Name: intraperitoneal injection into adult zebrafish
Uploaded: 2010-08-30T17:00:00
Duration: 9 min 19 s
Description: Watch this Scientific Journal Video about Intraperitoneal Injection into Adult Zebrafish at JoVE.com

This study was supported by Juvenile Diabetes Research Foundation grant 5-2007-97 (to V.E.P.), by National Institute of Diabetes and Digestive and Kidney Diseases grants R01DK064973 (to V.E.P.), R01DK48494 (to L.H.P.), T32DK07074 (supporting S.C.E.), K01DK083552 (to M.D.K), and by P60DK20595 to The University of Chicago Diabetes Research and Training Center. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIDDK or the NIH.

1. Pre-injection Preparations
<ol>
 <li> Fast the fish for at least 24 hours prior to injection. This will empty the intestinal bulb (stomach) contents. The basic fasting protocol is to transfer the fish, at their normal density, to a clean tank, then withhold food. For longer-term fasting that requires more rigorous conditions (e.g., for blood glucose studies), see additional considerations in the Discussion.</li>
 <li> Prepare Cortland salt solution (Perry et al., 1984). 
 For a 100 mL volume, dissolve the following in distilled water: 
 725 mg NaCl (124.1 mM) 
 38 mg KCl (5.1 mM) 
 41 mg Na2HPO4 (2.9 mM) 
 24 mg MgSO4&#8729;7H2O (1.9 mM) 
 16 mg CaCl2&#8729;2H2O (1.4 mM) 
 100 mg NaHCO3 (11.9 mM) 
 4 g Polyvinylpyrrolidone (PVP) (4%) 
 1,000 USP units Heparin 
 Filter, sterilize and store at 4&deg;C.</li>
 <li> Prepare the microscope.
 <ul>
 <li> Cover the microscope base with plastic wrap for protection in case of spills.</li>
 <li> Put a paper towel on top of the plastic wrap. The surgical table will sit on top of the paper towel.</li>
 <li> Pre-adjust focus by viewing the surgical table and focusing on the sponge.</li>
 </ul>
 Tip: Put your finger on top of the sponge and focus on that. This will eliminate or minimize further focal adjustment once the fish is on the surgical table.</li>
 <li> Weigh the fish.
 <ul>
 <li> Fill a 500 mL beaker about 1/3 full with fish facility water.</li>
 <li> Tare the balance.</li>
 <li> Collect the fish using a net. Wick excess water away from the net and fish by briefly dabbing the net on paper towels. Transfer the fish to the beaker.</li>
 <li> Weigh the fish.</li>
 <li> Transfer the fish to a clean tank.</li>
 <li> Transfer each weighed fish to its own labeled tank.</li>
 <li> Calculate the injection volume for each fish based on fish weight.</li>
 </ul>
 </li>
 <li> Prepare the syringe and related injection equipment. For injection, we recommend a 35G beveled steel needle and a 10 &mu;l NanoFil microsyringe. Prepare the NanoFil syringe and silflex tubing following the manufacturer's instructions. It is important to remove any bubbles from the syringe and tubing. After filling the syringe and tubing, mount the syringe on the pump, and program the injection volume for the first fish.</li>
 <li> Prepare the surgical table.
 <ul>
 <li> Cut a soft sponge (such as #L800-D, Jaece Industries) so that it is approximately 20 mm in height. On the flat face, make a cut that is 10-15 mm deep. This cut is the trough that will hold the fish for injection.</li>
 <li> Set the sponge into a 60 mm Petri dish.</li>
 <li> Set the Petri dish with sponge into a suitably-sized pipette tip box lid. The lid needs to be large enough to hold water to help maintain sponge temperature, but it should be shallow enough to not get in the way. We use a lid from a P200 tip box that is 11.4 cm L x 7.7 cm W x 1.5 cm D.</li>
 </ul>
 These three items assembled together (sponge in petri dish in box lid) constitute the surgical table.</li>
 <li> Prepare the anesthetic.
 <ul>
 <li> Make crushed ice using cubes made from fish facility water.</li>
 </ul>
 Tip: Using typical ice cube trays, it will take 3 trays to anesthetize 10-12 fish.
 <ul>
 <li> Fill a clean ice bucket with the crushed ice.</li>
 <li> Put the surgical table into a larger container such as a 2.4 liter Rubbermaid food storage container.</li>
 <li> Pour some facility water (warm) into the outer container and the surgical table. Keep a reserve of warm facility water nearby.</li>
 <li> Put a thermometer into the outer container.</li>
 </ul>
 </li>
</ol>
2. Anesthesia, Injection and Recovery
<ol>
 <li> Place the anesthetic outer container plus surgical table adjacent to the microscope. Have the bucket of ice chips nearby.</li>
 <li> Bring the water temperature down to 17&deg;C by adding ice chips. Important: Don't go below 17&deg;C for this step.</li>
 <li> Use a net to transfer the fish to the outer container.</li>
 <li> Slowly add ice chips to the container to bring the temperature down to 12&deg;C, over the course of several minutes.</li>
 <li> Monitor fish behavior: At 17&deg;C or slightly lower, the fish typically will spread its pectoral fins horizontally, gasp, and have rapid operculum movements. As the temperature drops, the fish will swim more slowly and finally stop swimming. As the surgical plane of anesthesia is approached, gasping will stop and operculum movements will slow. The fish is ready for injection when it does not react to being handled. For most fish, 12&deg;C is sufficient. Larger fish may require colder water.</li>
 <li> As the required temperature is reached (~12&deg;C or colder), press on the sponge to saturate it.</li>
 <li> Keep your fingers in the cold water sufficiently so that they will not warm up the fish and bring it out of anesthesia during handling.</li>
 <li> With cold fingers, gently transfer the fish to the trough of the sponge. Position the fish with the abdomen up and the gills in the trough.</li>
 <li> Quickly transfer the surgical table to the microscope stage.</li>
 <li> Working quickly, carefully insert the needle into the midline between the pelvic fins. The needle should point cranially and be inserted closer to the pelvic girdle than to the anus. You should be able to feel when the needle is deep to the body wall. Inject the appropriate volume and withdraw the needle.</li>
 <li> After injection, immediately transfer the fish back to its warm-water (~28.5&deg;C) tank for recovery by releasing the fish from the sponge over the tank water. 
 Tip: If the fish does not begin swimming immediately, help it to recover by gently swirling water towards the gills.</li>
 <li> Check the needle. Occasionally a scale may be attached and should be removed prior to the next injection.</li>
 <li> For subsequent injections, use warm facility water to bring the anesthetic chamber water temperature back up to 17&deg;C before introducing the next fish.</li>
</ol>
3. Representative Results:
<img src="/files/ftp_upload/2126/2126fig1.jpg" alt="Figure 1" /> 
 Figure 1. Representative results following intraperitoneal injection of 0.5 mg/g glucose or vehicle. Fish were fasted for 72 hours prior to injection. The x-axis shows time, post-injection. Mean &plusmn; SEM.

<ol>
	<li>Perry, S. F., Davie, P. S., Daxboeck, C., Ellis, A. G., Smith, D. G., Hoar, W. S., Randall, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion+methods+for+the+study+of+gill+physiology.">Perfusion methods for the study of gill physiology.</a> Fish Physiology Volume X: Gills, Part B: Ion and Water. , 325-388 (1984).</li><li>Brown, L. A., Stoskopf, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthesia+and+restraint.">Anesthesia and restraint.</a> Fish Medicine. , 79-90 (1993).</li><li>Eames, S. C., Philipson, L., Prince, V. E., Kinkel, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+sugar+measurement+in+zebrafish+reveals+dynamics+of+glucose+homeostastis.">Blood sugar measurement in zebrafish reveals dynamics of glucose homeostastis.</a> Zebrafish. 7, 205-213 (2010).</li><li>Chavin, W., Young, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Factors+in+the+determination+of+normal+serum+glucose+levels+of+goldfish+Carassius+auratus+L.">Factors in the determination of normal serum glucose levels of goldfish Carassius auratus L.</a> Comp Biochem Physiol. 33, 629-653 (1970).</li><li>Groff, J. M., Zinkl, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hematology+and+clinical+chemistry+of+cyprinid+fish.+Common+carp+and+goldfish.">Hematology and clinical chemistry of cyprinid fish. Common carp and goldfish.</a> Vet Clin North Am Exot Anim Pract. 2, 741-776 (1999).</li><li>Brand, M., Granato, M., Nusslein-Volhard, C., Nusslein-Volhard, C., Dahm, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Keeping+and+raising+zebrafish.">Keeping and raising zebrafish.</a> Zebrafish: A Practical Approach. , 7-37 (2002).</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. , (1995).</li><li>Iwama, G. K., Ackerman, P. A., Hochachka, P. W., Mommsen, T. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anaesthetics.">Anaesthetics.</a> Biochemistry and Molecular Biology of Fishes, Volume 3: Analytical Techniques. , 1-15 (1994).</li><li>Reavill, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Common+diagnostic+and+clinical+techniques+for+fish.">Common diagnostic and clinical techniques for fish.</a> Vet Clin North Am Exot Anim Pract. 9, 223-235 (2006).</li><li>Stoskopf, M. K., Stoskopf, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surgery.">Surgery.</a> Fish Medicine. , 91-97 (1993).</li></ol>

No conflicts of interest declared.

Intraperitoneal injection involves five steps: fasting, weighing, anesthetizing, injection, and recovery. For each step there are best practices that can ensure success. Success includes a healthy fish patient as well as a good experimental outcome.

Fasting: A 24-hour fast should empty the intestinal bulb. This practice is taken from the fish veterinary literature (e.g., Brown 1993). Additional fasting considerations are discussed below.

Longer-term fasting: We have found that a 72-hour fast is required to decrease blood glucose to a baseline level prior to injection (Eames et al., 2010). We have also found that for glucose studies there are several procedures that are required to ensure that the fish are fasted properly. Start with a clean tank (no debris on the bottom). Tanks should be offline, clearly labeled as 'fasting', and in a location where enthusiastic fish care personnel will not feed them. Evaluate the external environment of the tank and take steps to prevent the fish from being stressed from disturbances, as stress is known to raise blood glucose (Chavin and Young, 1970; Groff et al., 1999). For example, we had a fasting experiment in which a radio was operated daily on the bench that was holding the fish tanks. We found that blood glucose was unusually high and concluded that the fish were stressed by the vibrations. Another stressor is overcrowding. Fish should be kept at a density that conforms with good fish husbandry practices. For recommendations, see Brand et al. (2002) and Westerfield (1995). We have had good results fasting our fish at a density of 10-12 fish in a 9 liter tank (with 3 layers of marbles taking up some of that volume). Separating the sexes may cause stress, so we recommend maintaining a mixed-sex population during the fast. This means that eggs can be laid, and the eggs need to be sequestered so that they will not be eaten. A simple way to sequester eggs is to cover the tank bottom with 2-3 layers of marbles. Water quality needs to be maintained by removing eggs and waste and by replacing about 10-15% of the tank water, daily. For removing eggs and waste, siphoning works well.

Weighing: When weighing fish that are not anesthetized, care should be taken to minimize water transfer from the net into the beaker, to ensure accurate weighing. If the net (with fish) is blotted on paper towels, the majority of the excess water can be removed, and the weight can be accurately measured. It may be easier to anesthetize the fish prior to weighing, but we have not tested the possible effects of anesthetizing a fish twice in one day. We have tested our technique by weighing the fish first with the netting/blotting method and then re-weighing the fish after it has been anesthetized, and gently blotted dry. We found no significant difference in weight between the methods (P = 0.7927, t-test). Additionally, we tested whether this netting/blotting method affected blood glucose, in comparison with simply transferring the fish to the beaker as soon as it is netted (no blotting). We found no significant difference in blood glucose level between the two transfer methods (P = 0.2241, t-test).

Anesthetizing: Chemical anesthesia may be suitable for many studies. Here we have demonstrated cold water anesthesia as an alternative, because many anesthetics (including tricaine/MS-222 (Brown, 1993)), raise blood glucose. In previous studies, we have determined that cold water does not raise blood glucose in zebrafish (Eames et al., 2010). 
For cold water anesthesia, the temperature should be decreased slowly. The rate of decrease seems to depend on the size of the fish, with smaller fish going under faster than larger fish. Following injection, you may observe that the fish is recovering too slowly from the anesthetic (see below). This can result when either the starting temperature is too low, or when the temperature is decreased too rapidly. The starting temperature is too low if the fish bends laterally upon entering the water. If the starting temperature is correct, the fish will maintain its balance initially. It will rotate its pectoral fins to a horizontal position, gasp, and have rapid operculum movements. Typically, it will swim. As temperature decreases, movements will decrease and the fish will lose equilibrium. A surgical plane of anesthesia is reached when the fish can be handled without reacting. To maintain the fish under surgical anesthesia, your fingers must be cold, so keep them in the water prior to handling the fish. The sponge must also be kept cold at the same temperature as the water used for anesthetizing the fish. It is important to saturate the sponge with water that is sufficiently cold to maintain anesthesia once the fish is placed onto it.

Injection: Prior to undertaking injections, you may want to dissect at least one fish to get a sense of body wall thickness. This can help you to judge how far the needle needs to insert to enter the abdominal cavity. Additionally, as you insert the needle, you can feel the body wall "give" when the needle enters the abdominal cavity. During the injection, take steps to keep the patient happy. Make sure the sponge is saturated with the correct temperature cold water to prevent the fish from reviving during injection. A well-saturated and soft sponge is important for minimizing damage to the scales and mucus covering of the skin. A well-saturated sponge is also important for keeping the gills aerated. We highly recommend the foam sponge listed below under Materials. Finally, once the fish is anesthetized, work quickly to minimize the time that the fish is under.

Recovery: The fish should recover from the anesthesia virtually upon entering the warm tank water. If the fish does not begin swimming immediately, gently swirl the water towards its gills to speed recovery. If recovery is slow, then the fish went under too quickly and you should adjust the anesthesia procedure appropriately. The possible causes of slow recovery are discussed under Anesthetizing.

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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>Foam Sponge</td>
<td>&nbsp;</td>
<td>Jaece Industries</td>
<td>L800-D</td>
<td>&nbsp;</td>
<tr>
<td>60 mm Petri dish</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Pipet tip box lid</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>not too deep, e.g. 1.5 cm</td>
</tr>
<tr>
<td>Plastic storage container</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>deep, e.g. 7 cm </td>
</tr>
<tr>
<td>Thermometer</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Crushed ice</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>made from facility water</td>
</tr>
<tr>
<td>Warm facility water</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>1 liter or more</td>
</tr>
<tr>
<td>500 ml beaker</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>for weighing</td>
</tr>
<tr>
<td>NanoFil syringe</td>
<td>&nbsp;</td>
<td>World Precision Instruments (WPI)</td>
<td>NANOFIL</td>
<td>or Hamilton syringe</td>
</tr>
<tr>
<td>35 gauge needle</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>NF35BV-2</td>
<td>beveled</td>
</tr>
<tr>
<td>Silflex tubing</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>SILFLEX-2</td>
<td>&nbsp;</td>
<tr>
<td>UltraMicroPump III and Micro4 controller</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>UMPS-1</td>
<td>&nbsp;</td>
<tr>
<td>Foot switch</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>15867</td>
<td>&nbsp;</td>
<tr>
<td>Dissecting microscope</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Plastic wrap</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Paper towels</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Cortland salt solution</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

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.

Watch this Scientific Journal Video about Intraperitoneal Injection into Adult Zebrafish at JoVE.com

Intraperitoneal Injection into Adult Zebrafish

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, ...

intraperitoneal-injection-into-adult-zebrafish

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Transplantation of Whole Kidney Marrow in Adult Zebrafish

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

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Injection of dsRNA into Female A. aegypti Mosquitos

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video protocol illustrates a method used by the James lab to inject dsRNA into female A. aegypti mosquitoes, which harbor the dengue virus.  The technique for calibrating injection needles, manipulating the injection setup, and injecting dsRNA into the thorax is illustrated.

Reverse genetic approaches have proven extremely useful for determining which genes underly resistance to vector pathogens in mosquitoes. This video protocol illustrates a method used by the James lab to inject dsRNA into female A. aegypti mosquitoes, which harbor the dengue virus. The technique for calibrating injection needles, manipulating the injection setup, and injecting dsRNA into the thorax is illustrated.

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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).

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 ...

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 ...

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 ...

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 ...