The authors wish to thank the members of the Dowling lab (Aaron Reifler, Trent Waugh, Angela Busta, and William Telfer) that contributed to the development of the technique and to the production of the manuscript. This work was funded by the Taubman Institute, the Department of Pediatrics at the University of Michigan, and in part from grants from the Muscular Dystrophy Association (JJD MDA186999) and the National Institutes of Health (JJD 1K08AR054835).

1. Preparation of Poly-L-Lysine Coated Coverslips (Time: 1 hr)
Coating coverslips allows for rapid myofiber settling and adhesion. This may be performed during the dissociation step of the myofiber isolation (step 2 below).
<ol>
	<li>Cut and place Parafilm on the bottom of a 60 mm Petri dish (any brand).</li>
	<li>Place microscope cover glass slips (12 mm diameter) on Parafilm in a 60 mm tissue culture dish or place them individually in single wells of 24-well plates.</li>
</ol>
Note 1: These small round coverslips help to concentrate myofiber numbers and reduce excess antibody usage.
Note 2: Other size coverslips will work; however, the volumes of reagents and embryos used will need to be adjusted accordingly.
<ol start="3">
	<li>Pipette 50-200 &#956;l of poly-L-lysine solution (0.01%) on each cover glass in the Petri dish.</li>
	<li>Allow the coverslips to sit at least one hour at 37 &#176;C. Longer incubations will not negatively influence results.</li>
	<li>Remove poly-L-lysine solution from the cover glass and wash twice with a minimum of 100 &#956;l 1x PBS.</li>
	<li>Allow coverslips to dry.</li>
	<li>Coverslips are now ready for myofibers.</li>
</ol>
Note 3: To ensure sterility, coating can be done in a hood and on autoclaved coverslips.
Note 4: Keep the 60 mm Petri dish with Parafilm on the bottom. This setup will be used during myofiber plating and immunolabeling (later steps).
Alternative 1: Instead of coating coverslips immediately prior to use, a coated coverslip stock can be used. A 60 mm Petri dish containing a minimum of 2 ml poly-L-lysine can be stored at 4 &#176;C containing numerous coverslips. With this alternative start at step 1.5 to process the coverslips for myofibers.
Alternative 2: We also coat coverslips in poly-L-ornithine. This is more labor intensive, but is useful for longer term culturing because poly-L-ornithine coated coverslips can be UV treated. With UV treatment and careful sterile technique live myofibers can typically be maintained in culture from 4-7 days.
2. Dissociation of Zebrafish Embryos and Plating of Myofibers (Time: 1 to 3 hr)
Note: the standard protocol applies best to 3 dpf (days post fertilization) embryos.
<ol>
	<li>Transfer zebrafish embryos into a standard 1.5 ml centrifuge tube and remove as much excess fish water as possible. Typically, 10-20 embryos per tube, though less can be used. Using more than 20-25 embryos often results in a preparation of excessive density.
	<ol>
		<li>Remove chorions prior to dissociation, manually using #5 forceps. Alternatively, chemical chorion removal is achieved with a 10-15 min Pronase treatment. Typically, the previous removal of the chorion is only needed when prior confirmation of a mutant phenotype or morphology is required, or when using stages where embryos are naturally hatched from their chorion.</li>
	</ol>
	</li>
	<li>Add 900 &#956;l of CO2&#160;independent media to the tube containing the embryos.</li>
	<li>Add 100 &#956;l of collagenase type II, final concentration 3.125 mg/ml (Stock collagenase solution at 31.25 mg/ml diluted in 1x PBS) to begin dissociation. Collagenase IV can also be used for the dissociation.</li>
	<li>Rotate embryos on an orbital shaker, and triturate every 30 min at RT using a P1000 Pipetman for the trituration.</li>
</ol>
Note 2: Carefully monitor embryo dissociation; over or under dissociation (especially over) are the most common reasons for protocol failure. Times for digestion will vary depending on intensity of trituration, number of embryos per tube, and age of embryos. It is also often less (in comparison to wild types) for embryos from skeletal muscle mutants.
Note 3: Average digestion time per embryo age: 1 dpf = 1 hr, 2 dpf = 1.5 hr, 3 dpf = 2 hr, and 4 dpf = 2-2.5 hr.
<ol start="5">
	<li>Stop digestion when no whole embryos are visible, yet solid pieces are still visible -&#160;this is essential to prevent overdigestion.</li>
	<li>Centrifuge tubes with dissociated embryos at 0.8-2.3 x g for 3-5 min to pellet cells.</li>
	<li>Remove supernatant from pelleted cells and wash 2x with CO2&#160;independent media.</li>
	<li>Add fresh CO2&#160;independent media to resuspend cells. 1 ml is typically used for preps of 10-20 embryos. The volume can be scaled based on embryo number.</li>
	<li>Using a P1000 pipette, pass embryo suspension through a 70 &#956;m filter . This helps remove unwanted debris from the prep.</li>
</ol>
Note 4: Embryo suspension can be filtered a second time through a 40 &#956;m filter to further remove unwanted debris. From a double filtration, recovery is approximately 800 &#956;l from a starting volume of 1 ml.
<ol start="10">
	<li>Add approximately 50-100 &#956;l of myofiber suspension to each poly-L-lysine coated coverslip.</li>
</ol>
Note 5: Perform step 2.9 in the Petri dishes with Parafilm on the bottom, previously prepared. The Parafilm keeps the myofiber suspension from running off the coated coverslips. Keep Petri dishes covered to prevent evaporation.
<ol start="11">
	<li>Allow myofibers to settle approximately 1 hr onto the coated coverslips at room temperature.</li>
</ol>
Note 6: Myofibers will begin to settle within 5-10 min. However, for good myofiber attachment, a minimum of 1 hr (1-2 hr) is recommended. Allowing myofibers to settle for longer will not harm the prep. For longer incubations (including overnight), antibiotics can be added to the media. With antibiotics and sterile technique live cultures can typically be maintained for 1-2 days.
<ol start="12">
	<li>Live myofibers can be observed at this point. Myofibers from embryos 2 dpf and later will be seen as striated and elongated cells (Figure 1). At this point, myofibers are now ready for live analysis or for immunolabeling.</li>
</ol>
Note 7: Skeletal muscle from 1 dpf embryos does not plate as elongated and clearly striated fibers. Instead, large myoballs are visible. In addition, during the pelleting phase post dissociation (step 2.6), 1 dpf myoballs need to be centrifuged for a minimum of 8 min at 5,000 x g to achieve a pellet. For analysis of embryos at this stage, it is recommended to use the transgenic line expressing EGFP specifically in skeletal muscle23. This will allow identification of cells from muscle origin versus other sources.
3. Fluorescence Immunostaining of Dissociated Zebrafish Myofibers (Time: approximately 1 day)
3.1.&#160;Immunolabeling
<ol>
	<li>Remove a portion of media from myofibers adhered to the coated coverslip</li>
	<li>Fix cells using 4% paraformaldehyde or methanol. For PFA, remove &#189; the volume of media and replace with the same amount of PFA. Fix for 10 min at RT, then remove total volume, replace with 50-100 &#956;l of PFA, and fix for an additional 10 min. For alcohol fixation, ice cold methanol or acetone can be applied at 4 &#176;C for 10 min or 5 min, respectively.</li>
	<li>Wash myofibers at least 3-5x with 1x PBS or 1x PBS plus 0.1% TWEEN. Average wash volume is 50-100 &#956;l per coverslip.</li>
	<li>Add blocking solution to myofibers (working stock) and incubate 20-60 min at room temperature. Blocking solution will vary depending on the primary and secondary antibodies. The two most common used in the lab are (1) 1x PBS, 2 mg/ml BSA, 1% sheep serum, 0.25% Triton X-100 final and (2) 0.2% Triton X-100, 2 mg/ml (0.2% BSA) and 5% sheep/goat serum.</li>
	<li>Remove blocking solution and add primary antibody diluted in either blocking solution or PBS. Incubate myofibers overnight at 4 &#176;C. Make sure to either Parafilm or wrap in saran wrap the Petri dish containing the myofiber-loaded coverslips coated with antibody. Small pieces of moistened paper towel can be added to create a humidified chamber.</li>
	<li>Remove primary antibody from coverslips and wash coverslips 3-5x for 5 min with PBS or blocking solution.</li>
	<li>Add secondary antibody at appropriate concentration diluted in 1x PBS or blocking solution for 1-2 hr at RT.</li>
	<li>Remove secondary antibody and wash myofibers 3-5x in PBS for 5 min each at RT.</li>
</ol>
3.2.&#160;Mounting coverslips
<ol start="9">
	<li>Apply 1-2 drops of antifade reagent to a microscope slide.</li>
	<li>Carefully pick-up the coated and immunolabled coverslip using forceps and place (myofibers down) the coverslip on the antifade reagent on the microscope slide.</li>
</ol>
Note 1: Attention to the orientation of the coated coverslip is critical.
<ol start="11">
	<li>Lay 1-2 Kimwipes on a hard solid surface. Quickly invert the slide with the coated coverslips resting on the antifade reagent and place it (coverslip down) on the Kimwipes.</li>
	<li>Apply light pressure to the corners of the microscope slide to remove excess antifade and form a tight seal between the slide and coverslip</li>
	<li>Allow the antifade remaining to dry for 5-10 min at RT, then the myofibers are ready to image (Figures 2A and&#160;B) or store at 4 &#176;C.</li>
</ol>
4. Live Cell Calcium Imaging Using GCaMP3
Live cell experiments can be performed on myofibers prior to fixation (following step 2.10). The following protocol describes live imaging in myofibers expressing GCaMP324, a genetically encoded calcium indicator, expressed by the skeletal muscle specific zebrafish &#945;-actin promoter (pSKM)23. Alternatively, this technique can be readily adapted to use calcium indicator dyes such as Fura-2.
<ol>
	<li>Inject embryos with pSKM: GCaMP3 construct at the 1-2 cell stage</li>
	<li>At 3 dpf, collect larvae and prepare myofibers as described in sections 1 and 2. Allow myofibers to adhere for a minimum of 1 hr. For this technique, preparing coverslips in 24-well dishes is required.</li>
	<li>Carefully remove any excess media, and add 300 &#956;l of fresh CO2&#160;independent media at RT.</li>
	<li>Observe cells under an inverted microscope. GCaMP3 positive myofibers should be visible under green fluorescence.</li>
	<li>Set up camera for recording, if desired.</li>
	<li>Prepare a 30 mM caffeine solution in CO2&#160;independent media. To stimulate cells, gently pipette 300 &#956;l of caffeine solution into the well under magnification. Within 5-10 sec, GCaMP positive myofibers should respond to caffeine with a rapid increase in fluorescence (Figure 3). Myofibers may also contract. Of note, caffeine induces calcium release from the sarcoplasmic reticulum stores25. Other agents, such as KCl26&#160;and ryanodine4, can be used to study inducible calcium release.</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>Schredelseker, J., Shrivastav, M., Dayal, A., Grabner, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-Ca2+-conducting+Ca2++channels+in+fish+skeletal+muscle+excitation-contraction+coupling.">Non-Ca2+-conducting Ca2+ channels in fish skeletal muscle excitation-contraction coupling.</a> Proc. Natl. Acad. Sci. U.S.A. 107, 5658-5663 (2010).</li></ol>

The authors declare no conflicting interests.

Zebrafish are a powerful vertebrate model system for studying muscle development and function&#160;in vivo25,27,28&#160;. They have also emerged as a valuable asset for modeling human muscle diseases14,15,20,29&#160;. While great strides have been taken to advance the use and application of zebrafish for the study of muscle function and muscle disease, there is a constant critical need to develop tools that will allow more in depth analysis that compliments the genetic, morphologic, behavio...

Skeletal muscle is a highly specialized tissue responsible for generating the contractile forces necessary for motility. Contraction is initiated through a process known as excitation-contraction (EC) coupling that converts electric signals to calcium release from intracellular stores1,2 . Intracellular calcium release activates the sarcomere to shorten and generate force. The many specific components of the molecular machinery responsible for mediating neuromuscular junction transmission3, EC coupling4,5 , and actin-myosin dependent contractions6 continue to be the ongoing subject of intense research. In addition, proteins that stabilize the muscle membrane during contraction7,8 and that mediate signaling between the myofiber and the extracellular matrix7,9 have been identified and studied in great detail.
Mutations in a number of genes important for muscle structure and function have been identified as causes of human skeletal muscle diseases (<a href="http://www.musclegenetable.org/" target="_blank">http://www.musclegenetable.org/</a>). These diseases, classified broadly as skeletal myopathies and muscular dystrophies based on clinical and histopathologic features, are associated with muscle weakness, lifelong disability, and early mortality10,11 . The zebrafish has proven to be an outstanding system for modeling and studying human skeletal muscle diseases8,12,13 . It has been employed to validate new gene mutations8, define new aspects of disease pathophysiology14,15 , and identify new therapeutic approaches15,16 . The power of the zebrafish for studying human muscle disease relates to the large number of offspring, the rapid development of muscle structure and function, the optical clarity of the zebrafish embryo, and the ease of genetic and pharmacologic manipulation of the developing zebrafish17.
We and others12,18,19 have recently developed a simple technique for the rapid and efficient isolation of myofibers from the developing zebrafish. This methodology has enabled the examination of myofibers in greater detail than can be provided by whole embryo analysis. The technique has been exploited for the characterization of protein subcellular localization20 as well as for the identification of important histopathologic characteristics as part of validation studies in newly developed disease models21. Furthermore, isolated myofibers can additionally be used for live imaging and for electrophysiological studies22, techniques that allow for the interrogation of key aspects of muscle function. The specific protocol for myofiber isolation, along with two examples of subsequent analytic experiments, is detailed in the remaining parts of this manuscript.

<table><tbody><tr><td>24-well culture plate</td><td>Corning</td><td>3524</td><td></td></tr><tr><td>10x PBS</td><td>Invitrogen Gibco</td><td>70011</td><td></td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; Independent medium</td><td>Invitrogen Gibco</td><td>18045</td><td></td></tr><tr><td>Collagenase Type II</td><td>Worthington Biochemical</td><td>LS004186</td><td>Lot 41H12764</td></tr><tr><td>Collagenase Type IV</td><td>Worthington Biochemical</td><td>L5004188</td><td></td></tr><tr><td>8% Paraformaldehyde</td><td>Electron Microscopy Sciences</td><td>157-8</td><td></td></tr><tr><td>Methanol</td><td>Sigma</td><td>322415</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>X100</td><td></td></tr><tr><td>BSA</td><td>Sigma</td><td>A2153</td><td></td></tr><tr><td>Sheep serum</td><td>Sigma</td><td>S3772</td><td></td></tr><tr><td>Goat serum</td><td>Sigma</td><td>G9023</td><td></td></tr><tr><td>Glass coverslips</td><td>Fischerbrand</td><td>12-545-82 12CIR-1D</td><td></td></tr><tr><td>Poly-L-Lysine</td><td>Sigma</td><td>P4707</td><td></td></tr><tr><td>Pronase</td><td>Sigma</td><td>P5147</td><td></td></tr><tr><td>40 &amp;mu;m Filter</td><td>BD Biosciences</td><td>352340</td><td></td></tr><tr><td>70 &amp;mu;m Filter</td><td>BD Biosciences</td><td>352350</td><td></td></tr><tr><td>Prolong Gold antifade reagent</td><td>Invitrogen</td><td>P36931</td><td></td></tr><tr><td>Anti-&amp;alpha;-Actinin antibody</td><td>Sigma</td><td>A5044</td><td></td></tr><tr><td>Anti-RYR antibody</td><td>Abcam</td><td>34C</td><td></td></tr><tr><td>Alexa Fluor antibody</td><td>Invitrogen</td><td>A-21425</td><td></td></tr><tr><td>TWEEN 20</td><td>Sigma</td><td>P1379</td><td></td></tr><tr><td>60 mm Petri dish</td><td>Fischerbrand</td><td>0875713A</td><td></td></tr><tr><td>Poly-L-Ornithine</td><td>Sigma</td><td>P4957</td><td></td></tr><tr><td>Microscope slide</td><td>Fischerbrand</td><td>12-550-15</td><td></td></tr><tr><td>Caffeine</td><td>Sigma</td><td>C0750</td><td></td></tr></tbody></table>

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.

Fluorescent immunolabeling of myofibers (Figure 2) 
Images showing expected fluorescent labeling pattern from myofibers immunostained after successful isolation and plating. The myofibers have been labeled with either anti-ryanodine receptor (1:100) (Figure 2A) or anti-&#945;-actinin (1:100) (Figure 2B) antibodies, and reveal immunostaining of the triad and the Z-band respectively. Secondary antibodies used were Alexa Fluor 555 (1:500...

Watch this Scientific Journal Video about Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations at JoVE.com

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

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 ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

11.6K Views.  University of Michigan. 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.

Video: Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

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.

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 ...

Developmental Biology

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 ...

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, ...

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 ...

Immunostaining of Dissected Zebrafish Embryonic Heart

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

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

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

Semi-automated Analysis of Mouse Skeletal Muscle Morphology and Fiber-type Composition

For years, distinctions between skeletal muscle fiber types were best visualized by myosin-ATPase staining. More recently, immunohistochemical staining of myosin heavy chain (MyHC) isoforms has emerged as a finer discriminator of fiber-type. Type I, type IIA, type IIX and type IIB fibers can now be identified with precision based on their MyHC profile; however, manual analysis of these data can be slow and down-right tedious. In this regard, rapid, accurate assessment of fiber-type composition and morphology is a very desirable tool. Here, we present a protocol for state-of-the-art immunohistochemical staining of MyHCs in frozen sections obtained from mouse hindlimb muscle in concert with a novel semi-automated algorithm that accelerates analysis of fiber-type and fiber morphology. As expected, the soleus muscle displayed staining for type I and type IIA fibers, but not for type IIX or type IIB fibers. On the other hand, the tibialis anterior muscle was composed predominantly of type IIX and type IIB fibers, a small fraction of type IIA fibers and little or no type I fibers. Several image transformations were used to generate probability maps for the purpose of measuring different aspects of fiber morphology (i.e., cross-sectional area (CSA), maximal and minimal Feret diameter). The values obtained for these parameters were then compared with manually-obtained values. No significant differences were observed between either mode of analysis with regards to CSA, maximal or minimal Feret diameter (all p &#62; 0.05), indicating the accuracy of our method. Thus, our immunostaining analysis protocol may be applied to the investigation of effects on muscle composition in many models of aging and myopathy.

Immunohistochemical staining of myosin heavy chain isoforms has emerged as the state-of-the-art discriminator of skeletal muscle fiber-type (i.e., type I, type IIA, type IIX, type IIB). Here, we present a staining protocol along with a novel semi-automated algorithm that facilitates rapid assessment of fiber-type and fiber morphology.

For years, distinctions between skeletal muscle fiber types were best visualized by myosin-ATPase staining. More recently, immunohistochemical staining of ...

Medicine

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 ...

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular ...

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

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

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

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