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)4,7,31,32,33. Higher doses of DOX, such as 50 mg/kg, show significant fish death during 1 wpi, so they can be used to study only acute cardiac toxic responses to DOX23,28.
During the implementation of this DIC model, we noticed that it was difficult for new investigators to reproduce the same results initially. Even for experienced investigators, about a 20% variation of mortality in the chronic phase can still be found among different injections (Figure 3), indicating uncharacterized biological confounding factors in the current model. Despite a less than perfect model, we still believe that this current DIC model is sufficient to make sound discovery because of the following evidence. First, after several practices, the results from the DIC model can be later registered among most, if not all, investigators. Second, based on this DIC model, we identified 4 meaningful genetic modifiers. Existing evidence from literature reports supported 3 of them as cardiomyopathy genes34,35,36,37. The 4th one is DnaJ homolog subfamily B member 6 (DNAJB6), which has been proven to be a new cardiomyopathy gene supported by a human genetic study28. Thus, we conclude that results based on intra-experimental groups using stringently age-matched controls are still highly repeatable, albeit the DIC model in its current form is not reliable for inter-experimental comparison.
The confounding factors that contribute to the inconsistency observed in our current DIC model likely include the following: (1) Aging and gender-difference have been regarded as crucial risk factors for cardiovascular diseases including DIC38,39,40. While both hypothesized cofounders remain to be tested in our model specifically, we did notice that aged fish tend to be more sensitive to DOX toxicity (data not shown). The inter-batch variations observed using both IP methods (Figure 3) are also likely contributed to these two cofounders. (2) Different from larger animal models, the size of an adult zebrafish is small. Thus, local damage caused by DOX injection can be more severe and variable. (3) Death of injected fish might result from DOX toxicity in other organs (e.g., renal toxicity), in addition to the heart. While more carefully designed experiments are needed to address each of the aforementioned confounding factors, previous work suggests that the following cautions will help reduce phenotypic variations: First, it is critical to guarantee the efficacy of DOX. DOX powder should always be kept in a dry and dark area, and solutions should be handled with care to reduce exposure to light. It is recommended that DOX working solutions be prepared freshly each time before fish injection. We typically do not use stock DOX solutions after 4 weeks in storage. When there is a doubt, perform the optional Section 3 in this protocol using the quick embryonic DIC model to calibrate the drug efficacy of each DOX batch. Second, adult fish synchronization is pivotal. The same fish strain needs to be raised at the same density, to ensure similar body size. We then pre-select fish with similar BWs for dose calculation. It is recommended to maintain at least twice as many fish in total for this pre-selection process. All fish are fasted for 24 h before the pre-selection process. Third, it is recommended to always using fish of similar age, and to sex the fish before DOX injection because of their different growth rates and potentially different cardiac responses to DOX.
We noticed that the majority of inconsistent conclusions could be ascribed to a questionable control group. Therefore, we recommend investigators new to the DIC model to practice DOX injections before conducting real experiments. When 20 mg/kg DOX is injected using the alternative IP injection method, a good injection technique can be indicated by a nearly zero fish death during the 1 wpi of DOX and a relatively consistent mortality rate at 2-3 months post-DOX injection. The ultimate evidence for successful modeling is the reduction of the cardiac-functional index, which could be quantified via echocardiography25,30, by using a casper; Tg(cmlc2:nusDsRed) transgenic line, or by using a newly developed ex vivo-based cardiac function assay (data not shown).
In addition to the IP injection, other drug delivery routes such as retro-orbital injection41, oral feeding42, and water incubation are also frequently used in adult zebrafish. We did not adopt the retro-orbital injection approach, despite its direct drug releasing into the circulation system, due to the lack of a method to validate successful drug delivery, as the redness of DOX can be easily masked by potential bleeding. We tried an oral delivery protocol through embedding DOX with absorbable gluten42, which was mixed with meals to feed adult fish. Unfortunately, with up to the accumulative dose of 150 mg/kg used within a 4-week feeding period, we did not observe any severe cardiotoxicity, suggesting the ineffectiveness of oral DOX delivery. Alternatively, an oral gavage technique could be further explored43. Moreover, the incubation protocol of soaking fish in a solution containing DOX is also a potential delivery route that could be tested in the future.
We acknowledge that one of the major limitations of the current DIC model is the single bolus injection method, which could result in high toxicity and local damage to the internal organs. This approach was designed to reduce workloads and to increase throughput, so that genetic screening can be conducted at a high throughput in adult fish28. In the future, models with multiple injections of DOX at lower doses should be pursued, which will better recapitulate the DIC observed in cancer patients treated with chemotherapy.

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&#39;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 &#956;L NanoFil micro-syringe</td>
			<td>WPI</td>
			<td>NANOFIL</td>
			<td>For injection</td>
		</tr>
		<tr>
			<td>34 gauge needle&#160;</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 effectively prevents the leakage (Figure 1B, 1C). Successful delivery of DOX into the peritoneum for both methods is evidenced by rapid distribution of the red color throughout the fish belly, which is visible on the opposite side of the injection locus.
Injection of DOX dosed at 50 mg/kg using the alternative IP method leads to severe toxicity, where the majority of fish die within one week (Figure 2). By contrast, injection of DOX dosed at 20 mg/kg using the alternative IP method causes almost no fish death during the first 2 weeks and ~10% fish death at 4 wpi (Figure 3A). Injection of DOX dosed at 20 mg/kg using the classic IP method exhibits ~30% fish death at 4 wpi (Figure 3C). Fish injected with either method exhibit a similar ~20% fish death from 4 wpi to 10 wpi (Figure 3B, D).
We have leveraged the casper; Tg(cmlc2:nusDsRed) fish to assess the progression of cardiac dysfunction in the DIC model (Figure 4A)23. The transparent body enables documentation of a red heart at both systolic (Figure 4B) and diastolic (Figure 4C) stages under a florescent microscope. After injection of 20 mg/kg DOX using the alternative IP method, ventricular function decline can be detected starting from 4 wpi (Figure 4D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57567/57567fig1.jpg" /> 
Figure 1: Injection routes. (1A) Schematics of the classic IP injection (b) and alternative IP injection (a) methods. The red circle indicates the common DOX releasing site for the two injection methods. Green circles indicate the needle penetration sites. The distances are estimated based on an adult WIK fish weighing approximately 0.3 g. (1B, 1C) Representative results indicating a successful DOX delivery using two intraperitoneal injection methods. Redness inside the adult fish belly can be noted immediately after the injection. Injected fish were checked again after 5 min recovery in a fresh system water. HBSS: 1x Hank&#39;s balanced salt solution. WT WIK zebrafish were employed. Scale bar: 5 mm. <a href="/files/ftp_upload/57567/57567fig1large.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/57567/57567fig2.jpg" /> 
Figure 2: Representative survival curves of adult zebrafish following DOX stress with 50 mg/kg DOX injection. Shown are 3 sets of DOX injections in different batches of WT adult fish at 3-6 months of age. In total, n = 24 fish were employed in the 1x HBSS control group, and n = 8 fish were employed in each batch injected with DOX. No difference in fish survivals were noted between the two injection methods. WT WIK zebrafish were employed. <a href="/files/ftp_upload/57567/57567fig2large.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/57567/57567fig3.jpg" /> 
Figure 3: Representative survival curves of adult zebrafish following DOX stress with 20 mg/kg DOX injection and the comparison between two intraperitoneal injection methods. (3A) Fish survival within 0-4 weeks following DOX delivery by classic intraperitoneal injection. Percent fish number at week 0 was regarded as 100%. (3B) Fish survival after 4 weeks post DOX delivery by classic intraperitoneal injection. Percent fish number at week 4 was regarded as 100%. (3C) Fish survival within 0-4 weeks following DOX delivery by alternative intraperitoneal injection. Percent fish number at week 0 was regarded as 100%. (3D) Fish survival after 4 weeks following DOX delivery by alternative intraperitoneal injection. Percent fish number at week 4 was regarded as 100%. Data shown in (3A) and (3B) are 9 different batches of DOX injection from a total 223 injected fish at 3-6 months of age. Data shown in (3C) and (3D) are 14 different batches of DOX injection from a total 335 injected fish at 2-6 months of age. The numbers of live fish are recorded weekly. Error bars represent standard deviation in percentage of survival among different batches at each week. WT WIK zebrafish were employed. <a href="/files/ftp_upload/57567/57567fig3large.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/57567/57567fig4.jpg" /> 
Figure 4: Representative cardiac function assessment following DOX stress using casper; Tg(cmlc2:nusDsRed) fish. (4A) A photograph of an adult casper; Tg(cmlc2:nusDsRed) fish. Scale bar = 1 cm. (4B) Representative image of a red ventricle at the end systolic stage of a matured casper; Tg(cmlc2:nusDsRed) fish. Dash line represents end systolic diameter (ESD). (4C) Representative image of a red ventricle at the end diastolic stage of a matured casper; Tg(cmlc2:nusDsRed) fish. Images in (4B) and (4C) were extracted from movies of a beating heart captured with a fluorescent dissection microscope using 6.3x magnification. Dash line represents end diastolic diameter (EDD). Scale bar = 1 mm. (4D) Representative cardiac function measured using casper;Tg(cmlc2:nuDsRed) adult zebrafish after 20 mg/kg DOX injection. EDD and ESD were measured as shown in (4B) and (4C) for individual fish, and ventricular fraction shortening is calculated by the formula (EDD &#8722; ESD)/EDD. Significantly decreased ventricular fraction shortening was detected at 4 weeks and thereafter. The alternative intraperitoneal method was used. Values are shown as the mean &#177; standard error. n &#8805; 3 in each group. The Student t test was used for comparison of two groups. *p &#60;0.05. These figures have been modified from Ding et al.23 <a href="/files/ftp_upload/57567/57567fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Medicine

9.7K Views.  Mayo Clinic Graduate School of Biomedical Sciences. 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.

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 ...

Assessing Teratogenic Changes in a Zebrafish Model of Fetal Alcohol Exposure

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, including mental retardation due to disordered and damaged brain development. Fetal alcohol spectrum disorder (FASD) is a term used to cover a continuum of birth defects that occur due to maternal alcohol consumption, and occurs in approximately 4% of children born in the United States. With 50% of child-bearing age women reporting consumption of alcohol, and half of all pregnancies being unplanned, unintentional exposure is a continuing issue2. In order to best understand the damage produced by ethanol, plus produce a model with which to test potential interventions, we developed a model of developmental ethanol exposure using the zebrafish embryo. Zebrafish are ideal for this kind of teratogen study3-8. Each pair lays hundreds of eggs, which can then be collected without harming the adult fish. The zebrafish embryo is transparent and can be readily imaged with any number of stains. Analysis of these embryos after exposure to ethanol at different doses and times of duration and application shows that the gross developmental defects produced by ethanol are consistent with the human birth defect. Described here are the basic techniques used to study and manipulate the zebrafish FAS model.

In order to understand the molecular mechanisms of the ethanol-induced developmental damage, we have developed a zebrafish model of ethanol exposure and are exploring the physical, cellular, and genetic alterations that occur after ethanol exposure1. We then seek to find potential interventions and rapidly test them in this animal model.

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, ...

Screening for Melanoma Modifiers using a Zebrafish Autochthonous Tumor Model

Genomic studies of human cancers have yielded a wealth of information about genes that are altered in tumors1,2,3. A challenge arising from these studies is that many genes are altered, and it can be difficult to distinguish genetic alterations that drove tumorigenesis from that those arose incidentally during transformation. To draw this distinction it is beneficial to have an assay that can quantitatively measure the effect of an altered gene on tumor initiation and other processes that enable tumors to persist and disseminate. Here we present a rapid means to screen large numbers of candidate melanoma modifiers in zebrafish using an autochthonous tumor model4 that encompasses steps required for melanoma initiation and maintenance. A key reagent in this assay is the miniCoopR vector, which couples a wild-type copy of the mitfa melanocyte specification factor to a Gateway recombination cassette into which candidate melanoma genes can be recombined5. The miniCoopR vector has a mitfa rescuing minigene which contains the promoter, open reading frame and 3'-untranslated region of the wild-type mitfa gene. It allows us to make constructs using full-length open reading frames of candidate melanoma modifiers. These individual clones can then be injected into single cell Tg(mitfa:BRAFV600E);p53(lf);mitfa(lf)zebrafish embryos. The miniCoopR vector gets integrated by Tol2-mediated transgenesis6 and rescues melanocytes. Because they are physically coupled to the mitfa rescuing minigene, candidate genes are expressed in rescued melanocytes, some of which will transform and develop into tumors. The effect of a candidate gene on melanoma initiation and melanoma cell properties can be measured using melanoma-free survival curves, invasion assays, antibody staining and transplantation assays.

A rapid way to screen for melanoma modifiers using a zebrafish autochthonous tumor model is presented. It takes advantage of the miniCoopR vector which allows for expression of candidate melanoma genes in melanocytes. A method to obtain melanoma-free survival curves, an invasion assay, a protocol for antibody staining of scale melanocytes and a melanoma transplantation assay are described.

Genomic studies of  human cancers have yielded a wealth of information about genes that are altered  in tumors1,2,3. A challenge arising from these ...

A Zebrafish Model of Diabetes Mellitus and Metabolic Memory

Diabetes mellitus currently affects 346 million individuals and this is projected to increase to 400 million by 2030. Evidence from both the laboratory and large scale clinical trials has revealed that diabetic complications progress unimpeded via the phenomenon of metabolic memory even when glycemic control is pharmaceutically achieved. Gene expression can be stably altered through epigenetic changes which not only allow cells and organisms to quickly respond to changing environmental stimuli but also confer the ability of the cell to "memorize" these encounters once the stimulus is removed. As such, the roles that these mechanisms play in the metabolic memory phenomenon are currently being examined.
We have recently reported the development of a zebrafish model of type I diabetes mellitus and characterized this model to show that diabetic zebrafish not only display the known secondary complications including the changes associated with diabetic retinopathy, diabetic nephropathy and impaired wound healing but also exhibit impaired caudal fin regeneration. This model is unique in that the zebrafish is capable to regenerate its damaged pancreas and restore a euglycemic state similar to what would be expected in post-transplant human patients. Moreover, multiple rounds of caudal fin amputation allow for the separation and study of pure epigenetic effects in an in vivo system without potential complicating factors from the previous diabetic state. Although euglycemia is achieved following pancreatic regeneration, the diabetic secondary complication of fin regeneration and skin wound healing persists indefinitely. In the case of impaired fin regeneration, this pathology is retained even after multiple rounds of fin regeneration in the daughter fin tissues. These observations point to an underlying epigenetic process existing in the metabolic memory state. Here we present the methods needed to successfully generate the diabetic and metabolic memory groups of fish and discuss the advantages of this model.

Metabolic memory is the phenomenon by which diabetic complications persist and progress unimpeded even after euglycemia is achieved pharmaceutically. Here we describe a diabetes mellitus zebrafish model which is unique in that it allows for the examination of the mitotically transmissible epigenetic components of metabolic memory in vivo.

Diabetes mellitus currently affects 346 million individuals and this is projected to increase to 400 million by 2030. Evidence from both the laboratory ...

A Possible Zebrafish Model of Polycystic Kidney Disease: Knockdown of wnt5a Causes Cysts in Zebrafish Kidneys

Polycystic kidney disease (PKD) is one of the most common causes of end-stage kidney disease, a devastating disease for which there is no cure. The molecular mechanisms leading to cyst formation in PKD remain somewhat unclear, but many genes are thought to be involved. Wnt5a is a non-canonical glycoprotein that regulates a wide range of developmental processes. Wnt5a works through the planar cell polarity (PCP) pathway that regulates oriented cell division during renal tubular cell elongation. Defects of the PCP pathway have been found to cause kidney cyst formation. Our paper describes a method for developing a zebrafish cystic kidney disease model by knockdown of the wnt5a gene with wnt5a antisense morpholino (MO) oligonucleotides. Tg(wt1b:GFP) transgenic zebrafish were used to visualize kidney structure and kidney cysts following wnt5a knockdown. Two distinct antisense MOs (AUG - and splice-site) were used and both resulted in curly tail down phenotype and cyst formation after wnt5a knockdown. Injection of mouse Wnt5a mRNA, resistant to the MOs due to a difference in primary base pair structure, rescued the abnormal phenotype, demonstrating that the phenotype was not due to &#8220;off-target&#8221; effects of the morpholino. This work supports the validity of using a zebrafish model to study wnt5a function in the kidney.

A Possible Zebrafish Model of Polycystic Kidney Disease: Knockdown of wnt5a Causes Cysts in Zebrafish Kidneys

We describe a method of generating a possible zebrafish model of polycystic kidney disease. We used Tg(wt1b:GFP) fish to visualize kidney structure. Knockdown of wnt5a was by morpholino injection. Pronephric cyst formation after wnt5a knockdown was observed in this GFP transgenic zebrafish.

Polycystic kidney disease (PKD) is one of the most common causes of end-stage kidney disease, a devastating disease for which there is no cure. The ...

Tachycardia-Induced Cardiomyopathy As a Chronic Heart Failure Model in Swine

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment methods. Here, we present such a model by tachycardia-induced cardiomyopathy, which can be produced by rapid cardiac pacing in swine.
A single pacing lead is introduced transvenously into fully anaesthetized healthy swine, to the apex of the right ventricle, and fixated. Its other end is then tunneled dorsally to the paravertebral region. There, it is connected to an in-house modified heart pacemaker unit that is then implanted in a subcutaneous pocket. 
After 4 - 8 weeks of rapid ventricular pacing at rates of 200 - 240 beats/min, physical examination revealed signs of severe heart failure - tachypnea, spontaneous sinus tachycardia, and fatigue. Echocardiography and X-ray showed dilation of all heart chambers, effusions, and severe systolic dysfunction. These findings correspond well to decompensated dilated cardiomyopathy and are also preserved after the cessation of pacing.
This model of tachycardia-induced cardiomyopathy can be used for studying the pathophysiology of progressive chronic heart failure, especially hemodynamic changes caused by new treatment modalities like mechanical circulatory supports. This methodology is easy to perform and the results are robust and reproducible.

Here, we present a protocol to produce tachycardia-induced cardiomyopathy in swine. This model represents a potent way to study the hemodynamics of progressive chronic heart failure and the effects of applied treatment.

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

Isolation of Atrial Myocytes from Adult Mice

The electrophysiological properties of atrial myocytes importantly affect overall cardiac function. Alterations in the underlying ionic currents responsible for the action potential can cause pro-arrhythmic substrates that underlie arrhythmias, such as atrial fibrillation, which are highly prevalent in many conditions and disease states. Isolating adult mouse atrial cardiomyocytes for use in patch-clamp experiments has greatly advanced our knowledge and understanding of the cellular electrophysiology in the healthy atrial myocardium and in the setting of atrial pathophysiology. In addition, studies using genetic mouse models have elucidated the role of a vast array of proteins in regulating atrial electrophysiology. Here we provide a detailed protocol for the isolation of cardiomyocytes from the atrial appendages of adult mice using a combination of enzymatic digestion and mechanical dissociation of these tissues. This approach consistently and reliably yields isolated atrial cardiomyocytes that can then be used to characterize cellular electrophysiology by measuring action potentials and ionic currents in patch-clamp experiments under a number of experimental conditions.

This protocol is used to isolate single atrial cardiomyocytes from the adult mouse heart using a chunk digestion approach. This approach is used to isolate right or left atrial myocytes that can be used to characterize atrial myocyte electrophysiology in patch-clamp studies.

The electrophysiological properties of atrial myocytes importantly affect overall cardiac function. Alterations in the underlying ionic currents ...

Murine Myocardial Infarction Model using Permanent Ligation of Left Anterior Descending Coronary Artery

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine models of MI with permanent ligation of left-anterior descending (LAD) coronary artery closely mimics MI in humans. Murine models benefit from the extensive genetic engineering available nowadays. Here we propose a reproducible murine surgical model of myocardial infarction by permanent LAD coronary ligation. Our technique comprises anesthesia with ketamine/xylazine that can be rapidly reversed by administration of an antagonist, intubation without tracheotomy for mechanical-assisted ventilation, ventilation with application of extrinsic positive end-expiratory pressure (PEEP) to avoid alveolar collapse, a thoracotomy method limiting to the minimum surgical lesions made to skeletal muscles, and lung inflation without thoracentesis. This method is sparsely invasive, reproducible and reduces post-surgery mortality and complications.

Herein we describe a surgical procedure showing how to achieve permanent ligation of the left-anterior descending coronary artery in mice. This model is of high relevance to investigate the pathophysiology of myocardial infarction and the concomitant biological processes.

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine ...

A Doxorubicin-Induced Murine Model of Dilated Cardiomyopathy In Vivo

Dilated cardiomyopathy (DCM) refers to a spectrum of heterogeneous myocardial disorders characterized by ventricular dilation and depressed cardiac performance in the absence of hypertension, valvular, congenital, or ischemic heart diseases, and which may be related to infection, autoimmune or metabolic abnormalities, or family inheritance. It can progress into congestive heart failure with a poor prognosis. Doxorubicin (Dox) is widely employed as a chemotherapeutic drug, but its use is limited because it causes DCM-like changes of the myocardium. Its myocardial toxicity is attributed to oxidative stress, chronic inflammation, and cardiomyocyte apoptosis. A model of DCM exploiting these Dox-induced DCM symptoms has not been established.

Described is a protocol to establish a Doxorubicin-induced dilated cardiomyopathy (DCM) model in mice via long-term intraperitoneal injection of Doxorubicin.

Dilated cardiomyopathy (DCM) refers to a spectrum of heterogeneous myocardial disorders characterized by ventricular dilation and depressed cardiac ...