We thank V. Zimmermann for excellent technical assistance and for fish care, D. K&#246;nig (University of Fribourg) for critical reading of the manuscript, D. Kressler (University of Fribourg) for help with zCNTF protein synthesis, F. Ruggiero (Institut de G&#233;nomique Fonctionnelle de Lyon) for providing ColXII antibody, and P. Martin (University of Bristol) for L-plastin antibody. We thank the imaging core facility and the proteomics platform at the University of Fribourg. This work was supported by the Swiss National Science Foundation, grant number 310030_179213, and by the Schweizerische Herzstiftung (Swiss Heart Foundation).

Animal care and all animal procedures described in the following protocol were approved by the cantonal veterinary office of Fribourg, Switzerland.
1. Tools and Solutions for Injections
<ol>
	<li>Pull microinjection-adapted borosilicate glass capillaries using a needle puller according to Figure 1A. Store pulled capillaries in a 9 cm Petri dish with rails of modeling clay or adhesive tape.</li>
	<li>Using common scissors, cut a piece of sponge (7 cm x 3 cm x 1 cm) and carve a fish-like silhouette in its middle.</li>
	<li>Prepare small aliquots of injection solution with the tested proteins or other compounds. Adjust their concentration dependently on the assay by diluting the substance in 1x Hanks balanced salt solution (HBSS) complemented with 10% phenol red. 
	NOTE: Here, the concentration of the tested protein was 100 ng/mL.</li>
	<li>To prepare a stock solution of buffered tricaine anesthetics, dissolve 4 g of tricaine in 980 mL of distilled water. Adjust pH to 7.0&#8722;7.4 using 1 M Tris-HCl pH 9, and fill up with water to 1,000 mL. Store the solution in dark at 4 &#176;C.</li>
	<li>To obtain the working concentration of anesthetics, add 1&#8722;2 mL of tricaine stock solution into 50 mL of fish water in a beaker. 
	NOTE: The working concentration of tricaine anesthetics should be prepared freshly before use.</li>
</ol>
2. Preparation of the Injection Station
<ol>
	<li>Turn on the stereomicroscope with the light from the top and adjust magnification to 16x.</li>
	<li>Soak the sponge with fish water, place it on a 9 cm Petri dish on the microscope stage and adjust focus.</li>
	<li>Under the stereomicroscope, cut the end of a microcapillary at ~7 mm from the basis using an iridectomy scissors as shown in Figure 1A. The ideal tip diameter would be ~20 &#181;m. 
	NOTE: Cutting the tip of the capillary in an oblique way is optimal for insertions into the tissue.</li>
	<li>Insert the microcapillary into the needle holder of the microinjector apparatus.</li>
	<li>Using microloader tips, load a control solution (e.g., 1x HBSS) to set up the pressure of injection, in order to obtain the appropriate flow ranging between 0.3 &#181;L/s and 0.5 &#181;L/s. Empty the needle.</li>
	<li>Load the selected volume of the injection solution (e.g., ciliary neurotrophic factor [CNTF] diluted in 1x HBSS) into the tip of the capillary (Figure 1B). There should be no air bubble in the capillary. 
	NOTE: The maximal volume of injection solution depends on the fish size. For a standard length of 2.5&#8722;3 cm (distance from the snout to the caudal peduncle), the maximum injection volume that prevents exessive thoracic swelling and bleeding was determined to be 5 &#181;L (Figure 1F). Larger volumes could be injected to bigger fish.</li>
</ol>
3. Preparation of the Fish for Intrathoracic Injection
<ol>
	<li>Catch an adult zebrafish (Danio rerio) with a net and transfer it into the anesthetic solution.</li>
	<li>After 1&#8722;2 min, when fish stops swimming and the movement of operculum is reduced, touch the fish with a plastic spoon to make sure it does not react to any contact.</li>
	<li>Quickly and carefully transfer the fish with the spoon into the groove of the wet sponge, with ventral side up. The head of the fish should point away from the operator&#8217;s dominant-hand.</li>
</ol>
4. Microinjection into the Pericardium
<ol>
	<li>Under the stereomicroscope, carefully observe the movement of the beating heart under the skin of the fish. Visually determine the injection point above the beating heart and in the middle of the triangle defined by the ventral cartilaginous plates (Figure 1D). Insert the tip of the capillary at 30&#8722;45&#176; degree angle relative to the body axis (Figure 1E). Gently penetrate the skin with the tip of the microcapillary into the pericardium (Figure 1C). An optimal entry point is closer to the abdomen than to the head. 
	NOTE: Do not insert the capillary too deeply into the body and the heart, as this will cause injury to the organ. In case of heart puncture, the needle generally fills with blood. If this happens, remove the capillary and exclude the fish from the experiment.</li>
	<li>Once the needle is inside the pericardium, complete injection by pressing the pedal of the microinjector device. 
	NOTE: Be careful not to inject air into the thoracic cavity.</li>
	<li>After injection, gently withdraw the capillary from the thorax and immediately transfer the fish into a tank with system water for recovery.</li>
	<li>Monitor the fish until total recovery from anesthesia.</li>
	<li>Collect heart at the desired time point and prepare it for further analysis. 
	NOTE: In case the fish does not resume movement of the operculum within 30 s, reanimate the fish by squeezing water into the gills with a plastic pipette.</li>
</ol>

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The authors have nothing to disclose.

Here, we describe a method for delivering exogenous compounds and proteins into the pericardial cavity in order to study their effects on the heart in adult zebrafish. The procedure is based on intrathoracic injection, which results in the delivery of a small volume of solution in the vicinity of the organ. This technique was developed and described for studying cardiac preconditioning and regeneration.
The critical step in this procedure is the penetration of the glass capillary into the thoracic cavity. This step depends on three parameters which are: the rigidity and sharpness of the capillary tip, the angle of penetration, and the puncture site. To optimize the penetration through the skin, the pulled part of the capillary should not to be too long, as such needles are too flexible and bend in contact with the skin. To avoid this, the rigidity can be adapted by reducing the tip size with the iridectomy scissor. Although the angle of penetration can vary between 30&#176; and 45&#176;, it can be adapted to the rigidity of the tip. Indeed, a thin tip will penetrate the skin better with a narrower angle.
In order to optimize the needle penetration, the insertion site should be immediately above the beating heart. The risk of heart puncture is usually low ranging between 5% and 8%. Insertion of the needle posterior to the heart increases the risk of heart puncture, as seen by enhanced bleeding. In such cases, the animals should be removed from the experiments.
Another source of trouble during the IT injection occurs at the capillary level. Indeed the capillary can break when lateral forces are exerted on it. To avoid this, the needle must move along the axis of injection in a straight way. Occasionally, the capillary can be blocked by tissue residues that prevent the liquid from flowing. The needle can be unblocked by gently withdrawing the tip whilst injecting. If this does not improve the flow, we recommend completely withdrawing the needle from the thorax and replacing the needle.
Lesions can be caused by a too deeply inserted needle in the pericardium. In order to avoid lesions in the pericardial sac, the needle must not be inserted too much (1&#8722;2 mm) into the thorax. Some leaks were observed when the injection volume was bigger than 8 &#181;L.
In zebrafish, the exact composition of the pericardial fluid is unknown. However, the volume of the pericardial cavity is estimated at ~10 &#181;L31. Given that the volume of the adult zebrafish ventricle is approximately 1&#8722;2 mm3, we assume that the pericardial cavity accordingly has a tiny volume, which has to be considered before injections. From our preliminary studies, we determined that the optimal range of the injected volume is between 0.5 and 3 &#181;L for fish measuring 2.5&#8722;2.8 cm (distance from the snout to the caudal peduncle). This volume can be adapted depending on the size of the fish. Injection of up to 5 &#181;L did not induce any lesion in the fish of this size. However, volumes from 8 &#181;L were sufficient to cause bulging and internal bleeding as shown in Figure 1F. Based on this data, we estimate that an amount of solution bigger than 3 &#181;L might cause physical and physiological stress on the organ. This limitation infers the need to choose a higher concentration of molecules instead of increasing the amount of the injected solution.
Another important factor is the osmotic property of the injected solution, which should be in the physiological range. Indeed, to avoid a risk of osmotic stress, we recommend HBSS as injection medium.
In zebrafish, the common methods used to deliver drugs are through water treatment and intraperitoneal injection30,35. Although both of these techniques are suitable for many applications, IT injections provide experimental and economic advantages, by decreasing the risks of undesired systemic side effects and reducing the usage of costly molecules, respectively. This method can be suitable for delivery of tamoxifen to activate the Cre-ERT2 transgenic system used for cell lineage tracing analysis, and guide modified RNAs for functional studies in regeneration research.
The IT-injection method in zebrafish has been previously described31,36. In those reports, intrathoracic injections were performed with insulin needle, puncturing from the anterior side. In contrast, our protocol presents an alternative strategy with the pulled glass capillary inserted from the posterior direction. Specifically, our approach takes into account the anatomy of the fish pericardium to optimize the injection with a reduced risk of heart puncture. Furthermore, during the procedure, the fish is not held by metallic forceps, but by a moist and soft sponge, which is a more suitable method to avoid any external injury of the fish. Thus, the presented method might be better suited for studies of cardiac homeostasis, preconditioning and regeneration in adult zebrafish.
IT injections have already been established in mammalian model organisms. Indeed, this method has also been applied in experiments with pigs and clinical studies in humans37,38. In mice, transthoracic intramyocardial injections guided by ultrasounds have been used to challenge their heart39. Within this article we propose a detailed protocol to ease the use of IT injection for zebrafish. This will be particularly valuable to the field, in order to complement genetic approaches in cardiac homeostasis, preconditioning and regeneration research.

Among vertebrates, zebrafish possess a remarkable capacity to regenerate their hearts1,2. This ability has been reported in several injury models, namely ventricular apex resection, cryoinjury (CI) and genetic cardiomyocyte ablation3,4,5,6,7. After invasive injuries, the damaged wall of the ventricle becomes transiently healed by fibrotic tissue, which is progressively replaced by a new myocardium8,9,10,11. The early wound healing response involves epicardium activation and recruitment of immune cells12,13,14,15. Concomitantly, cardiomyocytes near the injured myocardium become activated, dedifferentiate, proliferate and progressively replace the wounded area within 30&#8722;90 days16,17,18,19. Substantial progress in deciphering the molecular and cellular mechanisms of cardiac regeneration has been achieved thanks to the availability of genetic tools, such as cell-lineage tracing analysis, inducible gene overexpression, fluorescent tissue reporter lines, and CRISPR/Cas9 gene mutagenesis20,21.
We have recently established a model of cardiac preconditioning in the adult zebrafish by thoracotomy22,23. Preconditioning increases expression of cardioprotective genes and elevates the re-entry into the cell cycle in the intact and regenerating hearts. These processes are associated with the recruitment of immune cells and matrix remodeling22,24. The mechanisms of preconditioning are poorly understood, and the establishment of new techniques is required to foster this area of research. In particular, optimized administration of secreted signaling proteins or other chemical compounds is essential to further investigate this topic.
Being aquatic animals, zebrafish can naturally absorb various substances dissolved in water through their gills and skin. This offers a possibility for non-invasive drug delivery through immersion of fish in solutions with diverse chemicals, such as pharmacological inhibitors, steroid hormones, tamoxifen, BrdU and antibiotics. Indeed, numerous studies from various laboratories, including ours25,26,27, have taken advantage of this method, which is particularly valuable in the field of regenerative biology6,28. This approach is, however, not appropriate for delivery of peptides, DNA, RNA, morpholinos or molecules with a limited tissue permeability. In these cases, a more efficient delivery is achieved by microinjection into the body, for example, by inserting the capillary into the retro-orbital venous sinus, into the intraperitoneal or intrapericardial cavity29,30,31. Here, we describe a procedure of intrathoracic injection of a small amount of solution, as a suitable method to study heart regeneration and preconditioning in adult zebrafish.

<table>
	<tbody>
		<tr>
			<td>Hanks Balanced Salt Solution</td>
			<td>Gibco by Life technology</td>
			<td>14065-056</td>
			<td></td>
		</tr>
		<tr>
			<td>Iridectomy scissor</td>
			<td>Roboz Surgical Instruments Co</td>
			<td>RS-5602</td>
			<td></td>
		</tr>
		<tr>
			<td>Macroscope (binocular)</td>
			<td>M400</td>
			<td></td>
			<td>with Apozoom</td>
		</tr>
		<tr>
			<td>Micro-injector femtojet</td>
			<td>Eppendorf</td>
			<td>5247 0034 77</td>
			<td></td>
		</tr>
		<tr>
			<td>Microloaders femtotips</td>
			<td>Eppendorf</td>
			<td>5242 956.003</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette glass needles type C</td>
			<td>WPI</td>
			<td>TW100F-6</td>
			<td>thin-wall capillary</td>
		</tr>
		<tr>
			<td>Micropipette puller model P-87</td>
			<td>Flaming/Brown</td>
			<td>20081016</td>
			<td>filament box 2.5 x 4.5 mm</td>
		</tr>
		<tr>
			<td>Sponge</td>
			<td>any</td>
			<td>any</td>
			<td>dim. carved sponge 7cm x 3 cm x 1 cm</td>
		</tr>
		<tr>
			<td>Tricaine (Anestethic)</td>
			<td>Sigma</td>
			<td>E10521</td>
			<td></td>
		</tr>
		<tr>
			<td>Dyes and Antibodies</td>
			<td>Company</td>
			<td>Catalog number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>anti-Chicken Cy5</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td></td>
			<td>Concentration: 1 / 500</td>
		</tr>
		<tr>
			<td>anti-Guinea pig Cy5</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td></td>
			<td>Concentration: 1 / 500</td>
		</tr>
		<tr>
			<td>anti-Rabbit Cy5</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td></td>
			<td>Concentration: 1 / 500</td>
		</tr>
		<tr>
			<td>Chicken l-plastin</td>
			<td>gift from P. Martin, Bristol</td>
			<td></td>
			<td>Concentration: 1 / 1000</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma</td>
			<td>10236276001</td>
			<td>Concentration: 1 / 2000 (1&#181;g/ml); 1/100 IT injected</td>
		</tr>
		<tr>
			<td>Guinea pig anti-ColXII</td>
			<td>gift from Florence Ruggerio, Lyon</td>
			<td></td>
			<td>Concentration: 1 / 500</td>
		</tr>
		<tr>
			<td>Phalloidin-Atto-565 (F-actin)</td>
			<td>Sigma</td>
			<td>94072</td>
			<td>Concentration: 1 / 500</td>
		</tr>
		<tr>
			<td>Phalloidin-Atto-647 (F-actin)</td>
			<td>Sigma</td>
			<td>95906</td>
			<td>Concentration: 1 / 50 IT injected</td>
		</tr>
		<tr>
			<td>Rabbit anti-MCM5</td>
			<td>gift from Soojin Ryu, Heidelberg</td>
			<td></td>
			<td>Concentration: 1 / 500</td>
		</tr>
		<tr>
			<td>Stamping Ink 4K</td>
			<td>Pelikan</td>
			<td>1 4k 351 197</td>
			<td>Concentration: 1 / 1</td>
		</tr>
		<tr>
			<td>ISH probe primers</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cystatin</td>
			<td></td>
			<td></td>
			<td>gene number: ENSDARG00000074425 
			&#8203;fw primer: GATTCACTGTCGGGTTTGGG 
			Rev primer: ATTGGGTCCATGGTGACCTC</td>
		</tr>
	</tbody>
</table>

intrathoracic injection for the study of adult zebrafish heart

The adult zebrafish heart provides a powerful model in cardiac regeneration research. Although the strength of this system is based on transgenic approaches, a rapid delivery of exogenous factors provides a complementary technique in functional studies. Here, we present a method that relies on administration of a few microliters of solution into the pericardial cavity without causing myocardial damage. Intrathoracic (IT) injections can efficiently deliver proteins and chemical compounds directly onto the heart surface. The injected substances diffuse through the epicardium into the underlying cardiac tissues. Compared to intraperitoneal (IP) injections, the main advantage of intrathoracic injections is the focal administration of the tested factors on the target organ. The delivery of molecules directly into the pericardium is a suitable strategy for studies of cardiac preconditioning and regeneration in adult zebrafish.

After intrathoracic (IT) injections, the effects of exogenous solution can be analyzed. For this purpose, the fish should be euthanized and the hearts collected, fixed and histologically processed, according to previously published protocols32,33.
To validate the method, we first performed two test experiments by injecting color and fluorescent dyes. First, we euthanized fish and post-mortem injected 3 &#181;L of ink into the thorax. The hearts were collected after 5 min, washed in phosphate-buffered saline (PBS), fixed in 2% formalin, washed in PBS and photographed under the microscope. Second, we injected 3 &#181;L of 1 &#181;g/mL 4&#8242;,6-diamidino-2-phenylindole (DAPI) in vivo, and fixed the heart after 2 h. In both assays, whole-mount analysis revealed labeling of the entire heart including the ventricle, the atrium and the bulbus arteriosus (Figure 2A,B). These results reveal efficient spreading of the injected solution on the heart surface.
A common protocol for delivery of exogenous substances into the adult fish is intraperitoneal (IP) injection. To compare the suitability of IT versus IP injections for heart studies, we injected a similar amount of DAPI using both methods and fixed the hearts after 5 min and 120 min (Figure 3A). The hearts were sectioned and stained with phalloidin Alexa Fluor (AF) 568 that labels F-actin in cardiac muscle. No DAPI-positive cells were observed in the hearts after IP injection at both time points (Figure 3B). By contrast, IT injection resulted in the presence of DAPI-labeled nuclei in the myocardium (Figure 3B). These results demonstrate that IT injection improved the delivery of the compound to the heart, as compared to the IP injection.
To test the suitability of this method for heart regeneration studies, we cryoinjured ventricles8, and performed IT injections of 3 &#181;L of 1 &#181;g/mL DAPI and 1 &#181;g/mL phalloidin AF649 at 3 and 7 days post-cryoinjury (dpci) (Figure 4A). At 1 h post-injection, the hearts were collected, fixed, sectioned and stained with phalloidin AF568 to visualize the intact myocardium. We found that both the myocardium and the injured tissue contained numerous DAPI positive cells, indicating an efficient penetration of this dye into the intact heart and fibrotic tissue (Figure 4B). Furthermore, injected phalloidin AF649 was also incorporated by cardiomyocytes of the peri-injury zone and some recruited fibroblasts of the wounded area. This experiment reveals that the drugs can cross the epicardium and penetrate into the underlying myocardium.
After testing the efficiency of IT injections using dyes, we analyzed the effects of injected proteins on the heart. We synthetized a cytokine, called CNTF, which is upregulated after thoracotomy24. We investigated the effects of the exogenous CNTF&#160;on various processes, namely cardiomyocyte proliferation, extracellular matrix deposition, immune cell recruitment and cardioprotective gene expression. We found that all these biological aspects were activated by IT injection of CNTF, as compared to control immunoglobulins (Figure 5)24. These results demonstrate that the method of intrathoracic injection provides a suitable strategy for targeted delivery of proteins to study their effects on distinct heart tissues in a variety of assays.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59724/59724fig1.jpg" /> 
Figure 1: Intrathorascic (IT) injection in adult zebrafish. (A) Photograph of a pulled microinjection capillary with filament (6&#8221;, 1.0 mm in diameter) and values of the needle puller program used. (B) Photograph of a pulled microinjection capillary with filament (6&#8221;, 1.0 mm in diameter) filled up with 2.5 &#181;L of solution containing 10% phenol red. The pulled tip of the needle is maximally 7 mm long. (C) Schematic representation of the IT injection procedure. (D) Photographs of the IT injection procedure. This figure has been modified from Bise et al.24. Numbers in panels C and D correspond to the same steps of the procedure: (1) fish is placed ventral side up on a humidified sponge. The puncture site (red dot in the triangle) is located in the center of the chest near the gills. (2) Penetration of the needle into the pericardium. Red dot indicates puncture site. (3) Injection is monitored by observing the spread of the red solution in the pericardial cavity. (E) Scheme of IT injection. Angle between the injection capillary and the body axis should be between 30&#176; and 45&#176; to avoid heart puncture. (F) Photographs of fish thorax at 1 hour after IT injection of indicated volumes. White arrows are pointing at the redish tissue, which might indicate internal bleeding. <a href="https://www.jove.com/files/ftp_upload/59724/59724fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59724/59724fig2.jpg" /> 
Figure 2: IT injected solutions spread nearly uniformly on the heart surface. (A) Stereomicroscope images of whole hearts of fish subjected to post-mortem IT injection with 2.5 &#181;L HBSS or 2.5 &#181;L ink. Ink stained the surface of the ventricle (V), atrium (A) and bulbus arteriosus (Ba). Scale bar = 300 &#181;m. (B) Bright-field and fluorescent stereomicroscope images of whole hearts of fish subjected to IT injection with HBSS and 3 &#181;L of 1 &#181;g/mL DAPI. DAPI fluorescence is detected on the heart parts soon after IT injection. Scale bar = 300 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59724/59724fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59724/59724fig3.jpg" /> 
Figure 3: Comparison of two injection methods for the delivery of DAPI to the heart. (A) Scheme of the experimental design. Intraperitoneal (IP) and Intrathorascic (IT) injections were performed with the same amount of 1 &#181;g/mL DAPI (3 &#181;L). Hearts were collected at 5 and 120 min after injection. (B) Confocal microscopy images of heart sections stained with fluorescent phalloidin (red) that abundantly labels muscle fibers. Injected DAPI was visualized in the appropriate channel shown in green.&#160;After IP injection, DAPI is not detected in the heart at any time point. After IT injection, DAPI positive cells are present in the ventricle after both time points. Scale bar = 500 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59724/59724fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59724/59724fig4.jpg" /> 
Figure 4: IT injection to study heart regeneration. (A) Scheme of the experimental design. At 3 and 7 days following cryoinjury, a mix of DAPI and phalloidin AF649 was IT-injected (3 &#181;L of 1 &#181;g/mL). Hearts were collected 1 hour after IT injection, fixed, sectioned and stained with phalloidin AF568 (red). (B) Confocal microscopy images of longitudinal heart sections at 3 and 7 dpci. Injected DAPI (green) and phalloidin AF649 (blue) label cells of the injured area (delimited by white dashed line) and the intact myocardium (red staining). White arrows are pointing at DAPI (green) distribution through intact compact and trabeculated myocardia and the epicardium. Scale bar = 500 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59724/59724fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59724/59724fig5.jpg" /> 
Figure 5: Exogenous IT-injected CNTF stimulates several biological processes in the heart. (A) Scheme of the experimental design. First, 2.5 &#181;L of a solution containing 250 ng of zebrafish CNTF or control immunoglobulins (hIgG) was injected into the pericardium of transgenic fish expressing nuclear DsRed2 in cardiomyocytes. Hearts were collected at 7 and 1 days post-injection (dpi) and analyzed by immunofluorescence and in situ hybridization, respectively. (B-D) Confocal microscopy images of ventricular sections of control and CNTF-injected hearts. (B) Immunostaining against a cell cycle marker, minichromosome maintenance complex component 5 (MCM5; green), reveals a higher number of proliferating cardiomyocytes in response to exogenous CNTF. Scale bar = 500 &#181;m. (C) Immunostaining against collagen XII shows increased deposition of collagen XII in the myocardium after CNTF injection. In control heart, collagen XII is confined to the epicardium34. Scale bar = 500 &#181;m. (D) Immunostaining against an immune cell marker, L-plastin, detects an enhanced recruitment of immune cells in the CNTF injected fish. Scale bar = 500 &#181;m. (E) Bright-field microscope images of ventricular cross sections after in situ hybridization using an antisense mRNA probe against cystatin, a cardioprotective factor, displays transcriptional upregulation of this gene in the heart of CNTF-injected fish. Scale bar = 500 &#181;m. This figure has been modified from Bise et al.24. <a href="https://www.jove.com/files/ftp_upload/59724/59724fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Intrathoracic Injection for the Study of Adult Zebrafish Heart at JoVE.com

Intrathoracic Injection for the Study of Adult Zebrafish Heart

This method relies on the injection of 0.5&#8722;3 &#956;L of solution into the thorax of adult zebrafish. The procedure efficiently delivers proteins and chemical compounds into the proximity of the zebrafish heart without damaging the organ. The approach is suitable for testing effects of exogenous factors on various tissues of the heart.

The adult zebrafish heart provides a powerful model in cardiac regeneration research. Although the strength of this system is based on transgenic ...

intrathoracic-injection-for-the-study-of-adult-zebrafish-heart

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12.2K Views.  University of Fribourg. This method relies on the injection of 0.5−3 μL of solution into the thorax of adult zebrafish. The procedure efficiently delivers proteins and chemical compounds into the proximity of the zebrafish heart without damaging the organ. The approach is suitable for testing effects of exogenous factors on various tissues of the heart.

Video: Intrathoracic Injection for the Study of Adult Zebrafish Heart

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 ...

Computer-Generated Animal Model Stimuli

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In particular, understanding the constraints on visual signal design for communication has been of great interest. Traditional methods for investigating animal interactions have used basic observational techniques, staged encounters, or physical manipulation of morphology. Less intrusive methods have tried to simulate conspecifics using crude playback tools, such as mirrors, still images, or models. As technology has become more advanced, video playback has emerged as another tool in which to examine visual communication (Rosenthal, 2000). However, to move one step further, the application of computer-animation now allows researchers to specifically isolate critical components necessary to elicit social responses from conspecifics, and manipulate these features to control interactions. Here, I provide detail on how to create an animation using the Jacky dragon as a model, but this process may be adaptable for other species. In building the animation, I elected to use Lightwave  3D to alter object morphology, add texture, install bones, and provide comparable weight shading that prevents exaggerated movement. The animation is then matched to select motor patterns to replicate critical movement features. Finally, the sequence must rendered into an individual clip for presentation. Although there are other adaptable techniques, this particular method had been demonstrated to be effective in eliciting both conspicuous and social responses in staged interactions.

Computer-generated stimuli using the Jacky dragon as a model.

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In ...

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 ...

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 ...

Retro-orbital Injection in Adult Zebrafish

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

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

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

Dissection of Organs from the Adult Zebrafish

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

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

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

Intraperitoneal Injection into Adult Zebrafish

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

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

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

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 ...

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work examining post-embryonic through adult cardiac development has been limited7-10. Examining the changing morphology of the maturing and aging heart are restricted by the lack of techniques available for staging and isolating juvenile and adult hearts. In order to analyze heart development over the fish's lifespan, we dissect zebrafish hearts at numerous stages and photograph them for further analysis11. The morphological features of the heart can easily be quantified and individual hearts can be further analyzed by a host of standard methods. Zebrafish grow at variable rates and maturation correlates better with fish size than age, thus, post-fixation, we photograph and measure fish length as a gauge of fish maturation. This protocol explains two distinct, size dependent dissection techniques for zebrafish, ranging from larvae 3.5mm standard length (SL) with hearts of 100&mu;m ventricle length (VL), to adults, with SL of 30mm and VL 1mm or larger. Larval and adult fish have quite distinct body and organ morphology. Larvae are not only significantly smaller, they have less pigment and each organ is visually very difficult to identify. For this reason, we use distinct dissection techniques.
We used pre-dissection fixation procedures, as we discovered that hearts dissected directly after euthanization have a more variable morphology, with very loose and balloon like atria compared with hearts removed following fixation. The fish fixed prior to dissection, retain in vivo morphology and chamber position (data not shown). In addition, for demonstration purposes, we take advantage of the heart (myocardial) specific GFP transgenic Tg(myl7:GFP)twu34 (12), which allows us to visualize the entire heart and is particularly useful at early stages in development when the cardiac morphology is less distinct from surrounding tissues. Dissection of the heart makes further analysis of the cell and molecular biology underlying heart development and maturation using in situ hybridization, immunohistochemistry, RNA extraction or other analytical methods easier in post-embryonic zebrafish. This protocol will provide a valuable technique for the study of cardiac development maturation and aging.

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

A clear, standardized method for dissection and isolation of the zebrafish heart at multiple developmental stages are described. Annotation and quantification techniques are also discussed.

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work ...