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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E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+deletion+of+lysine+7+expands+the+clinical,+histopathological+and+genetic+spectrum+of+TPM2-related+myopathies.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neb:+a+zebrafish+model+of+nemaline+myopathy+due+to+nebulin+mutation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxidative+stress+and+successful+antioxidant+treatment+in+models+of+RYR1-related+myopathy.">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, L. I., Zon, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> 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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutations+affecting+development+of+the+zebrafish+inner+ear+and+lateral.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+models+for+human+FKRP+muscular+dystrophies.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drug+screening+in+Scn1a+zebrafish+mutant+identifies+clemizole+as+a+potential+Dravet+syndrome+treatment.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluoxetine+prevents+dystrophic+changes+in+a+zebrafish+model+of+Duchenne+muscular+dystrophy.">Fluoxetine prevents dystrophic changes in a zebrafish model of Duchenne muscular dystrophy.</a> Hum Mol Genet. 23 (17), 4651-4662 .</li><li>Horstick, E. J., Gibbs, E. M., Li, X., Davidson, A. E., Dowling, J. 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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 border="0" cellpadding="0" cellspacing="0" width="801">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Fluorescein isothiocyanate-dextran MW 10,000</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:140px;">FD10S</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Evan&#39;s Blue Dye</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:140px;">E2129</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Ethyl 3-aminobenzoate methanesulfonate salt</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:140px;">A5040</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">100 mm Petri dish</td>
			<td style="width:173px;">Fischerbrand</td>
			<td style="width:140px;">FB0875712</td>
			<td>Injection mold base</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:321px;">Thin wall glass capillaries</td>
			<td style="width:173px;">World Precision Instruments</td>
			<td style="width:140px;">TW100F-4</td>
			<td>For Injection needle</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Agarose</td>
			<td style="width:173px;">Bioshop</td>
			<td style="width:140px;">AGA001</td>
			<td>Injection mold</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Microinjection mold</td>
			<td style="width:173px;">Adaptive Science Tools</td>
			<td style="width:140px;">TU-1</td>
			<td>Injection mold</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Sodium chloride</td>
			<td style="width:173px;">Bioshop</td>
			<td style="width:140px;">SOD001</td>
			<td>Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Potassium chloride</td>
			<td style="width:173px;">Bioshop</td>
			<td style="width:140px;">POC888</td>
			<td>Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Magnessium chloride hexahydrate</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:140px;">M2670</td>
			<td>Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Sodium phosphate monobasic monohydrate</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:140px;">S9638</td>
			<td>Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">HEPES</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:140px;">H4034</td>
			<td>Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Glucose</td>
			<td style="width:173px;">BioBasic</td>
			<td style="width:140px;">GB0219</td>
			<td>Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Calcium chloride</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:140px;">C1061</td>
			<td>Ringer&#39;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 ...

analysis-zebrafish-larvae-skeletal-muscle-integrity-with-evans-blue

Research

JoVE Journal

Developmental Biology

8.9K vues.  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.

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

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced skeletal muscle regeneration is a widely used, powerful model system to study the events involved in muscle regeneration as well as the mechanisms and different players. Indeed, a detailed knowledge of this process is essential for a better understanding of the pathological conditions that lead to skeletal muscle degeneration, and it aids in identifying new targeted therapeutic strategies. The present work describes a detailed and reproducible protocol to induce acute skeletal muscle regeneration in mice through a single intramuscular injection of cardiotoxin (CTX). CTX belongs to the family of snake venom toxins and causes myolysis of myofibers, which eventually triggers the regeneration events. The dynamics of skeletal muscle regeneration is evaluated by histological analysis of muscle sections. The protocol also illustrates the experimental procedures for dissecting, freezing, and cutting the Tibialis Anterior muscle, as well as the routine Hematoxylin &#38; Eosin staining that is widely used for subsequent morphological and morphometric analysis.

This manuscript describes a detailed protocol to induce acute skeletal muscle regeneration in adult mice and subsequent manipulations of the muscles, such as dissection, freezing, cutting, routine staining, and myofiber cross-sectional area analysis.

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

Application of Chronic Stimulation to Study Contractile Activity-induced Rat Skeletal Muscle Phenotypic Adaptations

Skeletal muscle is a highly adaptable tissue, as its biochemical and physiological properties are greatly altered in response to chronic exercise. To investigate the underlying mechanisms that bring about various muscle adaptations, a number of exercise protocols such as treadmill, wheel running, and swimming exercise have been used in the animal studies. However, these exercise models require a long period of time to achieve muscle adaptations, which may be also regulated by humoral or neurological factors, thus limiting their applications in studying the muscle-specific contraction-induced adaptations. Indirect low frequency stimulation (10 Hz) to induce chronic contractile activity (CCA) has been used as an alternative model for exercise training, as it can successfully lead to muscle mitochondrial adaptations within 7 days, independent of systemic factors. This paper details the surgical techniques required to apply the treatment of CCA to the skeletal muscle of rats, for widespread application in future studies.

This protocol describes the use of the chronic contractile activity model of exercise to observe stimulation-induced skeletal muscle adaptations in the rat hindlimb.

Skeletal muscle is a highly adaptable tissue, as its biochemical and physiological properties are greatly altered in response to chronic exercise. To ...

Identification of Skeletal Muscle Satellite Cells by Immunofluorescence with Pax7 and Laminin Antibodies

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, antibodies for specific cell markers are applied on tissue sections. In adult skeletal muscle, satellite cells (SCs) are stem cells that contribute to muscle repair and regeneration. Therefore, it is important to visualize and trace the satellite cell population under different physiological conditions. In resting skeletal muscle, SCs reside between the basal lamina and myofiber plasma membrane. A commonly used marker for identifying SCs on the myofibers or in cell culture is the paired box protein Pax7. In this article, an optimized Pax7 immunofluorescence protocol on skeletal muscle sections is presented that minimizes non-specific staining and background. Another antibody that recognizes a protein (laminin) of the basal lamina was also added to help identify SCs. Similar protocols can also be used to perform double or triple labeling with Pax7 and antibodies for additional proteins of interest.

The precise identification of satellite cells is essential for studying their functions under various physiological and pathological conditions. This article presents a protocol to identify satellite cells on adult skeletal muscle sections by immunofluorescence-based staining.

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, ...

Isolation, Culture, Characterization, and Differentiation of Human Muscle Progenitor Cells from the Skeletal Muscle Biopsy Procedure

The use of primary human tissue and cells is ideal for the investigation of biological and physiological processes such as the skeletal muscle regenerative process. There are recognized challenges to working with human primary adult stem cells, particularly human muscle progenitor cells (hMPCs) derived from skeletal muscle biopsies, including low cell yield from collected tissue and a large degree of donor heterogeneity of growth and death parameters among cultures. While incorporating heterogeneity into experimental design requires a larger sample size to detect significant effects, it also allows us to identify mechanisms that underlie variability in hMPC&#160;expansion capacity, and thus allows us to better understand heterogeneity in skeletal muscle regeneration. Novel mechanisms that distinguish the expansion capacity of cultures have the potential to lead to the development of therapies to improve skeletal muscle regeneration.

We present techniques for isolating, culturing, characterizing, and differentiating human primary muscle progenitor cells (hMPCs) obtained from skeletal muscle biopsy tissue. hMPCs obtained and characterized through these methods can be used to subsequently address research questions related to human myogenesis and skeletal muscle regeneration.

The use of primary human tissue and cells is ideal for the investigation of biological and physiological processes such as the skeletal muscle ...

Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this failure is the use of inadequate laboratory models. It is challenging to model complex human diseases in traditional laboratory organisms, but this issue can be circumvented through the study of human xenografts. The surgical method we describe here allows for the creation of human skeletal muscle xenografts, which can be used to model muscle disease and to carry out preclinical therapeutic testing. Under an Institutional Review Board (IRB)-approved protocol, skeletal muscle specimens are acquired from patients and then transplanted into NOD-Rag1null IL2r&gamma;null (NRG) host mice. These mice are ideal hosts for transplantation studies due to their inability to make mature lymphocytes and are thus unable to develop cell-mediated and humoral adaptive immune responses. Host mice are anaesthetized with isoflurane, and the mouse tibialis anterior and extensor digitorum longus muscles are removed. A piece of human muscle is then placed in the empty tibial compartment and sutured to the proximal and distal tendons of the peroneus longus muscle. The xenografted muscle is spontaneously vascularized and innervated by the mouse host, resulting in robustly regenerated human muscle that can serve as a model for preclinical studies.

Complex human diseases can be challenging to model in traditional laboratory model systems. Here, we describe a surgical approach to model human muscle disease through the transplantation of human skeletal muscle biopsies into immunodeficient mice.

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this ...

Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

Primary myoblasts are undifferentiated proliferating precursors of skeletal muscle. They can be cultured and studied as muscle precursors or induced to differentiate into later stages of muscle development. The protocol provided here describes a robust method for the isolation and culture of a highly proliferative population of myoblast cells from young adult mouse skeletal muscle explants. These cells are useful for the study of the metabolic properties of skeletal muscle of different mouse models, as well as in other downstream applications such as transfection with exogenous DNA or transduction with viral expression vectors. The level of differentiation and metabolic profile of these cells depends on the length of exposure, and composition of the media used to induce myoblast differentiation. These methods provide a robust system for the study of mouse muscle cell metabolism ex vivo. Importantly, unlike in vivo models, the methods described here provide a cell population that can be expanded and studied with high levels of reproducibility.

Myoblasts are proliferating precursor cells that differentiate to form polynucleated myotubes and eventually skeletal muscle myofibers. Here, we present a protocol for efficient isolation and culture of primary myoblasts from young adult mouse skeletal muscles. The method enables molecular, genetic, and metabolic studies of muscle cells in culture.

Primary myoblasts are undifferentiated proliferating precursors of skeletal muscle. They can be cultured and studied as muscle precursors or induced to ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...

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, ...