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. Resuspend Morpholino
<ol>
 <li>Dilute 300 nM of fluorescein-tagged morpholino into 100 &mu;l of nuclease-free water to make an approximately 3 mM solution. The morpholino solution is aliquoted into multiple paraffin-sealed microcentrifuge tubes and stored at room temperature and away from light.</li>
 <li>To determine the exact morpholino concentration, dilute 5 &mu;l of morpholino solution (or water as a blank) in 95 &mu;l of 0.1 N HCl. Set a baseline on a spectrophotometer at 265 nm with the water blank and then determine the reading of the diluted morpholino solution. Multiply the morpholino absorbance by its determined constant and by the dilution factor to determine the concentration in ng/&mu;l. The morpholino constant is calculated as: 
Morpholino constant = molecular weight of the morpholino X 1000/molar absorbance. 
 The molecular weight and molar absorbance for the morpholino can be found on the "Oligo Properties" sheet provided with the product. Divide the morpholino concentration, in ng/&mu;l, by the molecular weight to determine the concentration in mM. Dilute the morpholino, if necessary, to a working concentration, typically 1.2 mM.</li>
</ol>
2. Fin Amputation
<ol>
 <li>Anesthetize adult zebrafish in either Tricaine or 2-phenoxyethanol at 1.0 mg/ml in tank water.</li>
 <li>Amputate the fin using a sterile scalpel or razor blade proximal to the first lepidotrichial branching point. This should be done at the same proximal/distal location on each animal (e.g. 7 bony segments distal from the fin girdle). IMPORTANT: make sure to cut perfectly perpendicular to the anterior/posterior plane of the animal. Angled cuts will result in uneven fin outgrowth of the dorsal and ventral halves of the fin.</li>
 <li>Return the fish to a tank. We typically maintain the fish at 33 &deg;C to increase the regeneration rate. Depending on the experimental design, wait 0-2 days post amputation (dpa) for fin regeneration to begin before introducing the morpholino. Alternate approaches are discussed in the Discussion, below.</li>
</ol>
3. Morpholino Injection
<ol>
 <li>The day prior to injecting the morpholino, make an injection plate (Figure 2). Make sure to cut out a notch at one end of the well, which helps stabilize the fish for the microinjection procedure.</li>
 <li>At 2 dpa, prepare the micro-injection apparatus and morpholino solution.</li>
 <li>Dilute the fluorescein-tagged morpholino to the proper concentration (recommend starting with 1.2 mM) and place in a 65 &deg;C water bath, for 5 minutes.</li>
 <li>Pull the glass needle for the micro-injection using a needle puller.</li>
 <li>Load the needle with the morpholino (note: depending on whether you back-fill or front-load your needle will determine whether you load the needle first, or cut the tip first).</li>
 <li>Cut the tip off the needle at an angle.</li>
 <li>Follow the directions of your micro-injection system to inject an approximately 5 nl bubble of morpholino per injection. Larger volumes of morpholino could disrupt the tissue.</li>
 <li>Anesthetize the fish and place on injection plate. Remove all excess liquid and orient the needle to the appropriate location, just distal to the bony ray (Figure 3). Using a microscope and microinjection apparatus appropriate for morpholino injections (see Table of Specific Reagents), inject the morpholino into the fin from anterior to posterior, as shown in Figure 3. Injecting from posterior to anterior causes the fin tissue to roll up during the injection and is very difficult.</li>
 <li>Insert the needle gently into the regenerative tissue, just distal to each bony ray (marked "1" in Figure 3) and push distally until located in the blastema (marked "2" in Figure 3). Be careful not to push the needle through the entire fin. When the needle is correctly localized, inject the morpholino (i.e. click one time). At 1.2 mM, the fluorescein-tagged morpholino solution appears yellow-green, even in normal light conditions. With each injection, a yellow "puff" of morpholino solution can be visualized, which helps localize the injection to the blastema. Approximately 75 nl (10-15 clicks) of morpholino solution should be injected per bony ray. Pause ~ 1 sec between clicks. We only inject the dorsal side, using the ventral as electroporation-only control. Alternatively, one could repeat this entire procedure using a control morpholino on the ventral half of the fin.</li>
</ol>
4. Electroporation of the Morpholino
<ol>
 <li>Immediately following injection of the morpholino, remove the injection plate from the micro-injection apparatus. Fill the injection plate well with the anesthesia solution until the fish is submerged. The electroporation is performed under the water line.</li>
 <li>A 3 mm diameter platinum plate tweezer electrode (CUY 650-P3 Tweezers, Protech International) localizes the pulses to approximately one-half of the fin. Be careful not to touch the electrodes to the fin tissue. To ensure that the electrodes do not touch the fin tissue, rotate the fish on its dorsal side and look straight down the midline for the electroporation.</li>
 <li>Electroporate both the dorsal and ventral (to control for a non-specific electroporation effects) sides of the fin using a CUY21 Square Wave Electroporator (Protech International, Inc.). Wipe the electrodes with a damp KimWipe after each electroporation to remove any bubbles that may have formed. If they are not removed, a mini-charge builds up, which will attract the tissue toward the electrode. The electroporation parameters should be set to ten consecutive 50 msec pulses, at 15 V with a 1 sec pause between pulses.</li>
 <li>Place the fish on a glass slide or Petri dish and quickly image the fin, making sure to note the dorsal/ventral orientation. This image will be used for regrowth analysis on the following day. Return the fish to the tank.</li>
</ol>
5. Analysis
<ol>
 <li>If the targeted protein that was knocked down in expression is required for proper fin regeneration, a dramatic difference between the dorsal and ventral halves of the fin should be evident 1 day post-electroporation (Figure 5B).</li>
 <li>At 3 dpa, take another picture of the fin of each fish and match up this image with the corresponding 2 dpa image (Figure 5B). The natural variation in the fin pigmentation pattern between animals will allow you to correctly identify individual fish.</li>
 <li>Using NIH Image, trace the 2 dpa and 3 dpa areas of both the dorsal and ventral halves (Figure 5B).</li>
 <li>To determine % area of dorsal versus ventral use the following formula: (Dorsal3dpa - Dorsal2dpa)/(Ventral3dpa - Ventral2dpa) X 100 = % area</li>
</ol>
6. Representative Results
<ol>
 <li>We always assay for fin outgrowth at 3 dpa, or 24 hpe (hours post electroporation). At this time, the fluorescein-tagged morpholino should be present in the dorsal half of the fin (Supplemental Figure 1 and Figure 6A). It is normal to see some trailing down to the level of the amputation.</li>
 <li>There should be no difference between the dorsal and ventral sides in the fin injected and electroporated with the control morpholinos (Figure 6B).</li>
 <li>If the experimental morpholino targets an essential protein for fin outgrowth, a dramatic difference between the dorsal and ventral sides should be observed (Figure 6C).</li>
 <li>If the electrode touched the fin during electroporation, the tissue will appear brown (almost burnt in appearance) and necrotic.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/3632/3632fig1.jpg" /> 
Figure 1. Schematic of the various events that occur during fin regeneration. The underlying times for each event are given in hours post-amputation (hpa) and correspond to a tank temperature of 33 &deg;C.
<img alt="Figure 2" src="/files/ftp_upload/3632/3632fig2.jpg" /> 
Figure 2. A. Schematic of the injection plate, which is made from agarose and contains a small well to hold the fish during microinjection of the morpholino.
<img alt="Figure 3" src="/files/ftp_upload/3632/3632fig3.jpg" /> 
Figure 3. Schematic of morpholino microinjection. A. Place the fish in the dish with the head of the fish in the notch cut out of the well, which will help the fish stay stable. B. At low magnification, arrange the needle so that it is close to the regenerating tissue of the fin. C. At higher magnification, inject the morpholino distal to each bony fin ray (i.e. in each blastema). The needle should enter the tissue just distal to the bony ray (1), and then continue to the location of the blastema (2). Note: the green circles in the schematic are only meant to show the location of the injection. The morpholino can briefly be visualized as a green/yellow "puff" following each injection; however, this does not persist as shown in the schematic.
<img alt="Figure 4" src="/files/ftp_upload/3632/3632fig4.jpg" /> 
Figure 4. Schematic of fin electroporation. A. Following microinjection, place the fish in a Petri dish full of anesthesia and electroporate both the dorsal and ventral halves. B. Make sure to not touch the fin tissue. Electrodes should be placed ~ 1 mm from the tissue.
<img alt="Figure 5" src="/files/ftp_upload/3632/3632fig5.jpg" /> 
Figure 5. Schematic of the methods used to calculate fin outgrowth inhibition. A. Take a picture of the fin of each fish at 2 dpa, either immediately before or after morpholino injection and electroporation. Trace the regenerative tissue of both the dorsal (green) and ventral (blue) halves of the fin using NIH Image, (black dashed lines). B. At 3 dpa, take another picture of each fin and again trace the dorsal and ventral areas of regrowth using NIH Image. C. Subtract the area of regrowth at 2 dpa from the total regrowth at 3 dpa for both the dorsal and ventral haves. The percent area of dorsal versus ventral re-growth can be calculated using the formula: ((D3dpa - D2 dpa)/(V3dpa - V2dpa)) X 100. Percent inhibition = 100 - Percent Area.
<img alt="Figure 6" src="/files/ftp_upload/3632/3632fig6.jpg" /> 
Figure 6. Examples of expected outcomes. A. Fluorescent image showing a fluorescein-tagged control morpholino in the dorsal half of the fin, 24 hours post-electroporation (hpe). B. Brightfield image of a fin that was injected and electroporated with a control morpholino in the dorsal half. The image shows equal regrowth of both the dorsal and ventral halves of a fin, 24 hpe. C. Brightfield image of a fin that was injected and electroporated with an experimental morpholino in the dorsal half. The image shows inhibition of regrowth on the dorsal/injected side. The line shows the amount of regrowth at 2 dpa, immediately prior to morpholino injection and electroporation.
<img alt="Figure 7" src="/files/ftp_upload/3632/3632fig7.jpg" /> 
Figure 7. Schematic of an alternate injection and electroporation procedure to target proteins involved in wound healing and blastema formation. A. Inject the morpholino between each bony fin ray on the dorsal half of the fin. B. Electroporate the morpholino as per normal. C. Amputate the fin immediately proximal (~ 1 bony segment) to the injection site. D. The fluorescein-tagged morpholino can be observed in wound epithelium and blastema at 24 hpe.
<img alt="Figure 8" src="/files/ftp_upload/3632/3632fig8.jpg" /> 
Figure 8. Using the technique to target wound epithelium and blastema formation. A. Brightfield image of a fin at 24 hpa that was injected and electroporated with a control morpholino immediately prior to amputation. B. Fluorescent image of fin shown in panel A. Note that the injected dorsal half of the fin shows good uptake of the morpholino in the regenerative tissue. C - C". Higher magnification of the dorsal half of the fin shown in panels A and B. The injection sites are often still visible (arrowheads), but many targeted cells have migrated to participate in wound epithelium and blastema formation (arrow).
<img alt="Figure 9" src="/files/ftp_upload/3632/3632fig9.jpg" /> 
Figure 9. Using the technique to target cells that migrate to form the blastema. A. Fluorescent and brightfield inset images showing a fin injected and electroporated with morpholino on both the dorsal and ventral halves. The fin was then amputated at two planes. The dorsal half was cut immediately distal to the injection sites, where as the ventral half was cut 9-10 bony segments distal to the injection sites. B. At 24 hpa, the fluorescent and brightfield inset images show that morpholino has migrated to the dorsal regenerate (1), but not the ventral regenerate (2), indicating that only the cell immediately proximal to the cut site participate in blastema formation. The two sets of white arrowheads show the level of each amputation plane. Panels marked 1 and 2 on the far right of the image show a higher magnification view of the dorsal and ventral halves of the fin, respectively.
<a href="https://www.jove.com/files/ftp_upload/3632/Suppl_Fig1_JoVE.mov">Supplemental Figure 1.</a> A video of a confocal z-stack of a region corresponding to the location of a blastema in a fin injected and electroporated with a control morpholino. The image was taken at 24 hpe. Since a single fluorescein molecule cannot be visualized, not all of the morpholino can be visualized or quantified. However, these images give some idea of the varying degrees of uptake that can be visualized in cells, from individual punctate dots, to entire cells full of fluorescent morpholino. On the still image, the orientation is shown. Scale bar: 25 microns.

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We have nothing to disclose.

Here, we describe a powerful loss-of-function approach to conditionally knockdown proteins of interest during fin regeneration in adult zebrafish. This technique has been used to study gap junction genes, signaling receptors, transcription factors, and microRNAs during regenerative fin outgrowth16, 20-22.
We anticipate that this technique could also be used to study genes required for wound healing and blastema formation by adapting the technique. For example, we injected and electroporated a control morpholino in the space between the bony fin rays on the dorsal half of the fin prior to amputation (Figure 7). We then amputated the fin immediately distal to the injection plane. 24 hpa, we observed that the morpholino-targeted cells had migrated distally to form both the wound epithelium and blastema (Figure 8), indicating that cells during these early stages of regeneration can also be targeted. 
The technique has a few notable limitations. For example, fluorescence from the fluorescein tag does not persist following fixation and processing for immunohistochemistry, which makes it impossible to correlate a particular cellular phenotype (i.e. cell proliferation) with the amount of morpholino present in a cell. In addition, we have been unable to consistently achieve the electroporation of plasmids into the regeneration tail fin, although a previous group did report the successful electroporation of DNA into fin tissue23. Finally, we have noted that the morpholino is only effective for ~48 hours post electroporation21, which prohibits using this technique in its current form for testing genes involved in the differentiation of new cell types. Additional testing and modification of the procedure may overcome these current limitations.
In addition, it is possible that this technique could be used to test proteins involved in cell migration from the underlying tissue to the blastema. For example, we injected and electroporated a control morpholino into both sides of the fin (as described in Figure 7) prior to amputation. We then amputated the dorsal half of the fin immediately distal to the injection plane and we amputated the ventral fin much more distally. At 24 hpa, the morpholino-positive cells on the dorsal side had migrated from the injection site to the overlying wound epithelium and blastema. However, that was not the case on the ventral side (Figure 9). This supports the idea that only the cells that underlie the amputation plane participate in the regenerative response. These data also suggest that proteins hypothesized to be required for cell migration could be tested using this technique.

<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></td>
 </tr>
 <tr>
 <td>Morpholino</td>
 <td>GeneTools, LLC</td>
 <td></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>Micro-injection pump</td>
 <td>World Precision Instruments</td>
 <td>PV830 Pneumatic PicoPump</td>
 <td>Many different microinjection systems could be used</td>
 </tr>
 <tr>
 <td>Micro-manipulator</td>
 <td>World Precision Instruments</td>
 <td>MMJR</td>
 <td>Right-handed (MMJL for left-handed)</td>
 </tr>
 <tr>
 <td>Micro-injection needles, 1.0 mm outside diameter</td>
 <td>World Precision Instruments</td>
 <td>1B100F-4</td>
 <td>These are borosilicate glass capillaries, pulled into a needle</td>
 </tr>
 <tr>
 <td>Needle holder</td>
 <td>World Precision Instruments</td>
 <td>5430-ALL</td>
 <td>Pico Nozzle Kit; make sure to inset the 1.0 mm pipette gasket</td>
 </tr>
 <tr>
 <td>Needle puller</td>
 <td>Sutter</td>
 <td>P-97</td>
 <td>Other micropipette/needle pullers should also work </td>
 </tr>
 <tr>
 <td>Microscope</td>
 <td>Leica, Nikon, Zeiss</td>
 <td>Number varies depending on the manufacturer</td>
 <td>Any stereomicroscope with 20X optics and the ability to work with micromanipulators</td>
 </tr>
 </tbody>
</table>

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.

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

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.

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 ...

in-vivo-electroporation-morpholinos-into-regenerating-adult-zebrafish

Research

JoVE Journal

Biology

16.8K Aufrufe.  University of Notre Dame. Wir beschreiben ein Verfahren zur bedingten Knockdown die Expression eines Zielproteins im Erwachsenenalter Zebrafisch Flossenregeneration. Diese Technik beinhaltet Mikrospritzvorrichtung und Elektroporation Antisense-Oligonukleotid in Morpholinos Flossengewebes das Testen des Proteins Rolle in verschiedenen Stadien der Flossenregeneration, einschließlich Wundheilung, Blastembildung und regenerative Auswachsen ermöglicht.

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

Transplantation of Whole Kidney Marrow in Adult Zebrafish

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an organism. The functional definition of a HSC is a transplanted cell that has the ability to reconstitute all the blood lineages of an irradiated recipient long term. This designation was established by decades of seminal work in mammalian systems. Using hematopoietic cell transplantation (HCT) and reverse genetic manipulations in the mouse, the underlying regulatory factors of HSC biology are beginning to be unveiled, but are still largely under-explored. Recently, the zebrafish has emerged as a powerful genetic model to study vertebrate hematopoiesis. Establishing HCT in zebrafish will allow scientists to utilize the large-scale genetic and chemical screening methodologies available in zebrafish to reveal novel mechanisms underlying HSC regulation. In this article, we demonstrate a method to perform HCT in adult zebrafish. We show the dissection and preparation of zebrafish whole kidney marrow, the site of adult hematopoiesis in the zebrafish, and the introduction of these donor cells into the circulation of irradiated recipient fish via intracardiac injection. Additionally, we describe the post-transplant care of fish in an "ICU" to increase their long-term health. In general, gentle care of the fish before, during, and after the transplant is critical to increase the number of fish that will survive more than one month following the procedure, which is essential for assessment of long term (&lt;3 month) engraftment. The experimental data used to establish this protocol will be published elsewhere. The establishment of this protocol will allow for the merger of large-scale zebrafish genetics and transplant biology.

In this article, we demonstrate a method to perform HCT in adult zebrafish.

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an ...

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 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 ...

Gavaging Adult Zebrafish

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or agents in solutions for scientific research. Current methods for administering compounds orally to adult zebrafish are inaccurate due to variability in voluntary consumption by the fish. A gavage procedure was developed to deliver precise quantities of infectious agents to zebrafish for study in biomedical research. Adult zebrafish over 6 months of age were anesthetized with 150 mg/L of buffered MS-222 and gavaged with 5 &mu;l of solution using flexible catheter implantation tubing attached to a cut 22-G needle tip. The flexible tubing was lowered into the oral cavity of the zebrafish until the tip of the tubing extended past the gills (approximately 1 cm). The solution was then injected slowly into the intestinal tract. This method was effective 88% of the time, with fish recovering uneventfully. This procedure is also efficient as one person can gavage 20-30 fish in one hour. This method can be used to precisely administer agents for infectious diseases studies, or studies of other compounds in adult zebrafish.

The increasing use of zebrafish as an animal model requires the development of effective methods for the delivery of known quantities of compounds and solutions. The gavage procedure described below allows for the oral delivery of precise volumes of solution reliably, safely and efficiently to adult zebrafish.

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or ...