This work was supported in part by a Scientist Development Grant from the American Heart Association (14SDG18160021 to YD), the US NIH R01 grants HL 81753 and HL 107304 to XX, and the Mayo Foundation to XX.

All procedures described here were performed in accordance with the Guide for the Care and Use of Laboratory Animals (National Academies Press. 2011), and they were approved by the Mayo Clinic Institutional Animal Care and Use Committee.
1. Adult Zebrafish Preparation
<ol>
	<li>Set up sufficient breeding pairs in crossing tanks to acquire at least twice as many as the total fish needed for DOX injection. If comparing fish with different genetic backgrounds, breed all fish within the same week to ensure age-matched controls.</li>
	<li>Collect the fish embryos the next morning, transfer them to 100 mm Petri dishes, and keep them in a 28.5 &#176;C incubator. Keep embryos at a low density (&#60;100 embryos/Petri dish).</li>
	<li>Refresh the embryo water daily to avoid sex imbalance, and manually remove dead eggs in a timely fashion using a transfer pipette.</li>
	<li>Put the same number of embryos in each tank (for example, 60 embryos/3 L medium tank initially) to ensure density-matched controls.</li>
	<li>Start paramecia feeding at 4 days post-fertilization (dpf).</li>
	<li>Inspect the fish daily during the juvenile stage. Adjust the fish number as needed to ensure similar fish density.</li>
	<li>When fish reach 4 weeks of age, transfer up to 20 fish into each new 3 L medium tank for further growth. Start to feed the fish with live hatched brine shrimp.</li>
</ol>
2. Preparation and Storage of DOX Stock Solution
NOTE: DOX can be purchased from various bio-companies. The compound is usually acquired as a powder in dark brown containers.
<ol>
	<li>Thoroughly dissolve the DOX powder in de-ionized water to ensure no clumps are visible, with a final concentration of 5 mg/mL as the stock solution. Aliquot 1 mL of DOX stock into each 1.5 mL safe-lock tube. Wrap the 1.5 mL tubes with aluminum foil paper to protect the DOX from light exposure. 
	NOTE: Perform this step in a chemical hood.</li>
	<li>Keep the DOX stock solution at 4 &#176;C for storage. For long-term storage (&#62;4 weeks) of DOX stock solution, perform the optional Section 3 described below.</li>
</ol>
3. Quality Control of DOX Using Zebrafish Embryos (Optional)
NOTE: DOX is both moisture- and light-sensitive, so it can lose its drug efficacy for modeling DIC after extended storage. For DOX purchased from different companies, or even different batches from the same company, it is useful to calibrate its drug efficacy using wild-type (WT) zebrafish embryos before conducting the experiments on adult fish. This method is derived from a reported zebrafish embryonic DIC model21.
<ol>
	<li>Collect WT zebrafish embryos from at least 2 pairs of fish. Dechorionate embryos at 24 h post-fertilization (hpf) manually by using a syringe with a micro needle. Alternatively, treat the embryo with proteinase K at final concentration of 10 &#181;g/mL for 10-15 min in a 30 &#176;C incubator. Refresh the embryo water after dechorionation. Remove dead embryos and maintain at least 36 embryos of each batch.</li>
	<li>Dilute the DOX stock solution in fresh embryo water to a final concentration of 100 &#181;M. The volume of solution is 100 &#181;L for every 3 embryos. Mix the diluted DOX solution by vortex. The final diluted solution should be a light, red color.</li>
	<li>Add 100 &#181;L/well of diluted DOX solution of a clean 96-well transparent plate.</li>
	<li>Take 3 dechorionated embryos with a plastic transfer pipette, and keep the embryos close to the end of the pipette tip. Put the pipette tip into each well with DOX solution. Let the end of the tip touch the solution and allow the embryos to swim into the well. 
	NOTE: Avoid manually pushing the embryos, which will add more water into the well and dilute the DOX solution.</li>
	<li>Refresh the DOX solution at 48 hpf. By this time, observe the wells under a microscope with 10X magnification to identify dead embryos (cessation of heartbeat) or embryos with edema. Count and remove any dead embryos in a timely fashion, otherwise the remaining embryos exposed to the solution with dead embryos can die quickly as well.</li>
	<li>Check the embryos at 72 hpf and count them. The DOX treatment is considered &#34;good drug efficacy&#34; if &#62;25% death (cessation of heartbeat) can be observed in both batches of embryos.</li>
</ol>
4. Pre-injection Preparation
NOTE: Fish of 8 weeks to 6 months of age are used for DOX injection. The body weights (BWs) of a matured Wild Indian Karyotype (WIK) fish to be injected can range from 0.2-0.5 g.
<ol>
	<li>Fast the fish for 24 h before injection.</li>
	<li>Anesthetize the fish with embryo water containing 0.16 mg/mL tricaine. Use a clean filter paper to dry the water from both sides of the body. Measure the BW of each fish on a scale. Group fish within 10% difference in BW together for later injection. 
	NOTE: To minimize the workload at this step, fish within 10% difference in BW are considered the same size; therefore, prepare one DOX working solution according to their mean BW.</li>
	<li>Plan to inject each adult fish with 5 &#181;L of solution. Calculate the DOX working concentration according to the fish numbers and BWs. 
	NOTE: To study chronic cardiotoxicity up to 6 months, use DOX at a dose of 20 mg/kg. To study acute cardiotoxicity of DOX, the DOX dose can be increased up to 50 mg/kg of BW.</li>
	<li>Dilute the DOX stock in 1x Hank&#39;s balanced salt solution (HBSS) for corresponding working concentrations. Vortex to mix the solution. Briefly spin down to collect the solution.</li>
</ol>
5. DOX Injection in Adult Fish
<ol>
	<li>Place a clean 100 mm Petri dish with a sponge inside it, under a dissection microscope, then adjust the focus. Cut the sponge to make a cavity of about 4 cm in length to hold a fish. Make a longer cavity for a larger fish.</li>
	<li>Prepare a 34 G needle with a 10 &#181;L micro-syringe. Rinse the needle with 1x HBSS buffer to remove any bubbles and blocks from the syringe and tubing.</li>
	<li>Anesthetize the adult fish in embryo water containing 0.16 mg/mL tricaine for 2 min. 
	NOTE: Prolonged anesthetization over 5 min followed by DOX injection can easily cause fish death.</li>
	<li>Soak the sponge in embryo water with tricaine, and transfer the fish onto the sponge for injection.</li>
	<li>Perform IP DOX injection by either of the two methods described below.
	<ol>
		<li>Classic IP injection29
		<ol>
			<li>Position the fish with the abdomen up in the cavity of the sponge. Quickly insert the needle, with a 45&#176; angle to the fish body into the midline between the pelvic fins, and penetrate approximately 1-2 mm. Release all DOX solution slowly. Wait 5 s before pulling out the needle. Check the DOX delivery by a visible red color in the fish belly.</li>
		</ol>
		</li>
		<li>Alternative IP injection
		<ol>
			<li>Place the fish laterally onto the sponge with the anterior to the right. Gently stabilize the fish using a blunt end forceps with the left hand, and hold the micro-syringe with the right hand.</li>
			<li>Position the needle below the lateral line above the pelvic fin, with the bevel facing up. Pointing at the 7 o&#39;clock position at a 45&#176; angle, insert the needle 3-4 mm to the fish cavity located between the pelvic and the anal fins, and then slowly depress the plunger. Check the DOX delivery by a visible red color in the fish belly.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Quickly transfer the injected fish to a clean crossing tank filled with fresh system water to allow the fish to recover. Rinse the needle once with 1x HBSS buffer between injections.</li>
</ol>
6. Post-injection Fish Management
<ol>
	<li>After injection, return the fish to the system with running circulation. If possible, maintain DOX-treated fish separately from the main system to avoid cross-contamination among different tanks that share the circulation.</li>
	<li>Fast the injected fish for another 24 h for recovery. Observe the fish daily during the first week. Remove the dead fish in a timely fashion to avoid infection to the other fish. 
	NOTE: Fish deaths within the first 24 h are likely due to physical lesions caused by the injection.</li>
	<li>Further maintain the DOX-stressed fish for longitudinal observations. Remove dead fish in time to avoid infections to other fish in the tank. 
	NOTE: Fish numbers are documented to generate a survival curve.</li>
	<li>Use different experimental assays to phenotype the DOX-stressed fish, such as echocardiography30, cardiac function reporter transgenic line23, swimming challenge26, and quantification of other pathological remodeling markers23.</li>
</ol>

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(34), (2009).</li><li>Zang, L., Morikane, D., Shimada, Y., Tanaka, T., Nishimura, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+protocol+for+the+oral+administration+of+test+chemicals+to+adult+zebrafish.">A novel protocol for the oral administration of test chemicals to adult zebrafish.</a> Zebrafish. 8 (4), 203-210 (2011).</li><li>Collymore, C., Rasmussen, S., Tolwani, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gavaging+adult+zebrafish.">Gavaging adult zebrafish.</a> J Vis Exp. (78), (2013).</li></ol>

The authors declare no conflicts of interests.

To model a progressive DIC, the dose of 20 mg/kg DOX was determined experimentally as the highest dose that does not cause significant fish death during 1 wpi but still results in fish death and the reduction of cardiac function after 4 wpi (Figure 3 and Figure 4C). This dose is comparable to those frequently used in rodent DIC models (15-25 mg/kg) and to the limit cumulative dose in humans (550 mg/m2, which is equivalent to 15 mg/kg)<sup class="xref"...

Doxorubicin (DOX), also named Adriamycin, has been developed as an anti-neoplastic drug since the 1960s1,2. It is now still actively used as an important chemotherapeutic drug for a broad spectrum of tumors. However, clinical application of DOX was hampered by its dose-dependent toxicities, especially cardiotoxicity characterized by variable symptoms ranging from asymptomatic electrocardiographic changes to pericarditis and decompensated cardiomyopathy1,2. To date, at least three major hypotheses have been raised to explain DIC, including activated reactive oxygen species (ROS)1,3,4,5, inhibition of topoisomerase II-&#946; (TOP2&#946;)6,7, and modulation of intracellular calcium release1,8,9. Accumulating evidence also suggests genetic predisposition as a pivotal risk factor for DIC10,11,12,13. Gene identities related to these DIC predispositions, however, remain largely unknown. Dexrazoxane is the only adjuvant agent approved by the US Food and Drug Administration (FDA) to treat DIC, but with limited implementation14,15,16, underscoring the need to identify additional therapeutic strategies. Animal models of DIC are therefore explored for these purposes. Owing to their accessibility and simplicity, mechanistic studies on DIC models could potentially have broader impacts on other types of cardiomyopathies: common pathogenesis might be shared among cardiomyopathies of different etiologies, especially at later pathological stages17,18,19,20.
In addition to rodent models of DIC, zebrafish DIC models with higher throughput have been developed to facilitate the discovery of both new genetic factors and therapeutics. An embryonic DIC model has been established in the transparent zebrafish embryos for screening therapeutic compounds21. Given that the cardiomyopathies are adult onset diseases with a progressive pathogenesis, adult zebrafish cardiomyopathy models have been developed22,23,24,25,26. We generated the first acquired model for cardiomyopathy resulting from chronic anemia24, followed by DIC as the second acquired cardiomyopathy model in adult zebrafish23. We found that injection of a single bolus of DOX into adult zebrafish induces cardiotoxicity that consists of an acute phase roughly within 1 week post-injection (wpi), followed by a chronic phase of cardiomyopathy up to 6 months post-injection. While haploinsufficiency of the mechanistic target of rapamycin(mtor) ameliorates cardiomyopathy at the chronic phase, it exaggerates fish mortality at the acute phase, underscoring the value of the adult DIC model to discern stage-dependent mechanisms23. We further demonstrated that the adult DIC model can be used to stress a collection of zebrafish insertional cardiac (ZIC) mutants that are being generated via a transposon-based insertional mutagenesis approach27. A pilot screen identified 3 known cardiomyopathy genes as well as DnaJ (Hsp40) homolog, subfamily B, and member 6b (dnajb6b) as new DIC susceptibility genes28. Therefore, the generation of the adult DIC model in zebrafish led to a new methodology that systematically enables identification of genetic modifiers for DIC, which complements the existing genome-wide association study (GWAS) and quantitative trait locus (QTL) analysis.
During the generation and implementation of the adult zebrafish DIC model, we noticed significant variations among different researchers and/or even among different injections performed by the same investigator. The longitudinal nature of the model imposes challenges to registering results from different investigators and to the sequential troubleshooting process. To facilitate the use of this simple cardiomyopathy-inducing stress method by the research community, we describe our protocol in detail, present two types of IP injection, and discuss considerations to reduce variations among different researchers.

<table><tbody><tr><td>Crossing tank</td><td>Aquaneering</td><td>ZHCT100</td><td>Fish breeding</td></tr><tr><td>Incubator</td><td>ThermoFisher</td><td></td><td>Maintaining embryo</td></tr><tr><td>3 L medium tank</td><td>Aquaneering</td><td>ZT280</td><td>Maintaining fish</td></tr><tr><td>Paramecia</td><td>Carolina</td><td>131560</td><td>Food for juvenile fish</td></tr><tr><td>Live hatched brine shrimp</td><td>in house</td><td></td><td>Food for adult fish</td></tr><tr><td>Doxorubicin hydrochloride</td><td>Sigma</td><td>D1515-10MG</td><td></td></tr><tr><td>1.5 ml safe-lock tube</td><td>Eppendorf</td><td>No. 022363204</td><td>For drug storage</td></tr><tr><td>Aluminum foil paper</td><td>Fisher</td><td>1213104</td><td>For preventing light exposure</td></tr><tr><td>Proteinase K</td><td>Roche</td><td>No. 03115887001</td><td>For dechorionating embryo</td></tr><tr><td>Hank&#039;s balanced salt solution (HBBS)</td><td>ThermoFisher</td><td>14025076</td><td>Vehicle for DOX</td></tr><tr><td>100 mm petri dish</td><td>Falcon</td><td>431741</td><td></td></tr><tr><td>10 &amp;mu;L NanoFil micro-syringe</td><td>WPI</td><td>NANOFIL</td><td>For injection</td></tr><tr><td>34 gauge needle</td><td>WPI</td><td>NF34BV-2</td><td>For injection</td></tr><tr><td>Tricaine</td><td>Argent</td><td>MS-222</td><td>Anesthetizing fish</td></tr><tr><td>96 well plate</td><td>Costar</td><td>3539</td><td>For embryo drug treatment</td></tr><tr><td>Transfer pipette</td><td>Bel-art product</td><td>F37898</td><td>For transfering embryo</td></tr></tbody></table>

a doxorubicin-induced cardiomyopathy model in adult zebrafish

The genetically accessible adult zebrafish (Danio rerio) has been increasingly used as a vertebrate model for understanding human diseases such as cardiomyopathy. Because of its convenience and amenability to high throughput genetic manipulations, the generation of acquired cardiomyopathy models, such as the doxorubicin-induced cardiomyopathy (DIC) model in adult zebrafish, is opening the doors to new research avenues, including discovering cardiomyopathy modifiers via forward genetic screening. Different from the embryonic zebrafish DIC model, both initial acute and later chronic phases of cardiomyopathy can be determined in the adult zebrafish DIC model, enabling the study of stage-dependent signaling mechanisms and therapeutic strategies. However, variable results can be obtained with the current model, even in the hands of experienced investigators. To facilitate future implementation of the DIC model, we present a detailed protocol on how to generate this DIC model in adult zebrafish and describe two alternative ways of intraperitoneal (IP) injection. We further discuss options on how to reduce variations to obtain reliable results and provide suggestions on how to appropriately interpret the results.

Here, two methods to perform IP injection to model DIC in adult zebrafish are presented. While using the classic, established IP injection method29, it was noted that the injected DOX solution (red color) could sometimes ooze out from the location where the needle penetrated. The alternative IP injection uses a different location for needle penetration that is 3-4 mm away from the peritoneum where the DOX is released (Figure 1A), which...

Watch this Scientific Journal Video about A Doxorubicin-induced Cardiomyopathy Model in Adult Zebrafish at JoVE.com

A Doxorubicin-induced Cardiomyopathy Model in Adult Zebrafish

A method to generate a doxorubicin-induced cardiomyopathy model in adult zebrafish (Danio rerio) is described here. Two alternative ways of intraperitoneal injection are presented and conditions to reduce variations among different experimental groups are discussed.

The genetically accessible adult zebrafish (Danio rerio) has been increasingly used as a vertebrate model for understanding human diseases such as ...

a-doxorubicin-induced-cardiomyopathy-model-in-adult-zebrafish

Research

JoVE Journal

Medicine

9.7K Views.  Mayo Clinic Graduate School of Biomedical Sciences. Een methode voor het genereren van een Doxorubicine-geïnduceerde cardiomyopathie model in volwassen zebravissen (Danio rerio) wordt hier beschreven. Twee alternatieve manieren van intraperitoneale injectie worden gepresenteerd en voorwaarden om de variaties tussen verschillende experimentele groepen worden besproken.

Video: A Doxorubicin-induced Cardiomyopathy Model in Adult Zebrafish

Induction of Myocardial Infarction in Adult Zebrafish Using Cryoinjury

The mammalian heart is incapable of significant regeneration following an acute injury such as myocardial infarction1. By contrast, urodele amphibians and teleost fish retain a remarkable capacity for cardiac regeneration with little or no scarring throughout life2,3. It is not known why only some non-mammalian vertebrates can recreate a complete organ from remnant tissues4,5. To understand the molecular and cellular differences between regenerative responses in different species, we need to use similar approaches for inducing acute injuries.
In mammals, the most frequently used model to study cardiac repair has been acute ischemia after a ligation of the coronary artery or tissue destruction after cryoinjury6,7. The cardiac regeneration in newts and zebrafish has been predominantly studied after a partial resection of the ventricular apex2,3. Recently, several groups have established the cryoinjury technique in adult zebrafish8-10. This method has a great potential because it allows a comparative discussion of the results obtained from the mammalian and non-mammalian species.
Here, we present a method to induce a reproducible disc-shaped infarct of the zebrafish ventricle by cryoinjury. This injury model is based on rapid freezing-thawing tissue, which results in massive cell death of about 20% of cardiomyocytes of the ventricular wall. First, a small incision was made through the chest with iridectomy scissors to access the heart. The ventricular wall was directly frozen by applying for 23-25 seconds a stainless steel cryoprobe precooled in liquid nitrogen. To stop the freezing of the heart, fish water at room temperature was dropped on the tip of the cryoprobe. The procedure is well tolerated by animals, with a survival rate of 95%.
To characterize the regenerative process, the hearts were collected and fixed at different days after cryoinjury. Subsequently, the specimen were embedded for cryosectioning. The slides with sections were processed for histological analysis, in situ hybridization and immunofluorescence. This undertaking enhances our understanding of the factors that are required for the regenerative plasticity in the zebrafish, and provide new insights into the machinery of cardiac regeneration. A conceptual and molecular understanding of heart regeneration in zebrafish will impact both developmental biology and regenerative medicine.

Zebrafish represents a valuable model to study the mechanisms of heart regeneration in vertebrates. Here, we present a protocol for induction of a heart infarct in adult zebrafish using cryoinjury. This method results in massive cell death within 20% of the ventricular wall, similar to that observed in mammalian infarcts.

The mammalian heart is incapable of significant regeneration following an acute injury such as myocardial infarction1. By contrast, urodele amphibians and ...

Displacement Analysis of Myocardial Mechanical Deformation (DIAMOND) Reveals Segmental Heterogeneity of Cardiac Function in Embryonic Zebrafish

Zebrafish are increasingly utilized as a model organism for cardiomyopathies and regeneration. Current methods evaluating cardiac function fail to reliably detect segmental mechanics and are not readily feasible in zebrafish. Here we present a semiautomated, open-source method for the quantitative assessment of four-dimensional (4D) segmental cardiac function: displacement analysis of myocardial mechanical deformation (DIAMOND). Transgenic embryonic zebrafish were imaged in vivo using a light-sheet fluorescence microscopy system with 4D cardiac motion synchronization. Acquired 3D digital hearts were reconstructed at end-systole and end-diastole, and the ventricle was manually segmented into binary datasets. Then, the heart was reoriented and isotropically resampled along the true short axis, and the ventricle was evenly divided into eight portions (I&#8211;VIII) along the short axis. Due to the different resampling planes and matrices at end-systole and end-diastole, a transformation matrix was applied for image registration to restore the original spatial relationship between the resampled systolic and diastolic image matrices. After image registration, the displacement vector of each segment from end-systole to end-diastole was calculated based on the displacement of mass centroids in three dimensions (3D). DIAMOND shows that basal myocardial segments adjacent to the atrioventricular canal undergo the highest mechanical deformation and are the most susceptible to doxorubicin-induced cardiac injury. Overall, DIAMOND provides novel insights into segmental cardiac mechanics in zebrafish embryos beyond traditional ejection fraction (EF) under both physiological and pathological conditions.

Displacement Analysis of Myocardial Mechanical Deformation DIAMOND Reveals Segmental Heterogeneity of Cardiac Function in Embryonic Zebrafish

The goal of this protocol is to detail a novel method for the assessment of segmental cardiac function in embryonic zebrafish under both physiological and pathological conditions.

Zebrafish are increasingly utilized as a model organism for cardiomyopathies and regeneration. Current methods evaluating cardiac function fail to ...

Developmental Biology

High-Frequency Ultrasound Echocardiography to Assess Zebrafish Cardiac Function

The zebrafish (Danio rerio) has become a very popular model organism in cardiovascular research, including human cardiac diseases, largely due to its embryonic transparency, genetic tractability, and amenity to rapid, high-throughput studies. However, the loss of transparency limits heart function analysis at the adult stage, which complicates modeling of age-related heart conditions. To overcome such limitations, high-frequency ultrasound echocardiography in zebrafish is emerging as a viable option. Here, we present a detailed protocol to assess cardiac function in adult zebrafish by non-invasive echocardiography using high-frequency ultrasound. The method allows visualization and analysis of zebrafish heart dimension and quantification of important functional parameters, including heart rate, stroke volume, cardiac output, and ejection fraction. In this method, the fish are anesthetized and kept underwater and can be recovered after the procedure. Although high-frequency ultrasound is an expensive technology, the same imaging platform can be used for different species (e.g., murine and zebrafish) by adapting different transducers. Zebrafish echocardiography is a robust method for cardiac phenotyping, useful in the validation and characterization of disease models, particularly late-onset diseases; drug screens; and studies of heart injury, recovery, and regenerative capacity.

We describe a protocol to assess heart morphology and function in adult zebrafish using high-frequency echocardiography. The method allows visualization of the heart and subsequent quantification of functional parameters, such as heart rate (HR), cardiac output (CO), fractional area change (FAC), ejection fraction (EF), and blood inflow and outflow velocities.

The zebrafish (Danio rerio) has become a very popular model organism in cardiovascular research, including human cardiac diseases, largely due to its ...

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 ...

Biology

Immunostaining of Dissected Zebrafish Embryonic Heart 

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology and genetics 1,2. About 100-200 embryos are readily available every week from a single pair of adult fish. The transparent embryos that develop ex utero make them ideal for assessing cardiac defects 3. The expression of any gene can be manipulated via morpholino technology or RNA injection 4. Moreover, forward genetic screens have already generated a list of mutants that affect different perspectives of cardiogenesis 5. 
Whole mount immunostaining is an important technique in this animal model to reveal the expression pattern of the targeted protein to a particular tissue 6. However, high resolution images that can reveal cellular or subcellular structures have been difficult, mainly due to the physical location of the heart and the poor penetration of the antibodies. 
 Here, we present a method to address these bottlenecks by dissecting heart first and then conducting the staining process on the surface of a microscope slide. To prevent the loss of small heart samples and to facilitate solution handling, we restricted the heart samples within a circle on the surface of the microscope slides drawn by an immEdge pen. After the staining, the fluorescence signals can be directly observed by a compound microscope. 
Our new method significantly improves the penetration for antibodies, since a heart from an embryonic fish only consists of few cell layers. High quality images from intact hearts can be obtained within a much reduced procession time for zebrafish embryos aged from day 2 to day 6. Our method can be potentially extended to stain other organs dissected from either zebrafish or other small animals.

Immunostaining of Dissected Zebrafish Embryonic Heart

A rapid way to conduct immunostaining of zebrafish embryonic heart is described. Compared to the whole mount immunostaining approach, this method dramatically increases the penetration of the antibodies, which allows obtaining high resolution images that reveal cellular/subcellular structures in the heart within a much reduced processing time.

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology ...

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 ...

In Vivo Surface Electrocardiography for Adult Zebrafish

The electrocardiogram waveforms of adult zebrafish and those of humans are remarkably similar. These electrocardiogram similarities enhance the value of zebrafish not only as a research model for human cardiac electrophysiology and myopathies but also as a surrogate model in high throughput pharmaceutical screening for potential cardiotoxicities to humans, such as QT prolongation. As such, in vivo electrocardiography for adult zebrafish is an electrical phenotyping tool that is necessary, if not indispensable, for cross-sectional or longitudinal in vivo electrophysiological characterizations. However, too often, the lack of a reliable, practical, and cost-effective recording method remains a major challenge preventing this in vivo diagnostic tool from becoming more readily accessible. Here, we describe a practical, straightforward approach to in vivo electrocardiography for adult zebrafish using a low-maintenance, cost-effective, and comprehensive system that yields consistent, reliable recordings. We illustrate our protocol using healthy adult male zebrafish of 12-18 months of age. We also introduce a rapid real-time interpretation strategy for quality validation to ensure data accuracy and robustness early in the electrocardiogram recording process.

Here, we present a reliable, minimally invasive, and cost-effective method to record and interpret electrocardiograms in live anesthetized adult zebrafish.

The electrocardiogram waveforms of adult zebrafish and those of humans are remarkably similar. These electrocardiogram similarities enhance the value of ...

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 ...

Isolation of Cardiac and Vascular Smooth Muscle Cells from Adult, Juvenile, Larval and Embryonic Zebrafish for Electrophysiological Studies

Zebrafish have long been used as a model vertebrate organism in cardiovascular research. The technical difficulties of isolating individual cells from the zebrafish cardiovascular tissues have been limiting in studying their electrophysiological properties. Previous methods have been described for dissection of zebrafish hearts and isolation of ventricular cardiac myocytes. However, the isolation of zebrafish atrial and vascular myocytes for electrophysiological characterization was not detailed. This work describes new and modified enzymatic protocols that routinely provide isolated juvenile and adult zebrafish ventricular and atrial cardiomyocytes, as well as vascular smooth muscle (VSM) cells from the bulbous arteriosus, suitable for patch-clamp experiments. There has been no literary evidence of electrophysiological studies on zebrafish cardiovascular tissues isolated at embryonic and larval stages of development. Partial dissociation techniques that allow patch-clamp experiments on individual cells from larval and embryonic hearts are demonstrated.

The present protocol describes the acute isolation of viable cardiac and vascular smooth muscle cells from adult, juvenile, larval, and embryonic zebrafish (Danio rerio), suitable for electrophysiological studies.

Zebrafish have long been used as a model vertebrate organism in cardiovascular research. The technical difficulties of isolating individual cells from the ...

Heart Dissection: A Method to Observe Zebrafish Cardiac Development

Source: Singleman C. et. al., <a href="https://www.jove.com/v/3165/" target="_blank">Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio</a>. J. Vis. Exp. (2011).
This video describes a technique of isolating the heart from a zebrafish at multiple developmental stages to study cardiac development, maturation, and ageing.

Source: Singleman C. et. al., Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio. J. Vis. Exp. (2011).
This video describes a technique ...

A Doxorubicin-induced Cardiomyopathy Model in Adult Zebrafish

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