normal splicing pattern.</li>
	<li>When the retinas are sectioned from nasal to temporal on the dorsal ventral axis, the shaded blue boxes (Figure 6) represent the areas of the retina that are most often damaged from the electroporation event itself. The shaded pink box represents the area that is targeted for morpholino introduction into retinal cells by the electroporation.</li>
</ol>
<img alt="Figure 1." src="/files/ftp_upload/3603/3603fig1.jpg" /> 
Figure 1. Schematic depicting the removal of the epithelial layers of the cornea. The left panel shows the major structural features of the eye, including the lens (L), the vitreous (Vit), the retina (R), the outer cornea (OC), the inner cornea (IC), and the corneal protrusion (CL). The orientation of the eye is shown in the lower right of the first panel and is labeled anterior (A), posterior (P), dorsal (D), and ventral (V). The red coloration indicates the outer cornea that is removed. The second panel shows how the forceps are used to grasp the corneal protrusion and to pull the outer cornea across the eye at a low angle (third panel). The right panel shows how a second pair of forceps can be used to steady the eye while pulling the cornea.
<img alt="Figure 2." src="/files/ftp_upload/3603/3603fig2.jpg" /> 
Figure 2. Schematic of the steps involved in the intravitreal injection of morpholino. A small incision is made in the eye where the pupil meets the iris (left panel). The resulting gap should be just large enough for a 33 gauge needle, which is filled with the morpholino solution, to be inserted (middle panel). Slowly inject 0.5 &mu;l of solution into the vitreous (right panel). You should visualize a small amount of vitreal solution come out of the incision as the vitreous is partially replaced by with lissamine-tagged morpholino (right panel).
<img alt="Figure 3." src="/files/ftp_upload/3603/3603fig3.jpg" /> 
Figure 3. Schematic of the general electroporation procedure. The anesthetized fish, which was intravitreally injected with the morpholino, is gently wrapped in damp paper towel (left panel). The paper towel should cover the gills, but not obstruct the eye (middle panel). Gently press the positive electrode down on the ventral part of the eye, allowing the dorsal half of the eye to rotate out of the socket. Place the negative electrode approximately 1 mm from the dorsal half of the eye and start the electroporation (right panel).
<img alt="Figure 4." src="/files/ftp_upload/3603/3603fig4.jpg" /> 
Figure 4. Confocal image comparing a retina electroporated without intravitreally injecting a lissamine-tagged morpholino (A), and a retina intravitreally injected and electroporated with the lissamine-tagged morpholino (B). The lissamine label is found in all the retinal layers and cell types. ROS, rod outer segments; ONL, outer nuclear layer; INL, inner nuclear layer; GCL, ganglion cell layer.
<img alt="Figure 5." src="/files/ftp_upload/3603/3603fig5.jpg" /> 
Figure 5. Confocal images comparing PCNA immunolocalization in retinas electroporated with a control morpholino (left panels) and retinas electroporated with an anti-PCNA morpholino (right panels) prior to light-induced photoreceptor cell death. Dark-adapted adult albino zebrafish were placed in constant intense light for 1, 2, or 3 days after electroporation of the morpholino. The control morpholino failed to repress increasing PCNA expression in the light-damaged retinas. The anti-PCNA morpholino efficiently knocked down PCNA protein expression through three days of constant intense light damage.
<img alt="Figure 6." src="/files/ftp_upload/3603/3603fig6.jpg" /> 
Figure 6. Schematic (left panel) showing enucleation of the eye for processing and cryosectioning, which is done along the dorsal ventral axis. The pink shaded box (right panel) shows the retinal region with the greatest amount of protein knockdown using this technique. The blue shaded boxes represent areas that are most often damaged by the electroporation event. The orientation of the eye is labeled anterior (A), posterior (P), dorsal (D), and ventral (V).

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Neurosci. 27, 1712-1724 (2007).</li><li>Hitchcock, P., Ochocinska, M., Sieh, A., Otteson, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistent+and+injury-induced+neurogenesis+in+the+vertebrate+retina.">Persistent and injury-induced neurogenesis in the vertebrate retina.</a> Prog. Retin. Eye Res. 23, 183-194 (2004).</li><li>Raymond, P. A., Barthel, L. K., Bernardos, R. L., Perkowski, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+characterization+of+retinal+stem+cells+and+their+niches+in+adult+zebrafish.">Molecular characterization of retinal stem cells and their niches in adult zebrafish.</a> B.M.C Dev. Biol. 6, 36-36 (2006).</li><li>Vihtelic, T. S., Hyde, D. 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We have nothing to disclose

Recently, two microarray analyses examined the gene expression changes that occurred during the regeneration of either the light-damaged retina or a surgically-excised retinal patch10, 16. Both studies revealed numerous genes that exhibited significant changes in expression, either increasing or decreasing, that likely are required for different events that occur during retinal regeneration, such as re-entry of the M&uuml;ller glia into the cell cycle, continued proliferation and migration of the neuronal progenitors to the damaged retinal layer, and the differentiation of the neuronal progenitors into the correct neuronal cell type. While a direct comparison of these two microarray data sets reveal candidate genes that may be involved in these cellular events, ultimately these genes and their encoded proteins must be tested to determine if they are necessary for neuronal regeneration. Here, we describe a powerful loss-of-function approach to conditionally knockdown proteins of interest in the adult zebrafish retina. This technique has been used to study both extracellular and intracellular proteins, as well as signaling molecules, transcription factors and proteins required for DNA replication 17-19. Thus, this method has greatly aided our understanding of the molecular mechanism underlying adult retinal regeneration in zebrafish.
		We tested multiple electroporation parameters (voltage, number of pulses, time between pulses) before a successful protocol was achieved. A previous report of electroporating morpholinos into the regenerating zebrafish caudal fin used 10 pulses at a low voltage (15 V) 20. Because the adult zebrafish retina is surrounded by multiple cell layers and is not readily accessible to tweezer electrodes, the parameters used in the fin were unsuccessful at efficiently electroporating morpholinos into the adult retina. Conversely, a high voltage (100 V) often resulted in death to the fish. Recently, 5 pulses of 80 V was described to electroporate plasmid DNA into the newborn mouse pup retina 21. We found that a slightly less intense 2 pulses of 75 V resulted in successful electroporation of the morpholinos into the adult zebrafish retina with survival of 100% of the treated fish. Additionally, we discovered that the removal of the outer most (or external-most) component of the cornea greatly increased the electroporation efficiency. Increasing the number of pulses may eliminate the need for removal of this tissue but may also cause greater tissue damage. Extensive control studies demonstrated that these parameters did not cause any significant retinal cell death or alter the specific cellular responses observed during photoreceptor regeneration 19. Gain-of-function studies are not currently possible using this technique in vivo, due to our inability to consistently electroporate large ribonucleic acids into all retinal cells. However, the success of electroporating plasmids into the mouse retina in vivo and the zebrafish retina in explants suggests it is still a possibility in zebrafish 21, 22.
		One of the strengths of this technique is the electroporation appears to efficiently introduce the morpholino into all of the different retinal cell types. However, one potential weakness was that the electroporation efficiently delivered the morpholino into only the dorsal and central retinal regions. A second electroporation event toward the ventral half of the retina results in good targeting to all layers, but often results in damage. This spatial restriction likely was due to the shape and placement of the electrodes. The use of custom-designed cup shaped electrodes that could be placed around the eye may improve this weakness. This spatial restriction of the morpholino delivery limits the use of this technique and precludes the use of an assay such as ERG analysis that would require a global assessment of the retina. It should be noted that as with any morpholino experiment, it is advisable to confirm your results using a second, non-overlapping morpholino to the target mRNA. In some cases, it is possible to also electroporate two different morpholinos simultaneously to knockdown the expression of two different proteins. We demonstrated this by knocking down the expression of both the Pax6a and Pax6b proteins, individually and in combination 18.
		Although we demonstrated the use of this technique with our light damage model, it could likely be used to study regeneration in other damage models 2-8 or used to examine the function of proteins in the undamaged retina. For example, electroporation of morpholinos could be used to knockdown the expression of specific channels or signal transduction molecules in ganglion or amacrine cells and then study the function of these signaling pathways in visual processing. However, as morpholinos only affect translation of new protein, one would have to wait until endogenous protein turnover occurs before an assay could be performed. Depending on the stability of the protein, that could range from hours to days 18.

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>CUY21-EDIT or CUY21-SC Square wave electroporator</td>
 <td>Protech International</td>
 <td>CUY21EDIT or CUY21SC</td>
 <td>Both units work for this protocol</td>
 </tr>
 <tr>
 <td>3-mm diameter paddle electrodes</td>
 <td>Protech International</td>
 <td>CUY 650-P3</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Morpholino</td>
 <td>GeneTools, LLC</td>
 <td>&nbsp;
 </td>
 <td>Morpholino should be custom designed to your protein of interest</td>
 </tr>
 <tr>
 <td>2-Phenoxyethanol</td>
 <td>Sigma</td>
 <td>77861-1L</td>
 <td>Anesthesia; dilute 1:1000 in fish system water for procedure, 1:500 for euthanasia</td>
 </tr>
 <tr>
 <td>Dumont #5 tweezers</td>
 <td>World Precision Instruments</td>
 <td>14095</td>
 <td>Any #5 tweezers with work</td>
 </tr>
 <tr>
 <td>Dumont #5B tweezers with angled tips</td>
 <td>World Precision Instruments</td>
 <td>500234</td>
 <td>Used to enucleate eye following euthanasia</td>
 </tr>
 <tr>
 <td>Sapphire blade, double edge lance, 1 mm wide, with Sapphire knife handle</td>
 <td>World Precision Instruments</td>
 <td>Blade: 500314 
 Handle: 500317</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Hamilton syringe with removable 33 gauge blunt-end needle</td>
 <td>Hamilton company</td>
 <td>Syringe: 87930 
 Needle: 7762-06</td>
 <td>Remove needle that comes with syringe and replace with 33-gauge blunt-end needle</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina at JoVE.com

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.

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 ...

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Research

JoVE Journal

Biology

Organotypic Culture of Adult Rabbit Retina

Organotypic culture systems of functional neural tissues are important tools in neurobiological research. Ideally, such a system should be compatible with imaging techniques, genetic manipulation, and electrophysiological recording. Here we present a simple interphase tissue culture system for adult rabbit retina that requires no specialized equipment and very little maintenance. We demonstrate the dissection and incubation of rabbit retina and particle-mediated gene transfer of plasmids encoding GFP or a variety of subcellular markers into retinal ganglion cells. Rabbit retinas cultured this way can be kept alive for up to 6 days with very little changes of the overall anatomical structure or the morphology of individual ganglion- and amacrine cells.

This article demonstrates the dissection and incubation of rabbit retina and particle-mediated gene transfer of plasmids encoding GFP or a variety of subcellular markers into retinal ganglion cells.

Organotypic culture systems of functional neural tissues are important tools in neurobiological research. Ideally, such a system should be compatible with ...

Single Cell Electroporation in vivo within the Intact Developing Brain

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. The distinct advantage of this technique is that experimental manipulations may be performed on individual cells while leaving the surrounding tissue unaltered, thereby distinguishing cell-autonomous effects from those resulting from global treatments. When combined with advanced in vivo imaging techniques, SCE of fluorescent markers permits direct visualization of cellular morphology, cell growth, and intracellular events over timescales ranging from seconds to days. While this technique is used in a variety of in vivo and ex vivo preparations, we have optimized this technique for use in Xenopus laevis tadpoles. In this video article, we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the albino Xenopus tadpole. We also discuss methods to optimize yield, and show examples of live two-photon fluorescence imaging of neurons fluorescently labeled by SCE.

Single-cell electroporation (SCE) is a specialized technique allowing delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. Here we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the Xenopus laevis tadpole.

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact ...

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 ...

Dissection of Organs from the Adult Zebrafish

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental analysis of the embryo and larva is extensive and the morphology has been well documented, descriptions of adult zebrafish anatomy and studies of development of the adult structures and organs, together with techniques for working with adults are lacking. The organs of the larva undergo significant changes in their overall structure, morphology, and anatomical location during the larval to adult transition.  Externally, the transparent larva develops its characteristic adult striped pigment pattern and paired pelvic fins, while internally, the organs undergo massive growth and remodeling.  In addition, the bipotential gonad primordium develops into either testis or ovary.  This protocol identifies many of the organs of the adult and demonstrates methods for dissection of the brain, gonads, gastrointestinal system, heart, and kidney of the adult zebrafish. The dissected organs can be used for in situ hybridization, immunohistochemistry, histology, RNA extraction, protein analysis, and other molecular techniques.  This protocol will assist in the broadening of studies in the zebrafish to include the remodeling of larval organs, the morphogenesis of organs specific to the adult and other investigations of the adult organ systems.

This protocol describes a procedure for identifying and dissecting organs from the adult zebrafish.

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental ...

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 ...

Transplantation of Cells Directly into the Kidney of Adult Zebrafish

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic diseases1. However, most organs are not easily accessible, necessitating the need to develop surgical methods to gain access to these structures. In this video article, we describe a method for transplanting cells directly into the kidney of adult zebrafish, a popular model to study regeneration and disease2. Recipient fish are pre-conditioned by irradiation to suppress the immune rejection of the injected cells3. We demonstrate how the head kidney can be exposed by a lateral incision in the flank of the fish, followed by the injection of cells directly in to the organ. Using fluorescently labeled whole kidney marrow cells comprising a mixed population of renal and hematopoietic precursors, we show that nephron progenitors can engraft and differentiate into new renal tissue - the gold standard of any cell-based regenerative therapy. This technique can be adapted to deliver purified stem or progenitor cells and/or small molecules to the kidney as well as other internal organs and further enhances the zebrafish as a versatile model to study regenerative medicine.

Cell transplantation is an essential technique for studying tissue regeneration and for developing cell-based therapies of disease. We demonstrate here a microsurgical technique that permits the transplantation of genetically labeled cells directly into the kidney of adult zebrafish fish.

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic ...

Dissection of the Adult Zebrafish Kidney

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem cells during normal homeostasis and disease states. The understanding of adult stem cells has broad reaching implications for the future of regenerative medicine1. For example, better knowledge about adult stem cell biology can facilitate the design of therapeutic strategies in which organs are triggered to heal themselves or even the creation of methods for growing organs in vitro that can be transplanted into humans1. The zebrafish has become a powerful animal model for the study of vertebrate cell biology2. There has been extensive documentation and analysis of embryonic development in the zebrafish3. Only recently have scientists sought to document adult anatomy and surgical dissection techniques4, as there has been a progressive movement within the zebrafish community to broaden the applications of this research organism to adult studies. For example, there are expanding interests in using zebrafish to investigate the biology of adult stem cell populations and make sophisticated adult models of diseases such as cancer5. Historically, isolation of the zebrafish adult kidney has been instrumental for studying hematopoiesis, as the kidney is the anatomical location of blood cell production in fish6,7. The kidney is composed of nephron functional units found in arborized arrangements, surrounded by hematopoietic tissue that is dispersed throughout the intervening spaces. The hematopoietic component consists of hematopoietic stem cells (HSCs) and their progeny that inhabit the kidney until they terminally differentiate8. In addition, it is now appreciated that a group of renal stem/progenitor cells (RPCs) also inhabit the zebrafish kidney organ and enable both kidney regeneration and growth, as observed in other fish species9-11. In light of this new discovery, the zebrafish kidney is one organ that houses the location of two exciting opportunities for adult stem cell biology studies. It is clear that many outstanding questions could be well served with this experimental system. To encourage expansion of this field, it is beneficial to document detailed methods of visualizing and then isolating the adult zebrafish kidney organ. This protocol details our procedure for dissection of the adult kidney from both unfixed and fixed animals.
Dissection of the kidney organ can be used to isolate and characterize hematopoietic and renal stem cells and their offspring using established techniques such as histology, fluorescence activated cell sorting (FACS)11,12, expression profiling13,14, and transplantation11,15. 
We hope that dissemination of this protocol will provide researchers with the knowledge to implement broader use of zebrafish studies that ultimately can be translated for human application.

The zebrafish kidney is home to both renal and hematopoietic adult stem/progenitor cells, and represents an outstanding opportunity to study these cell types and their progeny in a vertebrate model organism. Here, we demonstrate a detailed dissection procedure that enables the researcher to identify and surgically remove the adult zebrafish kidney, which can be used for applications such as cell isolation, transplantation, and expression studies of kidney and/or blood cell populations.

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem ...

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 ...

In Ovo Electroporation in Embryonic Chick Retina

Chicken embryonic retina is an excellent tool to study retinal development in higher vertebrates. Because of large size and external development, it is comparatively very easy to manipulate the chick embryonic retina using recombinant DNA/RNA technology. Electroporation of DNA/RNA constructs into the embryonic retina have a great advantage to study gene regulation in retinal stem/progenitor cells during retinal development. Different type of assays such as reporter gene assay, gene over-expression, gene knock down (shRNA) etc. can be performed using the electroporation technique. This video demonstrates targeted retinal injection and in ovo electroporation into the embryonic chick retina at the Hamburger and Hamilton stage 22-23, which is about embryonic day 4 (E4). Here we show a rapid and convenient in ovo electroporation technique whereby a plasmid DNA that expresses green fluorescent protein (GFP) as a marker is directly delivered into the chick embryonic subretinal space and followed by electric pulses to facilitate DNA uptake by retinal stem/progenitor cells. The new method of retinal injection and electroporation at E4 allows the visualization of all retinal cell types, including the late-born neurons1, which has been difficult with the conventional method of injection and electroporation at E1.52.

In Ovo Electroporation in Embryonic Chick Retina

The overall goal of this video is to show how to perform targeted retinal injection and in ovo electroporation of DNA/RNA constructs into the chick embryonic retina at the Hamburger and Hamilton stage 22-23, which is about embryonic day 4 (E4). This technique is very useful to study gene expression, gene regulation, and morphological change in developing chick retina.

Chicken embryonic retina is an excellent tool to study retinal development in higher vertebrates. Because of large size and external development, it is ...