We would like to thank Dr. Alex Fulcher, and Monash Micro Imaging for assistance with microscope maintenance and setup. The Australian Regenerative Medicine Institute is supported by grants from the State Government of Victoria and the Australian Government. This work was funded by a Muscular Dystrophy Association (USA) project grant to P.D.C (628882).

Zebrafish maintenance was carried out as per the standard operating procedures approved by the Monash University Animal Ethics Committee under breeding colony license ERM14481.
1. Determination of the genotype of live embryos using an embryo genotyping platform.
<ol>
	<li>Anesthetize 3 days post fertilization (dpf) zebrafish embryos by adding tricaine methanesulfonate to a final concentration of 0.016% (v/v) in embryo medium (5 mM NaCl, 0.17mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4 in water). Wait for 10 minutes to ensure that the fish are completely anesthetized, evident when the fish stop swimming.</li>
	<li>Prepare the 24 chamber DNA extraction chip by peeling the clear protective film from the top surface of the chip (Figure 1A).</li>
	<li>Cut the tip of a 20 &#181;L filter tip to widen the opening. Using this, pick a single embryo in a 13 &#181;L volume of embryo medium and load into the first chamber of the chip. Repeat this process until embryos have been dispensed into each chamber. 
	NOTE: This step should be performed in an area with minimal air drafts, as high air flow may cause excessive evaporation of the fluid subsequently resulting in reduced genotyping sensitivity and survival of the embryos.</li>
	<li>Once the desired number of embryos have been loaded onto the DNA extraction chip, load it onto the zebrafish embryo genotyping platform. This is best achieved by placing in one side first, followed by the rest of it. 
	NOTE: Avoid adding too much pressure onto the platform as this may overstress and damage the underlying springs.</li>
	<li>Affix the magnetic platform lid over the chip, which will prevent evaporation of embryo media during the DNA extraction protocol, and close the platform lid.</li>
	<li>Set the base unit to 2.4 volts, 0.051 A, and 0.12 W and start the DNA extraction protocol by pressing the &#34;ON/OFF&#34; button. The platform should start vibrating, which can be assessed by gently touching the lid. The protocol should be run for 8 minutes. 
	NOTE: The vigorous vibration results in shedding of epidermal cells, from which genomic DNA is extracted.</li>
	<li>While the program is running, prepare a 24 well plate by adding 1 mL of embryo medium to each well, which is necessary to separate individual embryos while downstream genotyping assays are being performed.</li>
	<li>Additionally, appropriately label 8 well strip tubes, which will be used for collection of DNA material from each embryo.</li>
	<li>At the completion of the 8 minute protocol, press the ON/OFF button to stop the vibration of the platform.</li>
	<li>Open the lid of the platform, and gently remove the magnetic lid ensuring minimal pressure is applied to the platform.</li>
	<li>Carefully remove the DNA extraction chip from the platform and place it on a flat surface.</li>
	<li>From the first chamber, remove 10 &#181;L of embryo media surrounding the embryo and place it into the appropriate well of the 8-well strip tube. The media contains the genetic material from that embryo, which will be used for downstream assays. 
	NOTE: Avoid touching the embryo with the pipette tip during media collection, as this can damage the embryo.</li>
	<li>Using a plastic pipette, immediately add two drops of fresh embryo medium to the same chamber. Collect the embryo and move it to the first well of a 24 well plate.</li>
	<li>Repeat steps 1.12 and 1.13 for all the remaining chambers. At the completion of this, the 24 well plate containing live embryos can be moved to a 28 &#176;C incubator.</li>
	<li>Perform appropriate downstream genotyping assays to determine the genotype of the embryos. 
	NOTE: The DNA obtained from the embryo genotyping platform can be successfully amplified by PCR and can be used for subsequent analysis including sequencing, gel electrophoresis, or high-resolution melt-analysis14. To genotype the lama2 strain described in the representative results, PCR, restriction digestion and gel electrophoresis were used. Given that the concentration of DNA obtained from each embryo is low, it is suggested to utilise most, if not all, of the genetic material obtained for the downstream assay.</li>
	<li>Once the genotype of each larvae has been identified, transfer them to 90 mm Petri dishes, placing each genotype in a different dish. Fill the dish with 25 mL embryo medium and keep them in the 28 &#176;C incubator.</li>
</ol>
2. Performing muscle injury using a needle stab
<ol>
	<li>Once the larvae are 4 dpf, anesthetize them by adding tricaine methanesulfonate to a final concentration of 0.016% (v/v) in embryo medium. Wait for 10 minutes to ensure that the fish are completely anesthetized, evident when the fish stop swimming. 
	NOTE: To avoid bias, the identity of the genotype should be blinded to the investigator performing the stabs and downstream analyses.</li>
	<li>During this time, prepare 24 well plates by filling each well with 1 mL of fresh embryo medium, and set up the stereomicroscope with a black background and high-power light, to facilitate the visualization of the somite borders.</li>
	<li>Using a plastic pipette, transfer one anesthetized larva into a new Petri dish.</li>
	<li>Carefully remove excess embryo medium with a pipette, and under a dissecting microscope, orient the fish such that the head is on the left, tail on the right, dorsal region up and ventral region down (Figure 1B).</li>
	<li>Working under a dissecting microscope, use a 30-gauge needle to perform a quick but precise stab in the epaxial muscle located above the horizontal myoseptum. To ensure consistency in anterior-posterior position, aim for 1-2 somites located above the anal pore (Figure 1B). 
	NOTE: Avoid stabbing the neural tube and notochord, and work quickly to avoid the drying of the fish during the process.</li>
	<li>Using a plastic pipette, pour a drop of embryo medium on the stabbed zebrafish and carefully transfer it into a well of the 24 well plate.</li>
	<li>Repeat steps 2.3 to 2.6. until all the fish have been stabbed. 
	&#8203;NOTE: Several larvae can be stabbed together depending on the processing speed and proficiency of the investigator, as long as the fish do not dry out during the process.</li>
	<li>Once all the larvae have been stabbed, place the 24 well plate in the 28 &#176;C incubator until subsequent imaging is performed.</li>
</ol>
3. Imaging of muscle injury and recovery
<ol>
	<li>At 1 day post injury (1 dpi), anaesthetize the stabbed larvae by adding tricaine methanesulfonate to a final concentration of 0.016% (v/v) in embryo medium. Wait for 10 minutes to ensure that the fish are completely anesthetized, evident when the fish stop swimming. 
	NOTE: At 0 dpi, the wound site contains a large amount of cellular debris, which makes it difficult to quantify the extent of muscle injury (Supplementary Figure 1A-B). It takes approximately 18-20 hours for this debris to be cleared from the wound site, and as such it is more reliable to image larvae at 1 dpi, rather than 0 dpi, to determine the extent of injury elicited.</li>
	<li>Place a clean and empty glass bottom based dish on the stage of the polarizing microscope and set background using the integrated software. 
	NOTE: Do not use plastic Petri dishes to image birefringence, as they do not appropriately transmit refracted light. Depending on the polarized microscope used, additional settings may be required as per the manufacturer&#39;s guidelines.</li>
	<li>Having set the background, remove the glass bottom based dish from the microscope stage, and using a glass pipette, transfer the anesthetized, stabbed fish onto the glass bottom based dish.</li>
	<li>Carefully remove the excess of embryo medium using a pipette, and orient the fish as per 2.5 - the head is on the left, tail on the right, dorsal region up and ventral region down. 
	NOTE: Ensure the larvae is mounted as flat as possible, as uneven mounting results in different birefringence intensities across different somites.</li>
	<li>Place the glass bottom based dish with the anesthetized larvae on the microscope stage and image the muscle using polarized light (Figures 1C &#38; 1D). 
	NOTE: Depending on the type of microscope/polarized lens used, the fish orientation on its anterior-posterior axis can affect the overall birefringence15. Ensure to include at least 5 somites on either side of the injury site when imaging.</li>
	<li>Using a plastic pipette, pour a drop of embryo medium to rehydrate the fish and place the fish in a well of a 24 well plate filled up with embryo medium. 
	NOTE: It is important to perform the imaging as quickly as possible to prevent the fish from drying.</li>
	<li>Repeat steps 3.3. to 3.6. until all the fish have been imaged.</li>
	<li>When all images have been acquired, save them in .tiff format for subsequent analyses.</li>
	<li>Put the fish back into the 28 &#176;C incubator until performing subsequent imaging at 3 dpi (7 dpf).</li>
	<li>When the fish are 3 dpi, image the fish as per 3.3 to 3.8.</li>
	<li>Once all fish have been imaged, euthanize them by adding tricaine methanesulfonate to a final concentration of 0.2% (v/v) in embryo medium. Wait for at least 10 minutes to ensure that the fish are euthanized, evident by the loss of swimming ability, pumping of the gill covers, and lack of a flight response following touch.</li>
</ol>
4. Quantification of muscle regeneration
<ol>
	<li>Open a 1 dpi image on an imaging analysis software such as the freely available Image J software.</li>
	<li>Using the polygon tool, draw around the wound site, and measure the area and mean birefringence intensity of this region (Figures 1C &#38; 1D). Copy these values to cells D3 and E3 in the template provided (Supplementary Table 1).</li>
	<li>Draw two additional regions, each spanning 1-2 uninjured somites, and measure the area and mean birefringence intensities of each of these regions (Figures 1C &#38; 1D). Copy these values to cells D4-D5 and E4-E5 in the template provided (Supplementary Table 1). 
	NOTE: While it is preferable to select the same uninjured somites in the 1 dpi and 3 dpi images, the sporadic detachment of muscle fibres and subsequent reduction in birefringence in mutants may make this impossible. Therefore, in the event the same somites cannot be selected at 1 dpi and 3 dpi due to the reduction in muscle integrity in mutants, select two different but unaffected areas at each of the timepoints.</li>
	<li>Calculate the normalized birefringence for each region by dividing the mean birefringence intensity of that region by the area - displayed in column F in the template provided (Supplementary Table 1).</li>
	<li>Repeat steps 4.1-4.4 for all 1 dpi and 3 dpi images. 
	NOTE: When using the template provided (Supplementary Table 1), the 1 dpi area and mean birefringence intensity values should be inserted in columns D and E respectively, and the 3 dpi area and mean birefringence intensity values should be inserted in columns J and K respectively.</li>
	<li>For each time point, calculate the average normalized birefringence of the two uninjured regions. This value provides a reference point of uninjured muscle. 
	NOTE: In the template provided (Supplementary Table 1), this value is computed in columns G and M, for the 1 dpi and 3 dpi images respectively.</li>
	<li>Next, determine the extent of muscle injury at 1 dpi, by dividing the normalized birefringence of injury region by the average normalized birefringence of the uninjured regions (calculated in 4.6). 
	NOTE: Using the above detailed needle stab procedure, wildtype larvae typically show a normalized birefringence of 48.5 &#177; 14.3% at the wound site at 1 dpi, indicating that the birefringence has reduced by approximately 50% when compared to uninjured somites. When using the template (Supplementary Table 1), this value is computed as a percentage in column H.</li>
	<li>To determine the extent of muscle regeneration, divide the normalized birefringence of the injury region in the 3 dpi image, by the average normalized intensity of the uninjured regions at this stage. 
	NOTE: At 3 dpi, wildtype larvae typically show a normalized birefringence of 60 &#177; 15.3% at the wound site. Given that the normalized birefringence within the wound site at 1 dpi was 48.5 &#177; 14.3% (step 4.7), the increase in birefringence at 3 dpi to 60 &#177; 15.3% indicates a recovery of approximately 11.5%. When using the template (Supplementary Table 1), this value is computed as a percentage in column N.</li>
	<li>Keeping fish within each genotype separate, perform a paired t-test comparing the normalized birefringence of the wound site at 3 dpi (step 4.8) with that of 1 dpi (step 4.7). This will reveal the trajectory of muscle regeneration displayed by each fish in each genotype, and highlight if the extent of muscle regeneration displayed by each genotype has significantly altered.</li>
	<li>Finally, calculate the regenerative index by dividing the value obtained in step 4.8, which is the extent of muscle regeneration at 3 dpi, by the value obtained in step 4.7, which is the extent of muscle injury at 1 dpi. A regenerative index of 1 indicates that the injury at 1 dpi is comparable to 3 dpi and that muscle regeneration has not occurred; a value above 1 indicates that at 3 dpi new muscle has formed in the wound site highlighting that the muscle has regenerated; and a regenerative index of less than 1 highlights that the wound at 3 dpi is worse than 1 dpi, and that muscle regeneration in impaired. A t-test or a one-way ANOVA can be performed to statistically compare the extent of muscle regeneration between the different genotypes. When using the template provided (Supplementary Table 1), this value is calculated in column O. 
	NOTE: It is recommended to perform the entire experiment in triplicates, with fish from each experiment obtained from different biological parents, and each experiment performed on different days, to avoid any bias.</li>
</ol>

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three+muscular+dystrophies:+loss+of+cytoskeleton-extracellular+matrix+linkage.">Three muscular dystrophies: loss of cytoskeleton-extracellular matrix linkage.</a> Cell. 80 (5), 675-679 (1995).</li><li>Cerletti, M., 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=Highly+efficient,+functional+engraftment+of+skeletal+muscle+stem+cells+in+dystrophic+muscles.">Highly efficient, functional engraftment of skeletal muscle stem cells in dystrophic muscles.</a> Cell. 134 (1), 37-47 (2008).</li><li>Dumont, N. 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The authors have nothing to disclose.

Skeletal muscle regeneration is driven by obligate tissue resident muscle stem cells, whose function is altered in many muscle diseases such as muscular dystrophy, subsequently impeding the process of muscle regeneration. Here, we describe a high throughput protocol to examine muscle regeneration in live zebrafish models of muscle disease. The first step of the pipeline utilizes a embryo genotyping platform14, which is a user-friendly and accurate method to determine the genotype of live larvae, b...

Skeletal muscle accounts for 30-50% of human body mass, and is not only indispensable for locomotion, but it also serves as a critical metabolic and storage organ1. Despite being postmitotic, skeletal muscle is highly dynamic and retains a tremendous regenerative capacity following injury. This is attributed to the presence of tissue resident stem cells (also called satellite cells), located under the basal lamina of myofibers and marked by the transcription factors paired box protein 7 (pax7) and/or paired box protein 3 (pax3), among others2,3. Following injury, the satellite cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. The highly conserved cascade of pro-regenerative signals regulating satellite cell activation and robust muscle repair is&#160;affected in various conditions such as myopathies and homeostatic ageing4,5.
One such diverse group of myopathies is muscular dystrophy, characterized by progressive muscle wasting and degeneration6. These diseases are the consequence of genetic mutations in key proteins, including dystrophin and laminin-&#945;2 (LAMA2), responsible for the attachment of muscle fibers to the extracellular matrix7,8. Given that proteins implicated in muscular dystrophy play such a central role in maintaining muscle structure, for many years it was believed that a failure in this process was the mechanism responsible for disease pathogenesis9. However, recent studies have identified defects in the regulation of muscle stem cells and subsequent impairment in muscle regeneration as a second possible basis for the muscle pathology observed in muscular dystrophy10,11. As such, further studies are needed to investigate how an impairment in muscle stem cell function and associated niche elements contributes to muscular dystrophy.
Over the past decade, zebrafish (Danio rerio) has emerged as an important vertebrate model for disease modeling12. This is attributed to the rapid external development of the zebrafish embryo, coupled with its optical clarity, which allows the direct visualization of muscle formation, growth, and function. Additionally, not only is the development and structure of muscle highly conserved in zebrafish, they also display a highly conserved process of muscle regeneration13. Consequently, zebrafish represent an excellent system to study the pathobiology of muscle diseases, and explore how muscle regeneration is affected in it. To this end, we have developed a method that enables the timely study of skeletal muscle regeneration in zebrafish models of muscle disease. This high throughput pipeline involves a method to genotype live embryos14, following which a needle-stab injury is performed and the extent of muscle regeneration is imaged using polarizing light microscopy. The utilization of this technique will therefore reveal the regenerative capacity of muscle in zebrafish models of muscle disease.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>24 well plates</td>
			<td>Thermo Fischer</td>
			<td>142475</td>
			<td></td>
		</tr>
		<tr>
			<td>30 gauge needles</td>
			<td>Terumo</td>
			<td>NN-3013R</td>
			<td></td>
		</tr>
		<tr>
			<td>90 mm Petri Dishes</td>
			<td>Pacific Laboratory Products PT</td>
			<td>S9014S20</td>
			<td></td>
		</tr>
		<tr>
			<td>DNA extraction chips</td>
			<td>wFluidx</td>
			<td>ZEG chips</td>
			<td></td>
		</tr>
		<tr>
			<td>Embryo genotyping platform</td>
			<td>wFluidx</td>
			<td>ZEG base unit</td>
			<td>Zebrafish Embryo Genotyper</td>
		</tr>
		<tr>
			<td>Glass pipette</td>
			<td>Hirschmann</td>
			<td>9260101</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass plate dish</td>
			<td>WPI</td>
			<td>FD35-100</td>
			<td>Commonly referred to as FluoroDish</td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>Thermoline Scientific</td>
			<td>TEI-43L</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic pipette</td>
			<td>Livingstone</td>
			<td>PTP03-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Polarizing microscope</td>
			<td>Abrio</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The ability to quantify birefringence of skeletal muscle provides a non-invasive but highly reproducible method to examine and compare levels of muscle damage, and examine muscle regeneration in vivo. Birefringence results from the diffraction of polarised light through the pseudo-crystalline array of the muscle sarcomeres15, and following injury or damage to the muscle, a reduction in birefringence is evident. Likewise, the activation and differentiation ...

Watch this Scientific Journal Video about Examining Muscle Regeneration in Zebrafish Models of Muscle Disease at JoVE.com

Examining 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, ...

examining-muscle-regeneration-in-zebrafish-models-of-muscle-disease

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Developmental Biology

4.8K Views.  Monash University. 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.

Video: Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Kidney Regeneration in Adult Zebrafish by Gentamicin Induced Injury

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is the nephron. Mammals are able to repair existing nephrons after injury, but lose the ability to form new nephrons soon after birth. In contrast to mammals, adult fish produce new nephrons (neonephrogenesis) throughout their lives in response to growth requirements or injury. Recently, lhx1a has been shown to mark nephron progenitor cells in the adult zebrafish kidney, however mechanisms controlling the formation of new nephrons after injury remain unknown. Here we show our method for robust and reproducible injury in the adult zebrafish kidney by intraperitoneal (i.p.) injection of gentamicin, which uses a noninvasive visual screening process to select for fish with strong but nonlethal injury. Using this method, we can determine optimal gentamicin dosages for injury and go on to demonstrate the effect of higher temperatures on kidney regeneration in zebrafish.

Here we present a reliable method to study adult kidney regeneration by inducing acute kidney injury by gentamicin injection. We show that injury is dependent on gentamicin dosage and environmental temperature using in situ hybridization to label lhx1a+ developing new nephrons.

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is ...

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 ...

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 ...

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 ...

Preparation and Culture of Myogenic Precursor Cells/Primary Myoblasts from Skeletal Muscle of Adult and Aged Humans

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting occurs. Loss of muscle mass and function is associated with muscle fiber type atrophy, fiber type switching, defective muscle regeneration associated with dysfunction of satellite cells, muscle stem cells, and other pathophysiological processes. These changes are associated with changes in intracellular as well as local and systemic niches. In addition to most commonly used rodent models of muscle ageing, there is a need to study muscle homeostasis and wasting using human models, which due to ethical implications, consist predominantly of in vitro cultures. Despite the wide use of human Myogenic Progenitor Cells (MPCs) and primary myoblasts in myogenesis, there is limited data on using human primary myoblast and myotube cultures to study molecular mechanisms regulating different aspects of age-associated muscle wasting, aiding in the validation of mechanisms of ageing proposed in rodent muscle. The use of human MPCs, primary myoblasts and myotubes isolated from adult and aged people, provides a physiologically relevant model of molecular mechanisms of processes associated with muscle growth, atrophy and regeneration. Here we describe in detail a robust, inexpensive, reproducible and efficient protocol for the isolation and maintenance of human MPCs&#160;and their progeny&#160;&#8212; myoblasts and myotubes from human muscle samples using enzymatic digestion. Furthermore, we have determined the passage number at which primary myoblasts from adult and aged people undergo senescence in an in vitro culture. Finally, we show the ability to transfect these myoblasts and the ability to characterize their proliferative and differentiation capacity and propose their suitability for performing functional studies of molecular mechanisms of myogenesis and muscle wasting in vitro.

This protocol describes a robust, reproducible and simple method of isolation and culture of myoblast progenitor cells from the skeletal muscle of adult and aged people. The muscles used here include foot and leg muscles. This approach enables the isolation of an enriched population of primary myoblasts for functional studies.

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting ...

Evaluation of Injury-induced Senescence and In Vivo Reprogramming in the Skeletal Muscle

Cellular senescence is a stress response that is characterized by a stable cellular growth arrest, which is important for many physiological and pathological processes, such as cancer and ageing. Recently, senescence has also been implicated in tissue repair and regeneration. Therefore, it has become increasingly critical to identify senescent cells in vivo. Senescence-associated &#946;-galactosidase (SA-&#946;-Gal) assay is the most widely used assay to detect senescent cells both in culture and in vivo. This assay is based on the increased lysosomal contents in the senescent cells, which allows the histochemical detection of lysosomal &#946;-galactosidase activity at suboptimum pH (6 or 5.5). In comparison with other assays, such as flow cytometry, this allows the identification of senescent cells in their resident environment, which offers valuable information such as the location relating to the tissue architecture, the morphology, and the possibility of coupling with other markers via immunohistochemistry&#160;(IHC). The major limitation of the SA-&#946;-Gal assay is the requirement of fresh or frozen samples.
Here, we present a detailed protocol to understand how cellular senescence promotes cellular plasticity and tissue regeneration in vivo. We use SA-&#946;-Gal to detect senescent cells in the skeletal muscle upon injury, which is a well-established system to study tissue regeneration. Moreover, we use&#160;IHC to detect Nanog, a marker of pluripotent stem cells, in a transgenic mouse model. This protocol enables us to examine and quantify cellular senescence in the context of induced cellular plasticity and in vivo reprogramming.

Evaluation of Injury-induced Senescence and In Vivo Reprogramming in the Skeletal Muscle

Here we present a detailed protocol to detect both senescent and pluripotent stem cells in the skeletal muscle upon injury while inducing in vivo reprogramming. This method is suitable for evaluating the role of cellular senescence during tissue regeneration and reprogramming in vivo.

Cellular senescence is a stress response that is characterized by a stable cellular growth arrest, which is important for many physiological and ...

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, ...

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from animals to humans. Therefore, the powerful Drosophila model system can be used to study concepts of muscle-tendon development that can also be applied to human muscle biology. Here, we describe in detail how morphogenesis of the adult muscle-tendon system can be easily imaged in living, developing Drosophila pupae. Hence, the method allows investigating proteins, cells and tissues in their physiological environment. In addition to a step-by-step protocol with helpful tips, we provide a comprehensive overview of fluorescently tagged marker proteins that are suitable for studying the muscle-tendon system. To highlight the versatile applications of the protocol, we show example movies ranging from visualization of long-term morphogenetic events &#8211; occurring on the time scale of hours and days &#8211; to visualization of short-term dynamic processes like muscle twitching occurring on time scale of seconds. Taken together, this protocol should enable the reader to design and perform live-imaging experiments for investigating muscle-tendon morphogenesis in the intact organism.

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Here, we present an easy-to-use and versatile method to perform live imaging of developmental processes in general and muscle-tendon morphogenesis in particular in living Drosophila pupae.

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from ...

X-ray Diffraction of Intact Murine Skeletal Muscle as a Tool for Studying the Structural Basis of Muscle Disease

Transgenic mouse models have been important tools for studying the relationship of genotype to phenotype for human diseases including those of skeletal muscle. Mouse skeletal muscle has been shown to produce high quality X-ray diffraction patterns on third generation synchrotron beamlines providing an opportunity to link changes at the level of the genotype to functional phenotypes in health and disease by determining the structural consequences of genetic changes. We present detailed protocols for preparation of specimens, collecting the X-ray patterns and extracting relevant structural parameters from the X-ray patterns that may help guide experimenters wishing to perform such experiments for themselves.

We present detailed protocols for performing small-angle X-ray diffraction experiments using intact mouse skeletal muscles. With the wide availability of transgenic mouse models for human diseases, this experimental platform can form a useful test bed for elucidating the structural basis of genetic muscle diseases

Transgenic mouse models have been important tools for studying the relationship of genotype to phenotype for human diseases including those of skeletal ...