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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</ol>

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 mm 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>&lt;strong&gt;Dyes and Antibodies&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</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: 1/1,000</td></tr><tr><td>DAPI</td><td>Sigma</td><td>10236276001</td><td>Concentration: 1/2,000 (1&amp;micro;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>&lt;strong&gt;ISH probe primers&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Cystatin</td><td></td><td></td><td>gene number: ENSDARG00000074425&lt;br/&gt;fw primer: GATTCACTGTCGGGTTTGGG&lt;br/&gt;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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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

Nanyang Technological University

Research

JoVE Journal

Biology

12.4K Views.  University of Fribourg. In het hartonderzoek naar volwassen zebravissen worden medicamenteuze behandelingen meestal systemisch uitgevoerd. Met intrathoracische injectie kan men lokaal oplossing leveren aan het hart zonder myocardiale schade te veroorzaken. Deze methode maakt de levering van kleine hoeveelheden drugs, eiwitten, of andere moleculen in oplossing precies rond het hart van een volwassen zebravis.Gebruik om met deze procedure te beginnen een naaldtrekker om met microinjectie aangepaste borosilicaat...

Video: Intrathoracic Injection for the Study of Adult Zebrafish Heart

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 ...

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, ...

Optimized Intravenous Injection in Adult Zebrafish

Intravenous (IV) injection is widely recognized as the most effective and commonly utilized method for achieving systemic delivery of substances in mammalian research models. However, its application in adult zebrafish for drug delivery, stem cell transplantation, and regenerative and cancer studies has been limited due to the challenges posed by their small body size and intricate blood vessels. To overcome these limitations, alternative injection techniques such as intracardiac and retro-orbital (RO) injection have been explored in the past for stem cell transplantation in adult zebrafish. However, these techniques have their drawbacks, including the need for meticulous injection techniques or increased risk of mortality.
In this study, we have developed a refined and optimized IV injection procedure specifically tailored to adult zebrafish, addressing the challenges associated with their unique anatomy. To demonstrate the effectiveness of this technique, we performed successful IV injections of whole kidney marrow cells from Tg(mpo: EGFP) fish and FITC-dextran dye into adult Casper fish. The subsequent visualization of injected cells and dyes using a fluorescence microscope confirmed their successful delivery and engraftment within the zebrafish. Furthermore, we demonstrated that compared with the intracardiac and RO injections, the IV injection resulted in improved survival rates and engraftment efficiency in treated zebrafish. This approach enables precise delivery and localization of substances and holds great potential for large-scale drug and chemical screening using adult zebrafish. Additionally, the ability to visually track the injected cells and dyes provides invaluable insights into their engraftment, migration, and interactions with host tissues, enabling a more comprehensive evaluation of therapeutic effects and biological processes in zebrafish models.

Here, we present an optimized and effective protocol for the intravenous injection of cells or pharmacological agents into adult zebrafish, resulting in enhanced cell engraftment and increased survival rates of the treated zebrafish.

Intravenous (IV) injection is widely recognized as the most effective and commonly utilized method for achieving systemic delivery of substances in ...

Cancer Research

Intramyocardial Injection for the Study of Cardiac Lymphatic Function in Zebrafish

Zebrafish have proved to be an important model for studying cardiovascular formation and function during postembryonic development and regeneration. The present protocol describes a method for injecting fluorescent tracers into the zebrafish myocardium to study interstitial fluid and debris uptake into cardiac lymphatic vessels. To do so, microspheres (200 nm diameter) and quantum dots (&#60;10 nm diameter) are introduced into the myocardium of live zebrafish, which can be tracked using ex vivo confocal microscopy. These tracers are then tracked intermittently over several hours to follow clearance from the myocardium into cardiac lymphatic vessels. Quantum dots&#160;are transported through cardiac lymphatic vessels away from the heart, while larger microspheres&#160;remain at the injection site for over three weeks. This method of intramyocardial injection can be extended to other uses, including the injection of encapsulated MS or hydrogels to locally release cells, proteins, or compounds of interest to a targeted region of the heart.

The present protocol allows efficient and stable delivery of fluorescent microspheres (MS) and quantum dots (QD) into the myocardium of live fish that can be tracked (traced) over time.

Zebrafish have proved to be an important model for studying cardiovascular formation and function during postembryonic development and regeneration. The ...

Developmental Biology

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 ...

Analysis of Zebrafish Cardiac Contraction with Virtual Reality Interaction

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for gaining insights into structural and functional changes in response to injuries. However, obtaining high-resolution and high-speed 4-dimensional (4D, 3D spatial + 1D temporal) images of the zebrafish heart to assess cardiac architecture and contractility remains challenging. In this context, an in-house light-sheet microscope (LSM) and customized computational analysis are&#160;used to overcome these technical limitations. This strategy, involving LSM system construction, retrospective synchronization, single cell tracking, and user-directed analysis, enables one to investigate the micro-structure and contractile function across the entire heart at the single-cell resolution in the transgenic Tg(myl7:nucGFP) zebrafish larvae. Additionally, we are able to further incorporate microinjection of small molecule compounds to induce cardiac injury in a precise and controlled manner. Overall, this framework allows one to track physiological and pathophysiological changes, as well as the regional mechanics at the single-cell level during cardiac morphogenesis and regeneration.

Author Spotlight: High-Resolution 4D Light-Sheet Imaging and Virtual Reality in Zebrafish for Single-Cell Analysis of Heart Function

This protocol utilizes light-sheet imaging to investigate cardiac contractile function in zebrafish larvae and gain insights into cardiac mechanics through cell tracking and interactive analysis.

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for ...

Bioengineering

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

Optimized Intraperitoneal Injection for Compound Delivery in Adult Zebrafish

An Intraperitoneal Injection Technique in Adult Zebrafish that Minimizes Body Damage and Associated Mortality

The adult zebrafish (Danio rerio), which is genetically accessible, is being employed as a valuable vertebrate model to study human disorders such as cardiomyopathy. Intraperitoneal (IP) injection is an important method that delivers compounds to the body for either testing therapeutic effects or generating disease models such as doxorubicin-induced cardiomyopathy (DIC). Currently, there are two methods of IP injection. Both&#160;methods have limitations when handling toxic compounds such as doxorubicin, which result in side effects manifesting as severe damage to the body shape and fish death. While these shortcomings could be overcome by extensive investigator training, a new IP injection method that has minimal side effects is desirable. Here, a unique IP injection method that is able to handle toxic compounds is reported. Consistently reduced cardiac function can result without incurring significant fish death. The technique can be easily mastered by researchers who have minimal experience with adult zebrafish.

Author Spotlight: Novel Intraperitoneal Injection Method for Zebrafish Cardiotoxicity Models

A new intraperitoneal (IP) injection method in adult zebrafish is described. When handling toxic compounds such as doxorubicin, this procedure is more effective than the two previously reported IP methods. The technique is designed to be easily adopted by researchers with limited experience in the zebrafish model.

The adult zebrafish (Danio rerio), which is genetically accessible, is being employed as a valuable vertebrate model to study human disorders such as ...

Medicine

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid external development, tractable genetics, and translucency of the zebrafish, Danio rerio, LSFM has delivered insights into cardiac form and function at high spatial and temporal resolution without significant phototoxicity or photobleaching. Imaging of beating hearts challenges existing sample preparation and microscopy techniques. One needs to maintain a healthy sample in a constricted field of view and acquire ultrafast images to resolve the heartbeat. Here we describe optimized tools and solutions to study the zebrafish heart in vivo. We demonstrate the applications of bright transgenic lines for labeling the cardiac constituents and present novel gentle embedding and immobilization techniques that avoid developmental defects and changes in heart rate. We also propose a data acquisition and analysis pipeline adapted to cardiac imaging. The entire workflow presented here focuses on zebrafish embryonic heart imaging but can also be applied to various other samples and experiments.

We describe optimized tools to study the zebrafish heart in vivo with light sheet fluorescence microscopy. Specifically, we suggest bright cardiac transgenic lines and present new gentle embedding and immobilization techniques that avoid developmental and heart defects. A possible data acquisition and analysis pipeline adapted to cardiac imaging is also provided.

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid ...

Intrathoracic Injection: Compound Delivery to the Adult Zebrafish Heart

Source: Bise, T., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/59724/" target="_blank">Intrathoracic Injection for the Study of Adult Zebrafish Heart</a>. J. Vis. Exp. (2019).
This video describes the technique of delivering compounds into the heart of adult zebrafish by using intrathoracic injection.

Source: Bise, T., et al. Intrathoracic Injection for the Study of Adult Zebrafish Heart.  J. Vis. Exp. (2019).
This video describes the technique of ...

Intrathoracic Injection for the Study of Adult Zebrafish Heart

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