This work was funded by the US National Institutes of Health (NIH) grants R01DK069403 and P30DK079307.

Part of the following protocol is based on published methods reported by (Hentschel et al. 2005) and (Weinstein et al. 1995) with slight modifications.
Part 1: Microinjections
Set up in advance for microinjection
<ul>
	<li>The fish are injected at 2 days post-fertilization (dpf). Therefore, adult zebrafish are bred 3 nights prior to a planned experiment. Collect embryos and place them in a Petri dish with E3 water (5mM NaCl, 0.17mM KCL, 0.33mM CaCl2, 0.33mM MgSO4 ) at 28.5&#176;C. At the end of the first day remove dead embryos from the Petri dish and separate embryos so that there are 80-100 embryos per Petri dish.</li>
	<li>Prepare the glass needles pulling from capillary tubes (4 inches in length, 0.63/ 0.20 OD/ID millimeter (mm) R-6 Custom Glass Tubing Drummond scientific company, Broomall, PA 9-000-3000) with an electrode puller. The tip diameter should be of 10-20 &#956;m. Under a dissecting microscope (Leica S6E), with a permanent marker draw approximately 10 1mm lines along the needle, with the help of a ruler, and store the needles in a large Petri dish on clay ramps.</li>
	<li>Prepare the holding pipette: Fire-polish the tip of a thin walled borosilicate glass capillary tube (152 mm, 1 / 0.75 OD/ID mm TW100-6, World Precision Instruments, Inc.) in a Bunsen burner flame until the inner diameter of the tip is 0.4-0.5 mm. Such dimension allows for good grip of the yolk sac of a 2 dpf zebrafish larvae. Hold one end of a piece of glass capillary using long forceps, and heat the opposite tip with of the capillary with the Bunsen burner, placing it vertically over the light blue inner cone of the flame, the hottest part, for 1-2 seconds, rolling it to obtain an even melt. Repeat this a few times then examine the capillary under the microscope, on a stage micrometer (Ward&#39;s 94 W 9910). This is a critical step, it might be necessary to fire-polish a few holding pipettes and select the appropriate size later, while you are manipulating the larvae. If you are careful, one holding pipette lasts for multiple microinjection sessions.</li>
	<li>Fill the Manual Microsyringe Pump (MMP, World Precision Instruments, Inc.,) with mineral oil, following the instructions given by the manufacturer. The pipette holder is inserted in a joystick manipulator (Narishige, MN-151), connected with a magnetic stand (Narishige, MN 151) to an iron plate (Narishige, IP).</li>
</ul>
Microinjection
<ul>
	<li>Prepare 20&#956;l of the injection mixture: 2.4&#956;l gentamicin (Sigma-Aldrich G8648, 50mg/ml in saline solution) and 17.6&#956;l 10-kDa lucifer yellow dextran (Invitrogen D1825, (1mg/ml in saline solution). In the control fish the gentamicin is replaced with saline solution. The fluorescent dextran (many different colors can be used) is sufficient to verify the accuracy of the injection and select larvae that have been injected with the nephrotoxicant.</li>
	<li>Prepare 2 30mm Petri dishes with 2ml mineral oil for placing the injection mixture and control mixture under, to prevent evaporation of the solutions.</li>
	<li>Prepare the anesthetic: 400 mg tricaine powder (ethyl-m-aminobenzoate methanesulfonate SIGMA A-5040) 97.7 ml DD water, 2.1 ml 1M Tris (pH 9). Adjust to pH 7. Store this stock solution in the freezer, in aliquots of 4.2 ml. To use tricaine as an anesthetic dilute the 4.2 ml tricaine stock solution in 100 ml E3 water (Westerfield 1993) and leave at room temperature.</li>
</ul>
The following steps are performed under a dissecting microscope
<ul>
	<li>Manually remove any chorions remaining with fine forceps and anesthetize the zebrafish transferring them in a Petri dish with tricaine warmed at room temperature. Zebrafish should be exposed to tricaine at least 15 minutes before starting the microinjections.</li>
	<li>Open the tip of the injection needle using a razor blade (VWR scientific 55411-050). Hold the blade at a slight angle to create a bevel and cut the tip such as to obtain a tip opening of about 10-20&#956;m diameter.</li>
	<li>Turn on power, vacuum and air supply of the microinjection apparatus (World precision instruments Inc., PV820 pneumatic picopump). Insert the needle into the needle holder mounted on the micromanipulator and tilting base (Word Precision Instruments, Inc., Model M3301R, TB-1).</li>
	<li>To fill the needle, place a droplet (10&#956;l) of the solution to be injected in the 30mm Petri dish filled with mineral oil (Fisher Chemicals, 0121-1). Turn the manual micromanipulator so as to submerge the tip of the needle in the drop of solution, set the picopump &#34;hold/vent&#34; switch to &#34;vent&#34; (&#34;vent&#34; equates to vacuum on the PV820), and fill the needle by pulling solution in through the needle tip. This will take a few minutes, if the solution gets into the needle too fast, that means that the diameter of the tip is too big, and the injections won&#39;t be accurate. After the needle is filled, set the picopump to &#34;hold&#34; (&#34;hold&#34; equates to inject on the PV820).</li>
	<li>To calibrate the needle, set the picopump &#34;gated/timed&#34; switch to &#34;gated&#34;, press the foot pedal to discharge the needle, and record the time it takes (seconds) for the meniscus to travel from one marking to the next (1 mm total distance). Repeat 3 times, take the average time (in seconds), and divide the value by 30 (volume in nanoliters (nl) corresponding to 1 mm linear distance) to determine the number of milliseconds required to deliver 1 nl of solution. Adjust the range period knob to this number, and move the &#34;gated/timed&#34; switch to &#39;timed&#39;. Now the picopump is set to deliver a 1nl pulse of solution.</li>
	<li>Remove the mineral oil dish and position the Petri dish with the larvae under the microscope. Group the fish in the centre of the Petri dish and focus.</li>
	<li>Place the holding pipette in the manual microsyringe pump (MMP, World precision instruments) on the opposite side of the microinjector, advance its tip to the Petri dish, and into the plane of focus of the microscope. Bend it with a fairly shallow angle, so that the very tip of the holding pipette is touching the bottom of the Petri dish and is close to horizontal. Make sure not to have any air bubble in the system, which would slow the response time and decrease the precision of the MMP.</li>
	<li>Change to a higher magnification on the microscope and use a superfine eyelash with handle (Ted Pella 113) to position the larval fish dorsal side down with the yolk very close to the holding pipette. Hold the larval fish by sucking the yolk into the holding pipette by turning counterclockwise the micrometer drive controlling the manual microsyringe pump. Take care not to turn the micrometer too far, as the yolk can burst if pulled too deep into the holding pipette. Focus on the forming common cardinal vein, which lies over the yolk, just beneath the periderm and is the site of very high blood flow. With the micromanipulator of the injection apparatus, guide the needle into the developing common cardinal vein. It is typically necessary to pull the needle back a bit after insertion, as the common cardinal vein is very superficial. Inject 1nl of solution by pressing the foot pedal, then release the larvae from the holding pipette and return it to embryo water. This is a critical step, especially the first time, because it requires a certain amount of practice in positioning the fish and controlling the suction and pressure of the rig. If the yolk of the embryo is properly held with the holding pipette, it won&#39;t roll as the needle enters.</li>
	<li>To verify the success of the microinjection, monitor the presence of the lucifer yellow dextran in the heart of the zebrafish under a fluorescent microscope. The lucifer yellow remains visible in the heart for approximately 20 minutes before dissipating into the circulatory system If the nephrotoxic solution is injected correctly, within 2 hours of the injection you will see very little lucifer yellow dextran remaining at the site of injection. If the solution is injected in the yolk sac, a spot of lucifer yellow dextran will remain at the site of injection. (Fig. 1). Return the fish to E3 medium.</li>
	<li>Two days post-injection, the larvae develop pericardial edema as a consequence of decreased kidney function (Fig. 2).</li>
</ul>
Part 2: Fixation and immunofluorescence with Na+/K+ ATPase antibody (&#945;-6F)
<ul>
	<li>At 4 dpf, put between 10 and 15 larval zebrafish in 4ml glass vials (Weathon, 225012) and fix them in 1ml Dent&#39;s (80% MeOH 20% DMSO) for 4 hours at RT.</li>
	<li>Rehydrate the embryos in a graded MeOH/PBT (PBS with 0.5% Tween20) series, 75:25 50:50, 25:75, 100% PBT, pipetting the solution out and adding new solution to the same vial (1ml of solution per vial), 20 minutes in each solution. Larvae should never be exposed to air, leave a little solution covering the larvae when changing solutions.</li>
	<li>Remove PBT and replace 1ml with blocking solution (PBS with 1% DMSO, 0.5% Tween20, 10% normal goat serum) for 3 hours at 4 &#176;C on a nutator.</li>
	<li>Transfer the larvae in a 2ml microfuge. Remove blocking serum and replace with 300&#956;l primary antibody in incubation solution (PBS with 1% DMSO, 0.5% Tween20, 2% Normal goat serum) overnight at 4&#176;C on a nutator. We use a monoclonal Na+/K+ ATPase antibody (&#945;-6F, supernatant, Developmental Studies Hybridoma Bank, Iowa) at 1:25 dilution</li>
	<li>Remove the primary antibody and wash 3 x 30 minutes with incubation solution at RT.</li>
	<li>Incubate with secondary antibody (goat anti-mouse cy3, Jackson Immunoresearch Laboratories, Inc. 155-165-003) diluted 1:500 in incubation solution for 2 hours at RT.</li>
	<li>Wash 3 x 30 minutes with PBT.</li>
</ul>
Part 3: Plastic embedding with JB-4
<ul>
	<li>Dehydrate tissues through a graded ethanol series (50%; 75%; 95%; 100% x 2), 10 minutes per step.</li>
	<li>Prepare the infiltration solution: 25 ml of JB-4 Embedding Solution A (Polysciences 0226A) and 0.24g of catalyst (Polysciences 02618). It takes a while for the catalyst to dissolve, about 20-30 minutes. The solution can be stored at 4&#176;C up to 1 month.</li>
	<li>Decant off the 100% ethanol and replace with plastic infiltration solution. Place the vials a 4&#176;C on a rotator and allow infiltration to proceed until the larvae sink to the bottom of the glass vial. This process usually takes about 30 minutes.</li>
	<li>Embedding protocol: Place the plastic molding cup tray (Polysciences, Inc. 16643A) under a dissecting microscope. Transfer the larvae, along with infiltration solution to cover, into the cups. The plastic embedding media is prepared in a small plastic tube. We normally embed 4-5 fish at a time, one fish for each cup. Prepare 5-6 ml of embedding plastic (1.2 ml of embedding solution for each cup) adding 35 &#956;l of JB-4 Embedding Solution &#34;B&#34; (Polysciences Inc. 0226B) for each ml of infiltration solution. Add 35 &#956;l of JB-4 Embedding Solution &#34;B&#34; (Polysciences Inc. 0226B) for each ml of infiltration solution. Mix well, pipetting up and down with a plastic Pasteur pipette several times. This is important because if not mixed well the JB-4 won&#39;t polymerize properly. After 5ml of embedding plastic has been prepared, remove infiltration plastic from around samples with a Pasteur pipette. Fill the cup with embedding plastic. The plastic must ooze out of the cup, to ensure good attachment of the block holder to the specimen. To detect zebrafish renal tubules, transverse sections are the best. For this purpose, the fish are embedded vertically, head down. With the aid of an eyelash with handle or fine forceps, periodically position the fish vertically, until the plastic is hard enough that they don&#39;t move from their position. It takes about 20-30 minutes. Then, place the EBH-2 Block Holder (Polysciences Inc. 15899) over the cups and leave the tray at 4&#176;C overnight. When polymerization in complete, you can pop out the blocks like ice cubes.</li>
	<li>Alternative embedding protocol for multiple larvae: Fill molds halfway with complete JB-4 and allow it to polymerize. For the 15 mm square molds we use the JB-4 volume at 700&#956;l to fill the mold halfway. Next, add the JB-4 soaked larvae with hardener to the pre-polymerized half-filled molds. Orient the larvae with the head pointing toward the long side of the mold and align the larvae so the eyeballs line up (you can put 10 fish in a single mold in a line). After the block is polymerized you can cut the block edges, turn it so the side is facing the knife and superglue it to a plastic &#34;chuck&#34; that fits in the microtome. With this method you can position a large number of larvae in the same block, centered in the plastic, and ready for cross sections. Care has to be taken orienting the block to the knife so that equivalent sections of each fish are taken in a single section. This can be accomplished while sectioning through the eyes: i.e. orient the block so that equivalent sections of the eyes are taken for each fish. The end result is that you can analyze many fish concomitantly on a single microscope slide.</li>
</ul>
Part 4: Sectioning
<ul>
	<li>Set the specimen into the chuck holder of the microtome (we use a Shandon Finesse Thermo Electro Corporation microtome)</li>
	<li>Set the slide warmer to 45&#176;C.</li>
	<li>Pour DI water into a beaker and place it close to the microtome</li>
	<li>Before sectioning, be sure the block is oriented such as to obtain perfect transverse sections of the fish. The orientation head adjustment levers can be rotated enabling to position the specimen head accurately to the knife. Start cutting thick sections through the head of the fish. While sectioning through the eyes, orient the block so that equivalent sections of the eyes are taken for each fish. When the pectoral fins appear in the sections, that is when the pronephric tubules start. From this point, cut about 32 4 um sections from the block.</li>
	<li>Collect the sections with forceps and let them drop into the beaker with water, without touching the water with the forceps. The section will expand as soon as it hits the surface of the water, now you can collect it on a slide. Insert the slide into the water at a 45&#176; angle and approach the section with the help of an eyelash with handle, touching only the borders of the section.</li>
	<li>Incubate the slides on a slide warmer until completely dry.</li>
	<li>Image sections using epifluorescent or confocal microscopy.</li>
</ul>
Part 5: Representative Results
When performed correctly, injected fish appear like in figure 1, with little fluorescent dextran (in this case Lucifer Yellow) visible in the heart and in the circulatory system. If the injection is too deep, injected material can be seen concentrated in the yolk sac, or the heart can be punctured causing excessive bleeding. If the holding pipette is made correctly, the fish won&#39;t roll or move when injected, and this will allow a more accurate injection. Also, it&#39;s important to make sure not to have any air bubbles in the MMP, which would slow the response time and decrease the precision of the system. Generally, as a consequence of gentamicin nephrotoxicity, 80% of the injected fish develop pericardial edema (Fig. 2). We use a phenotype based classification system, in which fish with moderate edema show evident pericardial edema but no others observable phenotypes, whereas severely edematous fish show pericardial and trunk edema, with slight to moderate axis curvature. Using a renal tubule antibody against Na+/K+ ATPase (&#945;-6F) we can visualize the damaged tubules in transverse sections, with an evident loss of cell polarity and tubular disruption in damaged tubules as compared to controls (Fig. 3). This pattern is consistent through multiple transverse sections. In order to obtain perfect transverse sections of the renal tubules is important to position the fish correctly during the embedding. When comparing sections, pectoral fins and proximal convoluted tubules are used as anatomical markers.
<img alt="Figure 1" src="/files/ftp_upload/2079/2079Fig1.jpg" /> 
Figure 1. Nephrotoxin microinjection. (A) Successful injection of a 10kDa dextran conjugated to lucifer yellow, with faint expression in the common cardinal vein, white arrow. (B) Incorrect injection, resulting in a deposit of 10kDa dextran conjugated to lucifer yellow that does not dissipate into the circulatory system, red arrow.
<img alt="Figure 2" src="/files/ftp_upload/2079/2079Fig2.jpg" /> 
Figure 2. Gentamicin-mediated edema. (A) Control injected larval zebrafish. Gentamicin injected larval zebrafish with (B) moderate or (C) severe edema. All larvae are at 4 dpf with anterior to the left and dorsal is up. Black arrows point to pericardial edema in B and C.
<img alt="Figure 3" src="/files/ftp_upload/2079/2079Fig3.jpg" /> 
Figure 3. AKI immunohistochemistry. &#945;-6F antibody staining for Na+/K+ ATPase 48 hours-post gentamicin injection (4 dpf larvae) in control (A,B) and gentamicin injected (C,D) larvae. A, C are at 20X magnification and of B and D are 3X digital magnifications of the right tubules in A and C. Note loss of basolateral polarity and tubule disruption, white arrow, in D as compared to B.

<ol>
	<li>Hentschel, D. M., Park, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+renal+failure+in+zebrafish:+a+novel+system+to+study+a+complex+disease.">Acute renal failure in zebrafish: a novel system to study a complex disease.</a> Am J Physiol Renal Physiol. 288 (5), F923-F929 (2005).</li><li>Majumdar, A., Lun, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+no+isthmus+reveals+a+role+for+pax2.1+in+tubule+differentiation+and+patterning+events+in+the+pronephric+primordia.">Zebrafish no isthmus reveals a role for pax2.1 in tubule differentiation and patterning events in the pronephric primordia.</a> Development. 127 (10), 2089-2098 (2000).</li><li>Weinstein, B. M., Stemple, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gridlock,+a+localized+heritable+vascular+patterning+defect+in+the+zebrafish.">Gridlock, a localized heritable vascular patterning defect in the zebrafish.</a> Nat Med. 1 (11), 1143-1147 (1995).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book. , (1993).</li></ol>

No conflicts of interest declared.

In this video article we demonstrated an intravenous microinjection technique for an AKI model, by creating gentamicin mediated proximal tubular damage in zebrafish larvae. This damage results in the formation of pericardial and/or gross edema, reflecting an inability to regulate water balance. We also described an immunohistochemistry experiment with an antibody that marks differentiated kidney tubules (Majumdar et al. 2000), showing loss of cell polarity and a disruption of the proximal tubule in damaged kidneys. Intravenous microinjections represent an efficient method for delivering a soluble substance into the bloodstream of zebrafish larvae. This technique is an excellent tool for the introduction of a variety of substances and with the aid of a fluorescent marker uniform injections can be repeated many times over allowing for consistent results.

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<td>&alpha;6F mouse monoclonal </td>
<td>&nbsp;</td>
<td>Developmental study Hbridoma Bank</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Block Holder EBH-2</td>
<td>&nbsp;</td>
<td>Polysciences, Inc.</td>
<td>15899</td>
<td>&nbsp;</td>
<tr>
<td>Borosilicate Glass Capillaries</td>
<td>&nbsp;</td>
<td>World Precision Instruments, INC</td>
<td>TW100-6</td>
<td>&nbsp;</td>
<tr>
<td>Capillary tubes R-6 Custom Glass Tubing </td>
<td>&nbsp;</td>
<td>Drummond Scientific Company, Broomall PA</td>
<td>9-000-3000</td>
<td>&nbsp;</td>
<tr>
<td>Catalyst Powder</td>
<td>&nbsp;</td>
<td>Polysciences, Inc.</td>
<td>02618</td>
<td>&nbsp;</td>
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<td>Cy3</td>
<td>&nbsp;</td>
<td>Jackson Immunoresearch Laboratories, Inc.</td>
<td>155-165-003</td>
<td>&nbsp;</td>
<tr>
<td>Dextran 10k Lucifer yellow, lysine fixable</td>
<td>&nbsp;</td>
<td>Molecular probes</td>
<td>D1825</td>
<td>&nbsp;</td>
<tr>
<td>DMSO</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>D8418</td>
<td>&nbsp;</td>
<tr>
<td>Eyelash</td>
<td>&nbsp;</td>
<td>Ted Pella</td>
<td>113</td>
<td>&nbsp;</td>
<tr>
<td>Gentamicin sulfate</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich </td>
<td>G8648</td>
<td>&nbsp;</td>
<tr>
<td>Glass vials 4ml</td>
<td>&nbsp;</td>
<td>Weathon</td>
<td>225012</td>
<td>&nbsp;</td>
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<td>Iron plate</td>
<td>&nbsp;</td>
<td>Nariscige</td>
<td>IP</td>
<td>&nbsp;</td>
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<td>JB-4 Embedding Solution A</td>
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<td>Polysciences Inc.</td>
<td>0226A</td>
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<td>Polysciences, Inc.</td>
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<td>Magnetic stand</td>
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<td>MN-151</td>
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<td>Manual microsyringe pump</td>
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<td>World Precision Instruments, Inc</td>
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<td>Microinjection apparatus</td>
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<td>0121-1</td>
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<td>Kopf</td>
<td>720</td>
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<td>Plastic molding cup tray</td>
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<td>Polysciences, Inc.</td>
<td>16643A</td>
<td>&nbsp;</td>
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<td>Razor blade</td>
<td>&nbsp;</td>
<td>WVR Scientific</td>
<td>55411-050</td>
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<td>Slide warmer</td>
<td>&nbsp;</td>
<td>Lab line</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>Stage micrometer </td>
<td>&nbsp;</td>
<td>Wards </td>
<td>94 W 9910</td>
<td>&nbsp;</td>
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<td>Tilting base</td>
<td>&nbsp;</td>
<td>World Precision Instruments, Inc.</td>
<td>TB 1</td>
<td>&nbsp;</td>
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<td>Tricaine powder</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>A-5040</td>
<td>&nbsp;</td>
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<td>Tween 20</td>
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<td>P7949</td>
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intravenous microinjections of zebrafish larvae to study acute kidney injury

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections on 2 dpf zebrafish. This technique represents an efficient and rapid method to deliver soluble substances into the bloodstream of zebrafish larvae, allowing for the injection of 15-20 fish per hour. In addition to AKI studies, this microinjection technique can also be used for other types of experimental studies such as angiography.  We provide a detailed protocol of the technique from equipment required to visual measures of decreased kidney function. In addition, we also demonstrate the process of fixation, whole mount immunohistochemistry with a kidney tubule marker, plastic embedding and sectioning of the larval zebrafish. We demonstrate that zebrafish larvae injected with gentamicin show morphological features consistent with AKI: edema, loss of cell polarity in proximal tubular epithelial cells, and morphological disruption of the tubule.

Watch this Scientific Journal Video about Intravenous Microinjections of Zebrafish Larvae to Study Acute Kidney Injury at JoVE.com

Intravenous Microinjections of Zebrafish Larvae to Study Acute Kidney Injury

We describe a technique of microinjecting the aminoglycoside, gentamicin, into 2 days post-fetilization (dpf) zebrafish larvae to induce acute kidney injury (AKI). We also describe a method for whole mount immunohistochemistry, plastic embedding and sectioning of zebrafish larvae to visualize the AKI mediated damage.

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections ...

intravenous-microinjections-zebrafish-larvae-to-study-acute-kidney

Research

JoVE Journal

Biology

22.5K Aufrufe.  University of Pittsburgh. Wir beschreiben eine Technik der Mikroinjektion der Aminoglykosid, Gentamicin, in 2 Tagen nach fetilization (dpf) Zebrafischlarven zu akutem Nierenversagen (AKI) zu induzieren. Wir beschreiben ein Verfahren zur whole mount Immunhistochemie, Kunststoff eingebettet und Schnitte mit Zebrabärblingembryonen der AKI-vermittelten Schaden zu visualisieren.

Serie: Intravenous Microinjections of Zebrafish Larvae to Study Acute Kidney Injury

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of the visual system provide more empirical evidence for functional applications. Investigation into the sensory system provides information about the sensory capacity, learning and memory ability, and establishes known baseline behaviour in which to gauge deviations (Burghardt, 1977). However, unlike mammalian or avian systems, testing for learning and memory in a reptile species is difficult. Furthermore, using an operant paradigm as a psychophysical measure of sensory ability is likewise as difficult. Historically, reptilian species have responded poorly to conditioning trials because of issues related to motivation, physiology, metabolism, and basic biological characteristics. Here, I demonstrate an operant paradigm used a novel model lizard species, the Jacky dragon (Amphibolurus muricatus) and describe how to test peripheral sensitivity to salient speed and motion characteristics. This method uses an innovative approach to assessing learning and sensory capacity in lizards. I employ the use of random-dot kinematograms (RDKs) to measure sensitivity to speed, and manipulate the level of signal strength by changing the proportion of dots moving in a coherent direction. RDKs do not represent a biologically meaningful stimulus, engages the visual system, and is a classic psychophysical tool used to measure sensitivity in humans and other animals. Here, RDKs are displayed to lizards using three video playback systems. Lizards are to select the direction (left or right) in which they perceive dots to be moving. Selection of the appropriate direction is reinforced by biologically important prey stimuli, simulated by computer-animated invertebrates.

Testing visual sensitivity in lizards using an operant conditioning paradigm that employs video playback of random-dot kinematograms and computer-generated invertebrates.

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of ...

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 ...

Methods for the Study of the Zebrafish Maxillary Barbel

Barbels are skin sensory appendages found in fishes, reptiles and amphibians. The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal pair and a longer maxillary pair. Barbel tissue contains cells of ectodermal, mesodermal and neural crest origin, including skin cells, glands, taste buds, melanocytes, circulatory vessels and sensory nerves. Unlike most adult tissue, the maxillary barbel is optically clear, allowing us to visualize the development and maintenance of these tissue types throughout the life cycle. 
This video shows early development of the maxillary barbel (beginning approximately one month post-fertilization) and demonstrates a surgical protocol to induce regeneration in the adult appendage (&gt;3 months post-fertilization). Briefly, the left maxillary barbel of an anesthetized fish is elevated with sterile forceps just distal to the caudal edge of the maxilla. A fine, sterile spring scissors is positioned against the forceps to cut the barbel shaft at this level, establishing an anatomical landmark for the amputation plane. Regenerative growth can be measured with respect to this plane, and in comparison to the contralateral barbel. Barbel tissue regenerates rapidly, reaching maximal regrowth within 2 weeks of injury.
Techniques for analyzing the regenerated barbel include dissecting and embedding matched pairs of barbels (regenerate and control) in the wells of a standard DNA electrophoresis gel. Embedded specimens are conveniently photographed under a stereomicroscope for gross morphology and morphometry, and can be stored for weeks prior to downstream applications such as paraffin histology, cryosectioning, and/or whole mount immunohistochemistry. These methods establish the maxillary barbel as a novel in vivo tissue system for studying the regenerative capacity of multiple cell types within the genetic context of zebrafish.

The zebrafish maxillary barbel is an integumentary sense organ containing ectodermal, mesodermal and neural crest derivatives. Importantly, the adult barbel can regenerate after proximal amputation. This video introduces maxillary barbel development and demonstrates a surgical protocol to induce regeneration, followed by collection, embedding and downstream imaging of barbel specimens.

Barbels are skin sensory appendages found in fishes, reptiles and amphibians.  The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal ...

Using the optokinetic response to study visual function of zebrafish

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the visual functions of larval zebrafish1-5. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adult zabrafish. Here, we introduce how to measure the OKR of adult zebrafish with our simple custom-built apparatus using a new protocol which is established in our lab. Both our apparatus and step-by-step procedure of OKR in adult zebrafish are illustrated in this video. In addition, the measurements of the larval OKR, as well as the optomotor response (OMR) test of adult zebrafish, are also demonstrated in this video. This OKR assay of adult zebrafish in our experiment may last for up to 4 hours. Such OKR test applied in adult fish will benefit to visual function investigation more efficiently when the adult fish vision system is manipulated.
Su-Qi Zou and Wu Yin contributed equally to this paper.

Optokinetic response has been widely used to assess the visual functions of larval zebrafish. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adults1-5. Here, we introduce how to measure the OKR of adult zebrafish using a new protocol which is established in our lab.

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess 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 ...

Laser Ablation of the Zebrafish Pronephros to Study Renal Epithelial Regeneration

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in renal failure1. AKI can be caused by a number of factors including ischemia, drug-based toxicity, or obstructive injury1. This results in an inability to maintain fluid and electrolyte homeostasis. While AKI has been observed for decades, effective clinical therapies have yet to be developed. Intriguingly, some patients with AKI recover renal functions over time, a mysterious phenomenon that has been only rudimentally characterized1,2. Research using mammalian models of AKI has shown that ischemic or nephrotoxin-injured kidneys experience epithelial cell death in nephron tubules1,2, the functional units of the kidney that are made up of a series of specialized regions (segments) of epithelial cell types3. Within nephrons, epithelial cell death is highest in proximal tubule cells. There is evidence that suggests cell destruction is followed by dedifferentiation, proliferation, and migration of surrounding epithelial cells, which can regenerate the nephron entirely1,2. However, there are many unanswered questions about the mechanisms of renal epithelial regeneration, ranging from the signals that modulate these events to reasons for the wide variation of abilities among humans to regenerate injured kidneys.
The larval zebrafish provides an excellent model to study kidney epithelial regeneration as its pronephric kidney is comprised of nephrons that are conserved with higher vertebrates including mammals4,5. The nephrons of zebrafish larvae can be visualized with fluorescence techniques because of the relative transparency of the young zebrafish6. This provides a unique opportunity to image cell and molecular changes in real-time, in contrast to mammalian models where nephrons are inaccessible because the kidneys are structurally complex systems internalized within the animal. Recent studies have employed the aminoglycoside gentamicin as a toxic causative agent for study of AKI and subsequent renal failure: gentamicin and other antibiotics have been shown to cause AKI in humans, and researchers have formulated methods to use this agent to trigger kidney damage in zebrafish7,8. However, the effects of aminoglycoside toxicity in zebrafish larvae are catastrophic and lethal, which presents a difficulty when studying epithelial regeneration and function over time. Our method presents the use of targeted cell ablation as a novel tool for the study of epithelial injury in zebrafish. Laser ablation gives researchers the ability to induce cell death in a limited population of cells. Varying areas of cells can be targeted based on morphological location, function, or even expression of a particular cellular phenotype. Thus, laser ablation will increase the specificity of what researchers can study, and can be a powerful new approach to shed light on the mechanisms of renal epithelial regeneration. This protocol can be broadly applied to target cell populations in other organs in the zebrafish embryo to study injury and regeneration in any number of contexts of interest.

Acute kidney injury (AKI) in humans is a common clinical problem caused by damage to the epithelial cells that comprise kidney nephrons, and AKI is associated with high mortality rates of 50-70%1. Following epithelial cell destruction, nephrons have a limited ability to regenerate, though the mechanisms and limitations that guide this phenomenon remain poorly understood. In this video article, we describe our technique for targeted laser ablation of kidney nephron cells in the zebrafish embryo kidney, or pronephros. Our new method can be used to complement nephrotoxicity-induced models of AKI and gain a high-resolution understanding of the cell and molecular alterations that are associated with epithelial regeneration in the kidney nephron.

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

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

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

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

Microgavage of Zebrafish Larvae

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

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

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

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional measures of muscle health. In this video article, we demonstrate a method to measure force generation during contraction of zebrafish larval trunk muscle. Force measurements are accomplished by placing an anesthetized larva into a chamber filled with a salt solution. The anterior end of the larva is tied to a force transducer and the posterior end of the larva is tied to a length controller. An isometric twitch contraction is elicited by electric field stimulation and the force response is recorded for analysis. Force generation during contraction provides a measure of overall muscle health and specifically provides a measure of muscle function. Although we describe this technique for use with wild-type larvae, this method can be used with genetically modified larvae or with larvae treated with drugs or toxicants, to characterize muscle disease models and evaluate treatments, or to study muscle development, injury, or chemical toxicity.

Force measurements can be used to demonstrate changes in muscle function due to development, injury, disease, treatment or chemical toxicity. In this video, we demonstrate a method to measure force during a maximal contraction of zebrafish larval trunk muscle.

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional ...