We wish to thank&#160;Trent Waugh for his&#160;technical assistance. We also acknowledge the Department of Pediatrics at the Hospital for Sick Children and Cure Congenital Muscular Dystrophy (CMD) for their generous funding for this project.

1. Preparation of Agar Injection Plates (Time: 45 min)
<ol>
	<li>Boil 2% to 3% agarose in E3 media and allow solution to cool slightly on bench. Note: The number of injection plates being prepared dictates the amount of agarose required. Each injection plate needs approximately 35 ml of the agarose solution.</li>
	<li>After boiling, allow the agarose to cool until desired temperature is reached (e.g., 45 &#176;C) as per the injection mold manufacturer&#8217;s instructions.</li>
	<li>Pour approximately 35 ml of the cooled agarose into a 100 mm dish.</li>
	<li>Place one end of preferred injection mold into solution, then lay remainder of mold onto agarose solution (this will help reduce the occurrence of air bubbles).</li>
	<li>Allow the agarose solution to solidify either at RT or 4 &#176;C for approximately 30 min.</li>
	<li>Use a spatula to separate one end of the mold from the solid agarose. Slowly remove the remainder of the mold.</li>
</ol>
2. Preparation of Evans Blue Dye (EBD) Injection Mix (Time: 30 min)
<ol>
	<li>Make a 1% stock of EBD in 1X Ringer&#8217;s solution (155 mM NaCl; 5 mM KCl; 2 mM CaCl2; 1 mM MgCl2; 2 mM Na2HPO4; 10 mM HEPES; 10 mM glucose; pH to 7.2), which can be stored at RT.</li>
	<li>Make a stock solution of fluorescein isothiocyanate (FITC)-dextran MW 10,000 kDa at 25 mg/mL in 1X Ringer&#8217;s solution and store at -20 &#176;C.</li>
	<li>Prepare injection mix by diluting EBD to 0.1% directly in stock solution of FITC-dextran stock (i.e., for a final working volume of 100 &#181;l: Mix 10 &#181;l of 1% EBD in 90 &#181;l of FITC dextran stock).</li>
	<li>Thoroughly vortex injection mix (it should turn green) and keep out of direct light by wrapping injection mix tube in aluminum foil.</li>
</ol>
3. EBD Injection Preparation (Time: Approximately 30 min)
Note: Protocol works best with larvae from 3-7 days post fertilization (dpf).
<ol>
	<li>Pre-warm injection plate to RT.</li>
	<li>Set up injection apparatus by arranging micromanipulator on metal plate and stand next to the microscope being used for injection. Turn on air driven microinjection controller. Note: The preferred injection system will vary by lab and should not change outcome of analysis.</li>
	<li>Back fill injection needle with approximately 2-4 &#181;l of EBD mix.</li>
	<li>Calibrate injection volume to approximately 5 nl of EBD mix. Note: Injection volume calibration will depend on calibration method. A piston driven injection can be directly set to a given injection volume whereas gas pressure injectors will need the injection volume calibrated via volume bolus with the use of a micrometer.</li>
	<li>Wet injection plate with 1X Ringer&#8217;s solution and remove excess from wells.</li>
	<li>Pre-treat larvae with 0.04% ethyl 3-aminobenzoate methanesulfonate salt (tricaine) diluted in 1X Ringer&#8217;s solution to immobilize larvae prior to the start of injection. Note: Ensuring larvae are completely immotile is important as proper injection is difficult with any residual movement.</li>
	<li>Place anaesthetized larvae into wells of the agar injection plates using a glass pipette. Ensure that the larvae are completely within the well and lying on their side. Note: The number of larvae per well is up to the experimenter.</li>
	<li>After larvae are put in the wells, remove excess Ringer&#8217;s solution to minimize larvae movement within the well. Leave a residual amount of solution so that larvae do not dehydrate.</li>
</ol>
4. Pericardial Injection of Zebrafish Larvae with EBD (Time: Dependent on number of larvae injecting, estimated 1-3 hr)
<ol>
	<li>Place the injection plate containing the larvae on a dissecting scope where the injections will be performed.</li>
	<li>Position the injection pipette needle containing the EBD mix over a zebrafish larvae.</li>
	<li>Re-position the injection plate by rotating it so the injection needle is near the larvae&#8217;s heart and approximately 45&#176; ventrally from the anterior-posterior axis.</li>
	<li>Insert the injection needle into the common cardinal vein (CCV) in the region of the vein at the anterior portion of the yolk where the vein is initially turning in the dorsal direction (Figure 1). Note: A magnification of up to 40x may be useful to clearly see the CCV.</li>
	<li>Inject 5 nl of EBD mix and keep injection needle in position for 5-8 sec to minimize immediate leakage of EBD mix. Note: A good injection will have dye coloration seen in the heart chambers (Figure 1). If EBD mix is not observed in the heart, then injecting an additional 5 nl of EBD mix may be sufficient to induce dye uptake. Alternatively, the embryo can be discarded. 
	Note: In some situations, the heart may stop beating. If this occurs, continue to monitor the larvae for 20-40 sec. Typically, the heart resumes beating as the dye moves through the circulatory system.</li>
	<li>Move onto the next larvae and repeat.</li>
	<li>Identify successfully injected embryos by observing the presence of FITC-dextran in the vasculature immediately after injection (Figure 2).</li>
</ol>
5. Incubation and EBD Uptake (Time: 4-6 hr)
<ol>
	<li>After the desired number of larvae are injected, return injected larvae to 1X Ringer&#8217;s solution without tricaine in 100 mm dishes.</li>
	<li>Keep dishes wrapped in aluminum foil. Note: Keeping injected larvae in the dark significantly increases survival rates and ensures the greatest consistency in signal strength. Wrapping in aluminum foil is especially important for the period of time that the larvae are outside of the incubator.</li>
	<li>Allow larvae to incubate at 28.5 &#176;C for 4-6 hr to ensure sufficient EBD uptake.</li>
</ol>
6. Visualization of EBD in the Muscle (Time: Dependent on number of larvae injecting and type of microscopy, estimated 0.5-3 hr)
<ol>
	<li>Prior to imaging, anesthetize the larvae with 0.04% tricaine to prevent movement.</li>
	<li>View larvae under red fluorescence to determine if EBD uptake is occurring in the skeletal muscle (Figure 3).</li>
</ol>

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<li>Cavanna, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Molecular+and+genetic+mapping+of+the+mouse+mdx+locus%5BTitle%5D)+AND+%22Genomics%22%5BJournal%5D)">Molecular and genetic mapping of the mouse mdx locus.</a> Genomics. 3, 337-341 (1988).</li>
<li>Bassett, D. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dystrophin+is+required+for+the+formation+of+stable+muscle+attachments+in+the+zebrafish+embryo%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Dystrophin is required for the formation of stable muscle attachments in the zebrafish embryo.</a> Development. 130, 5851-5860 (2003).</li>
<li>Horstick, E. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stac3+is+a+component+of+the+excitation-contraction+coupling+machinery+and+mutated+in+Native+American+myopathy%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">Stac3 is a component of the excitation-contraction coupling machinery and mutated in Native American myopathy.</a> Nat Commun. 4, (1952).</li>
<li>Davidson, A. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Novel+deletion+of+lysine+7+expands+the+clinical%2C+histopathological+and+genetic+spectrum+of+TPM2-related+myopathies%5BTitle%5D)+AND+%22Brain%22%5BJournal%5D)">Novel deletion of lysine 7 expands the clinical, histopathological and genetic spectrum of TPM2-related myopathies.</a> Brain. 136, 508-521 (2013).</li>
<li>Telfer, W. R., Nelson, D. D., Waugh, T., Brooks, S. V., Dowling, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Neb%3A+a+zebrafish+model+of+nemaline+myopathy+due+to+nebulin+mutation%5BTitle%5D)+AND+%22Dis+Model+Mech%22%5BJournal%5D)">Neb: a zebrafish model of nemaline myopathy due to nebulin mutation.</a> Dis Model Mech. 5, 389-396 (2012).</li>
<li>Dowling, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Oxidative+stress+and+successful+antioxidant+treatment+in+models+of+RYR1-related+myopathy%5BTitle%5D)+AND+%22Brain%22%5BJournal%5D)">Oxidative stress and successful antioxidant treatment in models of RYR1-related myopathy.</a> Brain. 135, 1115-1127 (2012).</li>
<li>Detrich, H. W., Westerfield 3rd,, M,, Zon, L. I. Overview of the Zebrafish system. Methods Cell Biol. 59, 3-10 (1999).</li>
<li>Whitfield, T. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mutations+affecting+development+of+the+zebrafish+inner+ear+and+lateral%5BTitle%5D)+AND+%22%22%5BJournal%5D)">Mutations affecting development of the zebrafish inner ear and lateral.</a> , 123-241 (1996).</li>
<li>Kawahara, G., Guyon, J. R., Nakamura, Y., Kunkel, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Zebrafish+models+for+human+FKRP+muscular+dystrophies%5BTitle%5D)+AND+%22Hum+Mol+Genet%22%5BJournal%5D)">Zebrafish models for human FKRP muscular dystrophies.</a> Hum Mol Genet. 19, 623-633 (2010).</li>
<li>Baraban, S. C., Dinday, M. T., Hortopan, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Drug+screening+in+Scn1a+zebrafish+mutant+identifies+clemizole+as+a+potential+Dravet+syndrome+treatment%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">Drug screening in Scn1a zebrafish mutant identifies clemizole as a potential Dravet syndrome treatment.</a> Nat Commun. 4, (2013).</li>
<li>Waugh, T. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fluoxetine+prevents+dystrophic+changes+in+a+zebrafish+model+of+Duchenne+muscular+dystrophy%5BTitle%5D)+AND+%22Hum+Mol+Genet%22%5BJournal%5D)">Fluoxetine prevents dystrophic changes in a zebrafish model of Duchenne muscular dystrophy.</a> Hum Mol Genet. 23 (17), 4651-4662 Forthcoming.</li>
<li>Horstick, E. J., Gibbs, E. M., Li, X., Davidson, A. E., Dowling, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Analysis+of+embryonic+and+larval+zebrafish+skeletal+myofibers+from+dissociated+preparations%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Analysis of embryonic and larval zebrafish skeletal myofibers from dissociated preparations.</a> J Vis Exp. 50259, (2013).</li>
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<li>Guyon, J. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Genetic+isolation+and+characterization+of+a+splicing+mutant+of+zebrafish+dystrophin%5BTitle%5D)+AND+%22Hum+Mol+Genet%22%5BJournal%5D)">Genetic isolation and characterization of a splicing mutant of zebrafish dystrophin.</a> Hum Mol Genet. 18, 202-211 (2009).</li>
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<li>Majczenko, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dominant+Mutation+of+CCDC78+in+a+Unique+Congenital+Myopathy+with+Prominent+Internal+Nuclei+and+Atypical+Cores%5BTitle%5D)+AND+%22Am+J+Hum%22%5BJournal%5D)">Dominant Mutation of CCDC78 in a Unique Congenital Myopathy with Prominent Internal Nuclei and Atypical Cores.</a> Am J Hum. , (2012).</li>
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<li>Hall, T. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+zebrafish+candyfloss+mutant+implicates+extracellular+matrix+adhesion+failure+in+laminin+alpha2-deficient+congenital+muscular+dystrophy%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">The zebrafish candyfloss mutant implicates extracellular matrix adhesion failure in laminin alpha2-deficient congenital muscular dystrophy.</a> Proc Natl Acad Sci U S A. 104, 7092-7097 (2007).</li>
</ol>

The authors have no competing financial interests or other conflicts of interest.

Zebrafish are emerging as a powerful tool for the study of neuromuscular disease2,29. To date, the zebrafish system has been used to validate new muscle disease-causing mutations16,17,30, elucidate novel pathomechanisms18, and identify potentially new therapeutic drugs12,24. These collective efforts have established the utility of the zebrafish to model human neuromuscular diseases. However, despite the advances made with zebrafish and mammalian models, there are limited treatment options for patients within the wide spectrum of neuromuscular conditions. Therefore, a high demand exists for therapy development for this group of devastating diseases. Paralleling this demand for therapeutics is the corresponding need for ongoing experimental innovation, as well as rigorous analysis to verify new animal models and putative therapeutic strategies.
EBD analysis is commonly used in mouse models to study tissue and cellular damage in brain, heart, and skeletal muscle27,31. Most notably, EBD is used extensively in mouse models of various muscular dystrophy subtypes to show the severity of muscle membrane instability and damage8. The use of EBD to reveal muscle membrane damage is a supportive parameter establishing similarities of the animal model to the human disease state9. The power of EBD in mouse has led several labs, including our own, to develop and apply EBD to zebrafish models of neuromuscular disease. Due to the applicability of EBD analysis, this technique is actively being implemented to corroborate zebrafish models to the human disease state11,15,22,24,32. Larvae with damaged muscle membranes will have EBD uptake and therefore red fluorescence within muscle fibers. Fluorescence observed in the inter-fiber space, but not within individual muscle fibers may also be informative of fibers detaching from the basement membrane in the absence of membrane damage, providing useful diagnostic detail. EBD analysis has potential application beyond animal model validation. Efforts from our lab have recently demonstrated that EBD analysis is beneficial in validating potentially novel therapeutic drugs24. Determining if potential therapeutic treatments reduce or abolish EBD uptake in neuromuscular disease models can signify relevant therapeutic action8. This type of analysis can help establish the mechanism(s) of therapeutics and expands the application of EBD analysis.
As with many techniques, EBD analysis does have several caveats to be observed during experimental design and practice. For example, it can be challenging to identify the CCV due to the thickening of the tissue with age. Additionally, it is easy to damage larvae in preparation before and during the pericardial injection, reducing experimental counts and increasing the need to prep large numbers of larvae. Furthermore, physical damage done to the larvae during handling and injection could result in false positives as damaged muscle can take up EBD. In order to overcome some of these obstacles, we have described a co-injection strategy in this video article that allows easy and reliable identification of larvae with successful dye infusion immediately following injection and prior to subsequent analysis. The FITC-dextran co-injection controls for successful injection by allowing confirmation of EBD in the vasculature prior to its uptake by the muscle fibers. This can be particularly useful as EBD fluorescence becomes highly diffuse in larvae after several hours if not collected in the muscle fibers; as such, it can be difficult to detect. In addition, missing the CCV and injecting EBD into the yolk or body cavity can, after incubation, result in diffuse red fluorescence similar to control embryos, yet with the reduced likelihood of uptake by damaged muscle fibers. Collectively, these caveats suggest EBD injection requires patience and practice in order to achieve consistent and reliable results.
In all, we describe a practical and straightforward method to perform EBD analysis on zebrafish larvae. To date, the use of zebrafish as a model system, especially as a human disease model, has been rapidly expanding. This expansion is partially due to the continued development and modification of experimental techniques that improve upon the current advantages of the zebrafish system. The EBD injection technique provides an additional and powerful tool to a researcher&#8217;s arsenal for the validation and study of zebrafish muscle disease models. The ongoing implementation and modification of this technique has the potential to help uncover novel therapeutic strategies as well as disease causing mechanisms.

Muscular dystrophies constitute a group of prevalent and heterogeneous human muscle diseases with specific histopathological features1,2. Symptoms typically associated with this devastating group of diseases include muscle weakness, muscle degeneration, loss of motility, and early mortality1,3. The primary pathomechanisms of muscular dystrophies are the loss of proteins that stabilize the sarcolemma, anchor transmembrane complexes, and mediate cell signaling across the membrane4-6. For example, complete loss of the protein dystrophin, a primary scaffold protein of the dystrophin-glycoprotein complex, results in destabilization of the muscle membrane in Duchenne muscular dystrophy7. Due to the fact that most muscular dystrophies result from mutations in proteins that participate in the link between the extracellular matrix and the sarcolemmal cytoskeleton, a common observation at the cellular level is the loss of sacrolemmal integrity8,9. This understanding of the primary pathomechanism(s) associated with muscular dystrophies is the product of numerous years of research employing animal model systems2,10-15. However, despite advances in the field, there are still limited therapeutic options for treatment or management of the range of dystrophy subtypes. This limitation of viable therapies is due to several key factors: 1) the difficulty of gene therapy strategies, 2) a high frequency of de-novo disease cases and the corresponding lack of translatable animal models, and 3) the lack of rigorous strategies to test the physiological consequences of putative therapeutic agents with clear and reproducible outcome measures.
To overcome some of these limitations, numerous labs including our own are employing zebrafish as a system to model and study human neuromuscular diseases2. To date, zebrafish have proven a valuable tool in disease research. The zebrafish model has been used to identify and validate novel human disease causing mutations16,17, elucidate uncharacterized disease causing mechanisms17,18, and identify novel therapeutic strategies12,19. These advances were made, in part, by the canonical strengths of the zebrafish system such as their optical clarity, ease of genetic manipulation, and ability to breed in large numbers20. Zebrafish have additionally proven amendable to large-scale drug screens21, a valuable method for the identification of novel therapeutics22-24. Regarding muscle disease research, these strengths are complemented by the ability to isolate single zebrafish skeletal muscle fibers via dissociation25 and by the ability to examine myofiber organization in vivo using the optical phenomenon called birefringence26, which collectively allows for rapid determination of macroscopic muscle integrity. Regardless of these available utilities, further tool development is continuously required to advance investigation.
We, and others, have adapted a protocol for EBD injection and analysis in the zebrafish model. EBD is a vital, cell permeable dye that is taken up by damaged, degenerating, or apoptotic cells and then visualized under fluorescence27. To date, EBD analysis has extensively been used to analyze muscle membrane integrity in mouse models of skeletal muscle and heart diseases8,9,27. However, in mammalian preparations, harvested muscle typically requires laborious sectioning or dissection prior to analysis. In zebrafish, direct analysis is possible in high numbers using live and intact animals. In this video article, we will demonstrate the methodology to perform EBD injection and analysis in live zebrafish larvae, with representative images of EBD uptake in the zebrafish dystrophy mutant line sapje15,28. Furthermore, we present a co-injection strategy that allows for increased quantification of EBD preparations.

<table><tbody><tr><td>Fluorescein isothiocyanate-dextran MW 10,000</td><td>Sigma</td><td>FD10S</td><td></td></tr><tr><td>Evan&#039;s Blue Dye</td><td>Sigma</td><td>E2129</td><td></td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate salt</td><td>Sigma</td><td>A5040</td><td></td></tr><tr><td>100 mm Petri dish</td><td>Fischerbrand</td><td>FB0875712</td><td>Injection mold base</td></tr><tr><td>Thin wall glass capillaries</td><td>World Precision Instruments</td><td>TW100F-4</td><td>For Injection needle</td></tr><tr><td>Agarose</td><td>Bioshop</td><td>AGA001</td><td>Injection mold</td></tr><tr><td>Microinjection mold</td><td>Adaptive Science Tools</td><td>TU-1</td><td>Injection mold</td></tr><tr><td>Sodium chloride</td><td>Bioshop</td><td>SOD001</td><td>Ringer&#039;s solution</td></tr><tr><td>Potassium chloride</td><td>Bioshop</td><td>POC888</td><td>Ringer&#039;s solution</td></tr><tr><td>Magnessium chloride hexahydrate</td><td>Sigma</td><td>M2670</td><td>Ringer&#039;s solution</td></tr><tr><td>Sodium phosphate monobasic monohydrate</td><td>Sigma</td><td>S9638</td><td>Ringer&#039;s solution</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H4034</td><td>Ringer&#039;s solution</td></tr><tr><td>Glucose</td><td>BioBasic</td><td>GB0219</td><td>Ringer&#039;s solution</td></tr><tr><td>Calcium chloride</td><td>Sigma</td><td>C1061</td><td>Ringer&#039;s solution</td></tr></tbody></table>

analysis of zebrafish larvae skeletal muscle integrity with evans blue dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

The EBD injection mix was injected into the CCV of sapje homozygous mutants and wild-type siblings at 3 dpf. Injections that filled the heart chambers (Figure 1B) were then analyzed for successful injection by visualizing FITC-dextran in the vasculature under green fluorescence (Figure 2).
After a 4 hr incubation period, EBD uptake was examined at the somite level using fluorescence microscopy. Wild type siblings exhibited no EBD fluorescence within any visible muscle fibers (Figure 3A), whereas the sapje homozygous mutants showed EBD uptake, indicating damage to the muscle membrane15 (Figure 3B).
<img alt="Figure 1" src="/files/ftp_upload/53183/53183fig1.jpg" /> 
Figure 1.&#160;Injecting EBD injection mix into the common cardinal vein (CCV) of a zebrafish embryo. (A) Uninjected embryo. Arrow denotes ideal location for CCV injection. (B) Successful injection into the CCV. The dye enters the heart chambers (arrow) and begins to be pumped through the vasculature. (C) An unsuccessful CCV injection will result in some or all of the dye entering the yolk sac of the embryo (arrow). <a href="https://www.jove.com/files/ftp_upload/53183/53183fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53183/53183fig2.jpg" /> 
Figure 2. Embryos can be sorted for successful injection by observing FITC-dextran distribution under green fluorescence throughout the vasculature immediately following injection and prior to EBD uptake. <a href="https://www.jove.com/files/ftp_upload/53183/53183fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53183/53183fig3.jpg" /> 
Figure 3: EBD will be taken up by fibers with damaged membranes.&#160;(A) Wildtype siblings showing no EBD fluorescence in muscle fibers. (B) Sapje homozygous mutant with EBD fluorescence within multiple muscle fibers (arrows). All larvae were injected with the EBD injection mix and analyzed after a 4 hr incubation period at 3 dpf. Siblings and mutants were sorted by muscle fiber detachment prior to CCV injection. <a href="https://www.jove.com/files/ftp_upload/53183/53183fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye at JoVE.com

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

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Developmental Biology

9.1K Views.  The Hospital for Sick Children. In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

Video: Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many myopathies affecting skeletal muscle function can be quickly and easily assessed in zebrafish over the first few days of embryogenesis. By 24 hr post-fertilization (hpf), wildtype zebrafish spontaneously contract their tail muscles and by 48 hpf, zebrafish exhibit controlled swimming behaviors. Reduction in the frequency of, or other alterations in, these movements may indicate a skeletal muscle dysfunction. To analyze swimming behavior and assess muscle performance in early zebrafish development, we utilize both touch-evoked escape response and locomotion assays.
Touch-evoked escape response assays can be used to assess muscle performance during short burst movements resulting from contraction of fast-twitch muscle fibers. In response to an external stimulus, which in this case is a tap on the head, wildtype zebrafish at 2 days post-fertilization (dpf) typically exhibit a powerful burst swim, accompanied by sharp turns. Our method quantifies skeletal muscle function by measuring the maximum acceleration during a burst swimming motion, the acceleration being directly proportional to the force produced by muscle contraction.
In contrast, locomotion assays during early zebrafish larval development are used to assess muscle performance during sustained periods of muscle activity. Using a tracking system to monitor swimming behavior, we obtain an automated calculation of the frequency of activity and distance in 6-day old zebrafish, reflective of their skeletal muscle function. Measurements of swimming performance are valuable for phenotypic assessment of disease models and high-throughput screening of mutations or chemical treatments affecting skeletal muscle function.

Zebrafish are an excellent model to study muscle function and disease. During early embryogenesis zebrafish begin regular muscle contractions producing rhythmic swimming behavior, which is altered when the muscle is disrupted. Here we describe a touch-evoked response and locomotion assay to examine swimming performance as a measure of muscle function.

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many ...

Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics

Skeletal morphogenesis occurs through tightly regulated cell behaviors during development; many cell types alter their behavior in response to mechanical strain. Skeletal joints are subjected to dynamic mechanical loading. Finite element analysis (FEA) is a computational method, frequently used in engineering that can predict how a material or structure will respond to mechanical input. By dividing a whole system (in this case the zebrafish jaw skeleton) into a mesh of smaller 'finite elements', FEA can be used to calculate the mechanical response of the structure to external loads. The results can be visualized in many ways including as a 'heat map' showing the position of maximum and minimum principal strains (a positive principal strain indicates tension while a negative indicates compression. The maximum and minimum refer the largest and smallest strain). These can be used to identify which regions of the jaw and therefore which cells are likely to be under particularly high tensional or compressional loads during jaw movement and can therefore be used to identify relationships between mechanical strain and cell behavior. This protocol describes the steps to generate Finite Element models from confocal image data on the musculoskeletal system, using the zebrafish lower jaw as a practical example. The protocol leads the reader through a series of steps: 1) staining of the musculoskeletal components, 2) imaging the musculoskeletal components, 3) building a 3 dimensional (3D) surface, 4) generating a mesh of Finite Elements, 5) solving the FEA and finally 6) validating the results by comparison to real displacements seen in movements of the fish jaw.

Finite Element Analysis is a frequently used tool to investigate the mechanical performance of structures under load. Here we apply its use to modeling the biomechanics of the zebrafish jaw.

Skeletal morphogenesis occurs through tightly regulated cell behaviors during development; many cell types alter their behavior in response to mechanical ...

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be difficult to study, and can lead to confounding results. The protocol described here is a straightforward breeding paradigm to increase and decrease penetrance in lethal zebrafish skeletal mutants. Because lethal mutants cannot be selectively bred directly, the classic selective breeding strategy of progeny testing is employed. This method also includes protocols for Kompetitive Allele Specific PCR (KASP) genotyping zebrafish and staining larval zebrafish cartilage and bone. Applying the husbandry strategy described here can increase the penetrance of an interesting skeletal phenotype enabling more reproducible results in downstream applications. In addition, decreasing the mutant penetrance through this selective breeding strategy can reveal the developmental processes that most crucially require the function of the mutated gene. While the skeleton is specifically considered here, we propose that this methodology will be useful for all zebrafish mutant lines.

The goal of this protocol is to alter the penetrance of lethal skeletal mutant phenotypes in zebrafish by selective breeding. Lethal mutants cannot be grown to adulthood and bred themselves, therefore this protocol describes a method for tracking and selecting penetrance through multiple generations by progeny testing.

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be ...

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 ...

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, the muscle stem cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. In many muscle wasting conditions, including muscular dystrophy and ageing, this process is impaired resulting in the inability of muscle to regenerate. The process of muscle regeneration in zebrafish is highly conserved with mammalian systems providing an excellent system to study muscle stem cell function and regeneration, in muscle wasting conditions such as muscular dystrophy. Here, we present a method to examine muscle regeneration in zebrafish models of muscle disease. The first step involves the use of a genotyping platform that allows the determination of the genotype of the larvae prior to eliciting an injury. Having determined the genotype, the muscle is injured using a needle stab, following which polarizing light microscopy is used to determine the extent of muscle regeneration. We therefore provide a high throughput pipeline which allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, ...

Using Alizarin Red Staining to Detect Chemically Induced Bone Loss in Zebrafish Larvae

Chemically induced bone loss due to lead (Pb) exposure could trigger an array of adverse impacts on both human and animal skeletal systems. However, the specific effects and mechanisms in zebrafish remain unclear. Alizarin red has a high affinity for calcium ions and can help visualize the bone and illustrate skeletal mineral mass. In this study, we aimed to detect lead acetate (PbAc)-induced bone loss in zebrafish larvae by using alizarin red staining. Zebrafish embryos were treated with a series of PbAc concentrations (0, 5, 10, 20 mg/L) between 2 and 120 h post fertilization. Whole-mount skeletal staining was conducted on larvae at 9 days post fertilization, and the total stained area was quantified using ImageJ software. The results indicated that the mineralized tissues were stained in red, and the stained area decreased significantly in the PbAc-exposure group, with a dose-dependent change in bone mineralization. This paper presents a staining protocol for investigating skeletal changes in PbAc-induced bone defects. The method can also be used in zebrafish larvae for the detection of bone loss induced by other chemicals.

Here, we have used alizarin red staining to show that lead acetate exposure causes a bone mass change in zebrafish larvae. This staining method can be adapted to the investigation of bone loss in zebrafish larvae loss induced by other hazardous toxicants.

Chemically induced bone loss due to lead (Pb) exposure could trigger an array of adverse impacts on both human and animal skeletal systems. However, the ...

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

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

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

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

Biology

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

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

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

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