We thank the Animal House and the Integrated Microscopy Facilities of IGB-CNR. This work has benefited from research funding from the European Community's Seventh Framework Programme in the project ENDOSTEM (Activation of vasculature associated stem cells and muscle stem cells for the repair and maintenance of muscle tissue, grant agreement number 241440), the Italian Ministry of Education-University-Research (MIUR-PRIN2 010-2011) to G.M. and S.B. and PON Cluster IRMI to G.M., and the CARIPLO foundation to G.M. and S.B.

All experiments were conducted in strict accordance with the institutional guidelines for animal research and approved by the Department of Public Health, Animal Health, Nutrition and Food Safety of the Italian Ministry of Health in accordance with the law on animal experimentation.&#160;Cervical dislocation procedures may vary from institution to institution based on IACUC or its equivalent requirements.
1. Cardiotoxin Injection in the Tibialis Anterior Muscle
<ol>
	<li>Prepare a 10 &#956;M working solution of cardiotoxin (CTX) by diluting the cardiotoxin stock solution in sterile Phosphate-Buffered Saline (PBS) or in water before starting the procedure. 
	Note: CTX is soluble in water at 1 mg/mL. Prepare a 70 &#956;M stock solution. Aliquot and store at -20 &#176;C. CTX used for this protocol is a mixture of cardiotoxins having a molecular weight of approximately 7,000 Da. 
	CAUTION: It is suggested to wear double gloves, a hygiene mask, and safety glasses (alternatively to safety glasses, it is recommended to work under a hood) and to be very careful to avoid any contact with the solution, even if diluted.</li>
	<li>Anesthetize the mice by intraperitoneal injection of ketamine and xylazine (80 mg/kg and 10 mg/kg of body mass) or other approved available anesthetic.</li>
	<li>With the mouse face up, spray the hind limbs with 70% ethanol. To visualize the exact location of the Tibialis Anterior (TA) muscle, with the help of a scalpel, cut the hair away at the anterior part of the lower leg, under the knee. 
	Note: The TA muscle arises from the proximal body of the tibia, going down lateral to the bone and ending at the ankle. Its long tendon extends into the foot across the ankle (see Figure 1A).</li>
	<li>Draw ~ 100 &#956;L of the solution into the syringe by pulling the plunger back slowly.</li>
	<li>Remove air bubbles, holding the syringe upright (with the needle upward). Gently pull the plunger down and tap the side of the syringe to help any air bubbles attached to the inner part of the tube come upwards. Slowly press the plunger to release large air bubbles. A few drops of liquid will come out the needle. At this point, make sure to pick the liquid up again into the tube and not disperse it, given the toxicity of cardiotoxin.</li>
	<li>Withdraw the solution into the syringe until the desired dose is reached. Inject 20 &#956;L of CTX solution for each TA muscle.</li>
	<li>Insert the needle in the center of the TA by following the position of the Tibia bone as guidance, from the tendon towards the knee. Since the TA is a thin tissue, perform the injection without going too deep (insert the needle 2/3 mm deep, at approximately a 10&#176; to 20&#176; angle ), and avoid going beyond the muscle itself (Figure 1A).</li>
	<li>Do not pull out the needle immediately after the injection, but wait 2 - 3 s to prevent leakage of CTX drops from the muscle.</li>
	<li>After the procedure, place the cage on a heated plate (at 37 &#176;C) until the mice are awake, since the anesthetic and the ethanol used reduce body temperature.</li>
</ol>
2. Tibialis Anterior Isolation
Note: Muscles can be isolated at different time points after cardiotoxin injection according to experimental requirements.
<ol>
	<li>Sacrifice the mouse by cervical dislocation.</li>
	<li>Spray the hindlimbs with 70% ethanol. During the procedure, the animal can be fixed to a support to hold it down.</li>
	<li>Cut the skin at the level of the proximal tendon (above the foot), insert one of the blades of the scissors under the skin, and cut the skin upwards, in the direction of the knee. With the help of forceps, pull up the skin and expose the muscle entirely. Carefully remove all of the skin and the hair from the area of interest.</li>
	<li>Eliminate the fascia by slowly pinching or cutting (with thin-tip tweezers or with fine micro dissecting scissors) the extreme periphery of the muscle on the opposite side of the tibia (Figure 1B). Once broken, gently remove the fascia with the help of thin-tip tweezers; avoid damaging the underlying muscle. 
	Note: As with other muscles and organs, the TA is covered by fascia (deep fascia of the leg), a thin sheet of connective tissue that lies deep under the skin and that needs to be removed to easily isolate the underlying muscle.</li>
	<li>Visualize the distal TA tendon and the Extensor Digitorum tendon above the foot, which are juxtaposed. The tendon of the Extensor Digitorum Longus (EDL) muscle is located slightly below the TA tendon. Tendons of adult mice are visible to the naked eye.</li>
	<li>Remove the TA muscle following one of the two following techniques:
	<ol>
		<li>Cut both tendons with fine micro dissecting scissors and pull them towards the proximal side, holding the tendons with forceps. Separate the two tendons, pulling them gently in opposite directions to separate the muscles.</li>
		<li>With the help of thin-tip tweezers, separate the distal TA tendon from the Extensor Digitorum tendon by passing the tip below the tendon itself (Figure 1C, upper picture). Gently slide the tweezers below the TA to separate it from the underlying muscles (Figure 1C, bottom picture).</li>
		<li>Hold the tweezers under the muscle to keep it slightly raised, cut the TA tendon, and gently pull it upwards until only the area below the knee is attached (Figure 1D, left picture). Cut the TA below the knee, following the edge of the muscle with the scissors (Figure 1D, right picture). 
		Note: If the muscle remains slightly attached to the sides, use fine micro dissecting scissors to help detach it. 
		Note: If the forceps do not easily fit below the TA, it is possible that the fascia has not been completely removed, and it could still be visible on top of the muscle. In this case, completely remove the fascia before proceeding.</li>
	</ol>
	</li>
</ol>
3. Fresh Frozen Muscle Technique
<ol>
	<li>Place a small amount of tragacanth gum or a similar adhesive freezing compound (e.g., Optimal Cutting Temperature (OCT) compound) on a slice of cork. Label the cork&#39;s slice on the reverse side before freezing, if necessary (Figure 2A).</li>
	<li>In order to obtain transverse sections of the muscle, sink the TA into the tragacanth gum by inserting the distal tendon and leaving about 3/4 of the muscle outside, making sure to have the muscle in a perpendicular position with respect to the cork (Figure 2A).</li>
	<li>Fill an aluminum can (alternatively, a metal cup or a glass beaker), with isopentane. Suspend the can of isopentane in a Dewar containing liquid nitrogen, not allowing the liquid nitrogen to enter into the can. 
	Note: Isopentane needs to reach the proper temperature (from -150 to -160 &#176;C) for optimal freezing of the tissue. The correct temperature is reached when some white solid particles form at the bottom of the can and the isopentane becomes slightly viscous. 
	Note: Do not freeze the sample before the formation of the solid particles, as it can lead to freezing artifacts. On the other hand, if the isopentane becomes completely solid, leave it at room temperature until it is thawed, making sure that some white particles are still present.</li>
	<li>Using Mixter forceps, rapidly dip the cork with the muscle into the isopentane, keeping the muscle positioned downwards, and make sure to release the cork when it is completely immersed in the liquid. In this way, while the specimen floats, only the reverse side of the cork should be visible in the liquid.</li>
	<li>Leave the sample in the isopentane for 1 min.</li>
	<li>Using Mixter forceps (or similar forceps), take the muscle and immerse it in liquid nitrogen for at least 2 min.</li>
	<li>Wrap the specimens individually in labeled aluminum foil and immediately place them in dry ice (-70 &#176;C), or directly store them in a -80 &#176;C freezer. 
	Note: Throughout the procedure, from steps 3.4 to 3.7, it is extremely important to maintain the correct temperature and to prevent the specimens from thawing (e.g., during the transfer to the -80 &#176;C freezer). The rising temperature results in uncontrolled thawing and refreezing of the water microcrystals that are present within the tissue and eventually causes the formation of artifacts or the breakdown of muscle fibers (visualized on the tissue sections as large white holes in the center of the fiber).</li>
</ol>
4. Cryostat Sectioning of Frozen Muscles
<ol>
	<li>Set the cryostat chamber temperature to -20 to -22 &#176;C.</li>
	<li>Put the specimen stub, the blade, and the specimen in the cryostat chamber, allowing them to equilibrate for at least 30 min.</li>
	<li>Place a small amount of freezing compound (e.g., OCT compound) on the specimen stub and immerse the cork with the specimen before the freezing compound solidifies. In the meantime, place the blade in the blade holder.</li>
	<li>Align the specimen and the blade using the provided screws to orient the specimen holder with the blade holder (Figure 2B).</li>
	<li>Set the section thickness to 10 &#956;m. Proceed to cut the tissue. For each turn of the drive wheel, the specimen holder advances a controlled distance towards the blade.</li>
	<li>Place the sections on a polarized slide (8 to 10 section per slide) (Figure 2C).</li>
	<li>After sectioning, store the slides at -80 &#176;C. 
	Note: The specimen can be stored in a -80 &#176;C freezer and reused for further sectioning, if properly handled and stored.</li>
</ol>
5. Routine Histological Staining (Hematoxylin &#38; Eosin Stains)
Note: Several histological stains can be performed on muscle sections according to the analysis. A routine histological stain for morphological and morphometric analysis is the Hematoxylin &#38; Eosin (H&#38;E) bichromic stain. The hematoxylin stains the nuclei a deep purple. Nuclear staining is counterstained with eosin (pink/red), which stains eosinophilic structures, such as the myofibers in the cytoplasm.
<ol>
	<li>Completely air-dry frozen slides with sections for 10 - 15 min at room temperature, lodge the slides in a staining tray, and put the tray in a staining trough.</li>
	<li>Stain for 1 min with hematoxylin.</li>
	<li>Lay the staining trough under a fairly weak tap water jet for 10 min to wash out the hematoxylin.</li>
	<li>Immerse the slides for 5 min in 95% ethanol, if an alcoholic solution of eosin will be used (skip this step in case of an aqueous eosin solution).</li>
	<li>Counterstain for 1 min with eosin solution.</li>
	<li>Rinse with water to eliminate excessive eosin solution. 
	Note: Optimal timing of staining with either hematoxylin or eosin should be defined for each new batch.</li>
	<li>Dehydrate the samples in a graded ethanol series: 50% and 70% (a few seconds each). Rapidly immerse in 95% ethanol, followed by 2 changes in 100% ethanol, 5 min each.</li>
	<li>Clear in 2 changes of xylene, each 5 min or longer. This step must be carried out under a chemical hood.</li>
	<li>Mount the slides with a xylene-based mounting medium, applying a few drops of mounting medium on the slide and covering with a coverslip. The mounting medium can also be applied to the coverslip and the coverslip put on the slide.
	<ol>
		<li>Avoid forming air bubbles between the slide and the coverslip. To eliminate air bubbles, keep the slide vertical (after mounting) and squeeze out the excess mounting medium and air bubbles by gently pressing on the coverslip with blunt forceps (or a similar blunt tool).</li>
	</ol>
	</li>
	<li>Keep the slides under the hood for a few hours or overnight to let the xylene evaporate completely before proceeding to the analysis.</li>
	<li>Perform the morphological and morphometric analysis of H&#38;E-stained regenerating muscle sections by using non-overlapping serial images of at least one whole muscle section per mouse. Capture images with a bright-field microscope, with 20X magnification. 
	Note: A minimum number of 5 mice is recommended to obtain statistical significance of the measurements. 
	Note: Image analysis software is available for this purpose, such as the free download software ImageJ (http://imagej.nih.gov/ij/).</li>
</ol>

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The authors have nothing to disclose.

Here, we describe a protocol to induce acute injury in skeletal muscle (i.e., the intramuscular injection of CTX). It is widely used as a powerful tool to study the dynamics of skeletal muscle regeneration in vivo. CTX injection induces the degeneration of muscle fibers, which is caused by the depolarization of the sarcolemma and the contraction of the fibers12, and triggers the cascade of events that leads to muscle regeneration. Skeletal muscles are dissected at desired time points after the injection and injury, according to experimental requirements, and used for subsequent histological analyses. The dissected muscles can either be frozen, after inclusion in cryo-protectant media, or included in paraffin. However, these procedures require a step of pre-fixation with paraformaldehyde, which can generate artifacts and can eventually affect the subsequent analyses. Freezing the tissue directly can prevent this problem. Of note, many primary antibodies work exclusively or more effectively on fresh frozen muscle sections13, which makes this procedure more reliable. The morphometric analysis is usually performed on time course experiments, which allow for the evaluation of differences in the regeneration processes between groups of mice with the same age and with specific genetic mutations and/or pharmacological treatments. Although the protocol of CTX-induced damage is highly reproducible per se, a possible limitation is due to the operator-dependent variability of CTX injection. To overcome this limitation, it is preferable that the entire time course experiment is performed by the same operator.
The extent of muscle regeneration can be quantified as percentage of healthy fibers, necrotic areas and inflammation, and regenerating and regenerated areas over the total area. However, this approach is mostly used when studying events of chronic damage, such as in dystrophic mdx mice, rather than in models of acute injury. Indeed, while dystrophic muscles are characterized by asynchronous events of degeneration and regeneration, acute injury is followed by well-defined and consequential events. One limitation of these analyses, which require an empirical identification of the morphological features described above, is that they are not completely unbiased. To support experimental conclusions, the morphological analysis should always be complemented with further analysis, such as immunofluorescence analysis of specific molecular markers, to support experimental conclusions. For this reason, it is preferred to use parallel analysis to unequivocally interpret the results. For instance, regenerating myofibers are positively stained for embryonic Myosin Heavy Chain (eMyHC) that is expressed specifically in newly formed myofibers. Thus, quantification of eMyHC-stained myofibers can be performed side-by-side with the morphological analysis.
The CSA analysis is a more reliable quantification and can be performed either on H&#38;E-stained sections or on muscle sections stained with laminin, which marks the edge of the muscle fibers; quantification is performed using appropriate macros of ImageJ, as described above. In both cases, it is always necessary to first assess the quality of the sections and to exclude areas of tissue that appear deformed or curled.
Beside the morphological analyses of the tissues, H&#38;E-stained sections allow for the identification of other specific features, such as the presence of fibrosis and/or fatty tissue. Indeed, fibrosis derives from the excessive accumulation of extracellular matrix that occurs either if the regeneration is impaired or in chronic diseases14. Indeed, extracellular matrix accumulation is visible as pale material deposited between the regenerated myofibers in H&#38;E-histological staining. Multiple rounds of CTX injection can be used to mimic chronic disease in which the myofibers undergo waves of degeneration and regeneration and scar tissue accumulates aberrantly. However, more reliable protocols are available to induce muscle fibrosis15, and specific histological staining can be performed to identify and quantify these structures, such as Masson&#39;s Trichrome or Sirius Red staining. Formation of fatty tissue is clearly visible in H&#38;E-stained sections as rounded white structures between the myofibers, giving an important indication of the presence of fatty tissue, which is quantified by Oil Red staining. Finally, skeletal muscle sections obtained following the protocol described can be used to perform immunofluorescence analysis using specific antibodies and protocols.

Mammalian adult skeletal muscles are formed by groups of fascicules of multinucleated muscle cells (myofibers) that are specialized for contraction. Each myofiber is an elongated syncytium, surrounded by the sarcolemma (plasmatic membrane) and containing myofibrils, which are made up of regularly and repeatedly organized contractile proteins (actin and myosin filaments). In adult life and in resting conditions, skeletal muscles have a very low turnover of their myonuclei1; indeed, the myonuclei, which are located at the periphery of the myofiber, under the sarcolemma, are arrested in the G0 phase of the cell cycle and are unable to proliferate1,2.
Skeletal muscles have the peculiar ability to regenerate following damage, reaching homeostasis after several events of tissue remodeling that are tightly related to each other. After an acute injury or trauma, degeneration is induced, followed by regeneration processes that involve different cell populations, including a resident population of muscle cells, the satellite cells (SCs). Indeed, in the absence of any environmental stimuli, the satellite cells are in a quiescent state and are located in a specialized niche between the sarcolemma and the basal lamina3,4. Following an injury or disease, SCs become activated, proliferate, migrate to the damaged areas, and eventually differentiate, giving rise to newly forming myofibers5. Activated SCs establish cross-talk with different cell populations, mainly inflammatory cells, which are recruited in the site of trauma6-8. This cross-talk allows the cells to follow a regulated paradigm by which molecular signals drive structural modifications, eventually leading to homeostasis9. Besides SCs, inflammatory and interstitial cells, angiogenic processes, and re-innervation events are also involved, acting in a coordinated manner to repair this highly organized and specialized structure.
There is great interest in studying different aspects of skeletal muscle regeneration, not only to understand the physiology of the muscle, but also to improve therapeutic strategies that require deeper knowledge of the whole process. Several experimental approaches are currently available to study the identity and function of the different cell populations, the signaling pathways, and the molecular mechanisms involved. Mouse models of acute injury represent a powerful tool to investigate many aspects of this process. Different commonly used techniques to induce acute muscle damage allow researchers to follow the regeneration process in vivo, from the very early stages to the end of the process. This protocol describes the steps from the intramuscular injection of snake venom-derived cardiotoxin (CTX), which induces myolysis and triggers the regeneration process, up to the analysis of tissue samples. Following CTX injection, mice can be sacrificed at different time points depending on experimental requirements, and the skeletal muscles can be dissected and processed for further analysis. Finally, we describe the staining protocol of tissue sections to perform morphological observations and basic quantitative analyses. This protocol allows for the study of acute skeletal muscle regeneration in vivo in a highly reproducible manner10.

<table><tbody><tr><td>Cardiotoxin from&amp;nbsp;&lt;em&gt;Naja mossambica mossambica&lt;/em&gt;</td><td>SIGMA ALDRICH</td><td>C9759</td><td></td></tr><tr><td>Syringe For Insulin BD Micro-Fine+ Needle 30 G x 8 mm - Da 0.3 mL</td><td>BD</td><td>324826</td><td></td></tr><tr><td>Tragacanth Gum</td><td>MP BIOMEDICALS,LLC</td><td>104792</td><td></td></tr><tr><td>2-Methylbutane (Isopentane)</td><td>SIGMA ALDRICH</td><td>78-78-4.</td><td></td></tr><tr><td>OCT Killik Solution For Inclusion Cryostat</td><td>Bio-optica</td><td>&amp;nbsp;05-9801</td><td></td></tr><tr><td>Feather Microtome Blade S35</td><td>Bio-optica</td><td>&amp;nbsp;01-S35</td><td></td></tr><tr><td>Glass Slide Superfrost Plus</td><td>Menzel-Gl&amp;auml;ser</td><td>09-OPLUS</td><td></td></tr><tr><td>Dumon #5 Mirror Finish Forceps&amp;nbsp;</td><td>2BIOLOGICAL INSTRUMENTS</td><td>11251-23</td><td></td></tr><tr><td>Scissors Straight Sharp/Sharp</td><td>2BIOLOGICAL INSTRUMENTS</td><td>15024-10</td><td></td></tr><tr><td>Scissors Noyes Straight</td><td>2BIOLOGICAL INSTRUMENTS</td><td>15012-12</td><td></td></tr><tr><td>Fine Iris Scissors Straight Sharp/Sharp 10.5 cm</td><td>2BIOLOGICAL INSTRUMENTS</td><td>14094-11</td><td></td></tr><tr><td>Eukitt</td><td>Bio-optica</td><td>09-00100</td><td></td></tr><tr><td>Slide Coverslip</td><td>BIOSIGMA</td><td>VBS651</td><td></td></tr><tr><td>Xylene</td><td>SIGMA ALDRICH</td><td>214736</td><td></td></tr><tr><td>Ethanol 100%</td><td>sigma-Aldrich</td><td>02860-2.5L</td><td></td></tr><tr><td>Hematoxyline</td><td>J.T. BAKER</td><td>3873</td><td></td></tr><tr><td>Eosin</td><td>SIGMA ALDRICH</td><td>HT110116</td><td></td></tr><tr><td>Cryostat</td><td>LEICA</td><td>CM3050 S</td><td></td></tr></tbody></table>

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.

H&#38;E staining allows for the evaluation of the morphology of the regeneration process at specific time points during skeletal muscle regeneration. Figure 3 shows the time course analysis performed on injured TA muscles of wild type mice. Muscles have been isolated at 3, 7, 15, and 30 days after CTX injection, as schematized in Figure 3A. Representative pictures of H&#38;E-stained transverse sections show the dynamics of skeletal muscle repair over time (Figure 3B-E). At day 3 after injury, the structural architecture of the muscle is completely destroyed, and both degenerated myofibers (necrotic area) and mononucleated cells are clearly visible (Figure 3B). This heterogeneous group of cells mainly consists of transient amplifying satellite cells (myoblasts), inflammatory cells recruited from blood11, and interstitial and endothelial cells that participate in the regeneration process, which is triggered at the site of injury. In this context, new regenerating myofibers are generated by fusion and differentiation of myoblasts. The main feature of the regenerating area is the presence of small basophilic myofibers with centrally located nuclei and a dark-stained cytoplasm (Figure 3C). At this stage, inflammation is still visible, although it progressively decreases. The size of regenerating myofibers increases over time, giving rise to eosinophilic regenerated myofibers characterized by centrally located nuclei (Figure 3D, E). The regenerated myofibers are visible at later stages of regeneration and until the end of the process. The healthy myofibers appear highly organized and close to each other, with a pink cytoplasm and with the nuclei placed at the periphery of the fibers, under the sarcolemma. These myofibers are usually present far from the site of CTX injection and injury, and they more rarely represent areas where the regeneration is definitively complete. Indeed, regenerated fibers are characterized by the presence of centrally located nuclei that move to the periphery only several months after injury.
Digital images of the stained muscle sections are captured by adding the scale bars for spatial calibration, which is necessary to accurately measure the areas of interest by outlining the contours of the interested areas. The measurement of the myofiber cross-sectional area (CSA) is a reliable morphometric analysis, which is widely used to quantify the differences in the regeneration trend between different groups of mice. This analysis requires the measurement of areas of centrally nucleated myofibers, including at least 500 and up to 1,000 fibers per section. Both the average of the regenerating and regenerated myofiber areas and the Gaussian distribution of these areas are indicators of the regeneration process. Increased myofiber CSA is generally associated with an improved and/or accelerated regenerative response. On the contrary, failure of proper regeneration is associated with a decrease in CSA. Here, we reported the myofiber CSA analysis, indicated as both the average (Figure 3F) and distribution (Figure 3G) of myofiber CSA at different time points. The analysis highlights how the distribution of myofiber areas changes over time and how the CSA shifts towards larger-sized fibers as the process of regeneration proceeds.
<img alt="Figure 1" src="/files/ftp_upload/54515/54515fig1.jpg" /> 
Figure 1. Intramuscular Injection and Isolation of the Tibialis Muscle. A. Intramuscular injection of cardiotoxin in the Tibialis Anterior muscle. B. Removal of muscle fascia. The white arrow indicates the thin-tipped tweezers that pinch the muscle to remove the fascia. C. The top picture shows the anatomical position of the distal TA tendon. The bottom picture shows how to lift the TA and remove it while avoiding muscle damage. D. Left picture: the TA is lifted by pulling the distal tendon upward. The white arrow indicates the distal EDL tendon. Right picture: by holding the tendon with forceps, the top edge of the TA is cut off below the knee. The white arrow indicates the cutting site. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54515/54515fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54515/54515fig2.jpg" /> 
Figure 2. Fresh Freezing Muscle and Cryostat Sectioning of Frozen Muscles. A. Technique of fresh muscle inclusion in tragacanth gum. The TA muscle is immersed in the gum from the tendon, with about 3/4 outside and maintained in a perpendicular position with respect to the cork. B. Representative images of the procedure for cryostat sectioning. The cork is placed in a small amount of freezing compound on the specimen stub. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54515/54515fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54515/54515fig3.jpg" /> 
Figure 3. Time Course Analysis of Skeletal Muscle Regeneration. A. Experimental scheme of acute skeletal muscle injury. B-E. Representative pictures of Hematoxylin &#38; Eosin stained sections of cardiotoxin (CTX)-treated muscles at indicated days after injury. B. The black continuous line (on the left side of the picture), encloses a couple of necrotic fibers, likely invaded by inflammatory cells. The dashed line (right side of the picture) marks an area of infiltrating mononucleated cells. C. A single immature myofiber with centrally located nucleus is indicated by the black circle. D-E. Large regenerated eosinophilic myofibers, with centrally located nuclei are marked by black circles. Scale bars represent 100 mm. F. Average of centrally nucleated myofiber size values in TA muscle sections. Values are mean &#177; SEM, 5 mice/group. G. Myofiber cross-sectional area (CSA) distribution at 6, 15, and 30 days after CTX injection. Values are mean &#177; SEM, 5 mice/group. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54515/54515fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection at JoVE.com

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

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 ...

induction-acute-skeletal-muscle-regeneration-cardiotoxin

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

21.9K Views.  Institute of Genetics and Biophysics "A. Buzzati Traverso", CNR.  Dit manuscript beschrijft een gedetailleerd protocol voor acute skeletspier regeneratie bij volwassen muizen en latere manipulaties van de spieren, zoals dissectie, bevriezen, snijden, routine kleuring en myofiber dwarsdoorsnede analyse induceren. 

Video: Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional capacity. Transcutaneous surface stimulation of skeletal muscle involves a current flow between a cathode and an anode, thereby inducing excitement of the motor unit and the surrounding muscle fibers. 
NMES is an attractive modality to evaluate skeletal muscle adaptive responses for several reasons. First, it provides a reproducible experimental model in which physiological adaptations, such as myofiber hypertophy and muscle strengthening6, angiogenesis7-9, growth factor secretion9-11, and muscle precursor cell activation12 are well documented. Such physiological responses may be carefully titrated using different parameters of stimulation (for Cochrane review, see 13). In addition, NMES recruits motor units non-selectively, and in a spatially fixed and temporally synchronous manner14, offering the advantage of exerting a treatment effect on all fibers, regardless of fiber type. Although there are specified contraindications to NMES in clinical populations, including peripheral venous disorders or malignancy, for example, NMES is safe and feasible, even for those who are ill and/or bedridden and for populations in which rigorous exercise may be challenging. 
Here, we demonstrate the protocol for adapting commercially available electrodes and performing a NMES protocol using a murine model. This animal model has the advantage of utilizing a clinically available device and providing instant feedback regarding positioning of the electrode to elicit the desired muscle contractile effect. For the purpose of this manuscript, we will describe the protocol for muscle stimulation of the anterior compartment muscles of a mouse hindlimb.

A murine model of neuromuscular electrical stimulation (NMES), a safe and inexpensive clinical modality, to the anterior compartment muscles is described. This model has the advantage of modifying a readily available clinical device for the purpose of eliciting targeted and specific muscle contractions in mice.

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional ...

Medicine

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

Bioengineering

Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo

Skeletal muscle possesses an enormous capacity to regenerate after injury. This process is mainly driven by muscle stem cells, also termed satellite cells. Satellite cells are characterized by the expression of the transcription factor Pax7 and their location underneath the basal lamina in the resting skeletal muscle. Upon injury, satellite cells get activated, undergo self-renewal or differentiation to either form new myofibers or to fuse with damaged ones. The functionality of satellite cells in vivo can be investigated using a cardiotoxin based injury model of skeletal muscle. To study the function of one gene during the regeneration of skeletal muscle, transgenic mouse models are mostly used. Here, we present an alternative method to transgenic mice, to investigate the gene function in satellite cells during regeneration, e.g., in cases where transgenic mice are not available. We combine the cardiotoxin mediated injury of a specific skeletal muscle with the injection of a self-delivering siRNA into the regenerating muscle which is then taken up by satellite cells among other cells. Thereby, we provide a method to analyze gene function in satellite cells during regeneration under physiological conditions without the need for transgenic mice.

We describe an in vivo method to investigate the functional loss of a specific gene in satellite cells by using a combination of cardiotoxin mediated injury of skeletal muscle and the injection of a self-delivering siRNA.

Skeletal muscle possesses an enormous capacity to regenerate after injury. This process is mainly driven by muscle stem cells, also termed satellite ...

Genetics

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 ...

Human Skeletal Muscle Biopsy Procedures Using the Modified Bergstr&#246;m Technique

The percutaneous biopsy technique enables researchers and clinicians to collect skeletal muscle tissue samples. The technique is safe and highly effective. This video describes the percutaneous biopsy technique using a modified Bergstr&#246;m needle to obtain skeletal muscle tissue samples from the vastus lateralis of human subjects. The Bergstr&#246;m needle consists of an outer cannula with a small opening (&#8216;window&#8217;) at the side of the tip and an inner trocar with a cutting blade at the distal end. Under local anesthesia and aseptic conditions, the needle is advanced into the skeletal muscle through an incision in the skin, subcutaneous tissue, and fascia. Next, suction is applied to the inner trocar, the outer trocar is pulled back, skeletal muscle tissue is drawn into the window of the outer cannula by the suction, and the inner trocar is rapidly closed, thus cutting or clipping the skeletal muscle tissue sample. The needle is rotated 90&#176;&#160;and another cut is made. This process may be repeated three more times. This multiple cutting technique typically produces a sample of 100-200 mg or more in healthy subjects and can be done immediately before, during, and after a bout of exercise or other intervention. Following post-biopsy dressing of the incision site, subjects typically resume their activities of daily living right away and can fully participate in vigorous physical activity within 48-72 hr. Subjects should avoid heavy resistance exercise for 48 hr to reduce the risk of herniation of the muscle through the incision in the fascia.

The purpose of this video is to present the modified Bergstr&#246;m skeletal muscle biopsy technique on human subjects.

The percutaneous biopsy technique enables researchers and clinicians to collect skeletal muscle tissue samples. The technique is safe and highly ...

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from traumatic injury, disease and various congenital, genetic and acquired conditions are quite common. Tissue engineering and regenerative medicine technologies have enormous potential to provide a therapeutic solution. However, utilization of biologically relevant animal models in combination with longitudinal assessments of pertinent functional measures are critical to the development of improved regenerative therapeutics for treatment of VML-like injuries. In that regard, a commercial muscle lever system can be used to measure length, tension, force and velocity parameters in skeletal muscle. We used this system, in conjunction with a high power, bi-phase stimulator, to measure in vivo force production in response to activation of the anterior crural compartment of the rat hindlimb. We have previously used this equipment to assess the functional impact of VML injury on the tibialis anterior (TA) muscle, as well as the extent of functional recovery following treatment of the injured TA muscle with our tissue engineered muscle repair (TEMR) technology. For such studies, the left foot of an anaesthetized rat is securely anchored to a footplate linked to a servomotor, and the common peroneal nerve is stimulated by two percutaneous needle electrodes to elicit muscle contraction and dorsiflexion of the foot. The peroneal nerve stimulation-induced muscle contraction is measured over a range of stimulation frequencies (1-200 Hz), to ensure an eventual plateau in force production that allows for an accurate determination of peak tetanic force. In addition to evaluation of the extent of VML injury as well as the degree of functional recovery following treatment, this methodology can be easily applied to study diverse aspects of muscle physiology and pathophysiology. Such an approach should assist with the more rational development of improved therapeutics for muscle repair and regeneration.

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

We describe an in vivo protocol to measure dorsiflexion of the foot following stimulation of the peroneal nerve and contraction of the anterior crural compartment of the rat hindlimb. Such measurements are an indispensable translational tool for evaluating skeletal muscle pathology and tissue engineering approaches to muscle repair and regeneration.

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from ...

Minimally Invasive Muscle Embedding (MIME) - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new muscle fibers under the right conditions. Although SCs can be harvested from muscle tissue and cultured in vitro, the resulting myoblast cells are not very effective in promoting myogenesis when transplanted into host muscle. Surgically exposing the host muscle and grafting segments of donor muscle tissue, or the isolated muscle fibers with their SCs onto host muscle, promotes better myogenesis compared to myoblast transplantation. We have developed a novel technique that we call Minimally Invasive Muscle Embedding (MIME). MIME involves passing a surgical needle through the host muscle, drawing a piece of donor muscle tissue through the needle track, and then leaving the donor tissue embedded in the host muscle so that it may act as a source of SCs for the host muscle. Here we describe in detail the steps involved in performing MIME in an immunodeficient mouse model that expresses a green fluorescent protein (GFP) in all of its cells. Immunodeficiency in the host mouse reduces the risk of immune rejection of the donor tissue, and GFP expression enables easy identification of the host muscle fibers (GFP+) and donor-cell-derived muscle fibers (GFP-). Our pilot data suggest that MIME can be used to implant an extensor digitorum longus (EDL) muscle from a donor mouse into the tibialis anterior (TA) muscle of a host mouse. Our data also suggest that when a myotoxin (barium chloride, BaCl2) is injected into the host muscle after MIME, there is evidence of donor-cell-derived myogenesis in the host muscle, with approximately 5%, 26%, 26% and 43% of the fibers in a single host TA muscle showing no host contribution, minimal host contribution, moderate host contribution, and maximal host contribution, respectively.

Minimally Invasive Muscle Embedding MIME - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

We describe a novel experimental technique that we call Minimally Invasive Muscle Embedding (MIME), which is based on the evidence that skeletal muscle tissue contains viable myogenic cells that can facilitate donor-cell-mediated myogenesis when implanted into a host muscle.

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new ...

Isolation, Culture, and Transplantation of Muscle Satellite Cells

Muscle satellite cells are a stem cell population required for postnatal skeletal muscle development and regeneration, accounting for 2-5% of sublaminal nuclei in muscle fibers. In adult muscle, satellite cells are normally mitotically quiescent. Following injury, however, satellite cells initiate cellular proliferation to produce myoblasts, their progenies, to mediate the regeneration of muscle. Transplantation of satellite cell-derived myoblasts has been widely studied as a possible therapy for several regenerative diseases including muscular dystrophy, heart failure, and urological dysfunction. Myoblast transplantation into dystrophic skeletal muscle, infarcted heart, and dysfunctioning urinary ducts has shown that engrafted myoblasts can differentiate into muscle fibers in the host tissues and display partial functional improvement in these diseases. Therefore, the development of efficient purification methods of quiescent satellite cells from skeletal muscle, as well as the establishment of satellite cell-derived myoblast cultures and transplantation methods for myoblasts, are essential for understanding the molecular mechanisms behind satellite cell self-renewal, activation, and differentiation. Additionally, the development of cell-based therapies for muscular dystrophy and other regenerative diseases are also dependent upon these factors.

However, current prospective purification methods of quiescent satellite cells require the use of expensive fluorescence-activated cell sorting (FACS) machines. Here, we present a new method for the rapid, economical, and reliable purification of quiescent satellite cells from adult mouse skeletal muscle by enzymatic dissociation followed by magnetic-activated cell sorting (MACS). Following isolation of pure quiescent satellite cells, these cells can be cultured to obtain large numbers of myoblasts after several passages. These freshly isolated quiescent satellite cells or ex vivo expanded myoblasts can be transplanted into cardiotoxin (CTX)-induced regenerating mouse skeletal muscle to examine the contribution of donor-derived cells to regenerating muscle fibers, as well as to satellite cell compartments for the examination of self-renewal activities.

The isolation and culture of a pure population of quiescent satellite cells, a muscle stem cell population, is essential to the understanding of muscle stem cell biology and regeneration, as well as stem cell transplantation for therapies in muscular dystrophy and other degenerative diseases.&#160;

Muscle satellite cells are a stem cell population required for postnatal skeletal muscle development and regeneration, accounting for 2-5% of sublaminal ...

Biology

In Situ Immunofluorescent Staining of Autophagy in Muscle Stem Cells

Increasing evidence points to autophagy as a crucial regulatory process to preserve tissue homeostasis. It is known that autophagy is involved in skeletal muscle development and regeneration, and the autophagic process has been described in several muscular pathologies and age-related muscle disorders. A recently described block of the autophagic process that correlates with the functional exhaustion of satellite cells during muscle repair supports the notion that active autophagy is coupled with productive muscle regeneration. These data uncover the crucial role of autophagy in satellite cell activation during muscle regeneration in both normal and pathological conditions, such as muscular dystrophies. Here, we provide a protocol to monitor the autophagic process in the adult Muscle Stem Cell (MuSC) compartment during muscle regenerative conditions. This protocol describes the setup methodology to perform in situ immunofluorescence imaging of LC3, an autophagy marker, and MyoD, a myogenic lineage marker, in muscle tissue sections from control and injured mice. The methodology reported allows for monitoring the autophagic process in one specific cell compartment, the MuSC compartment, which plays a central role in orchestrating muscle regeneration.

In Situ Immunofluorescent Staining of Autophagy in Muscle Stem Cells

Active autophagy is associated with productive muscle regeneration, which is essential for Muscle Stem Cell (MuSC) activation. Here, we provide a protocol for the in situ detection of LC3, an autophagy marker in MyoD-positive MuSCs of muscle tissue sections from control and injured mice.

Increasing evidence points to autophagy as a crucial regulatory process to preserve tissue homeostasis. It is known that autophagy is involved in skeletal ...

A Massager Based on Ding's Roll Manipulation to Treat Muscle Injury in a Rat Model

Mimicking Ding's Roll Method on Notexin-Induced Muscle Injury in Rats

Ding's roll method is one of the most commonly used manipulations in traditional Chinese massage (Tuina) clinics and one of the most influential contemporary Tuina manipulations in China. It is based on the traditional rolling method commonly used in the one finger Zen genre and named Ding's roll method. Due to its anti-inflammatory and blood circulation-promoting effects, Ding's rolling method has sound therapeutic effects on myopathy. Because of the large area of force applied to human skin, Ding's roll method is challenging to perform on experimental animals with small skin areas, such as rats and rabbits. Additionally, the strength of Tuina applied to the human body differs from that applied to experimental animals, so it may happen that the strength is too high or too low to achieve the therapeutic effect of Tuina during the experiment. This experiment aims to create a simple massager suitable for rats based on Ding's rolling manipulation parameters (strength, frequency, Tuina duration). The device can standardize manipulation in animal experiments and reduce the variation in Tuina force applied to different animals due to subjective factors. A rat model of notexin-induced skeletal muscle injury was established, and plasma injury markers creatine kinase (CK) and fatty acid binding protein 3 (FABP3) were used to assess the therapeutic effect of Tuina on skeletal muscle injury. The results showed that this Tuina massager could reduce the levels of CK and FABP3 expression and slow down the degree of skeletal muscle injury. Therefore, the Tuina massager described here, mimicking Ding's roll method, contributes to standardizing Tuina manipulation in experimental research and is of great help for subsequent research on the molecular mechanism of Tuina for myopathy.

Author Spotlight: Performing Tuina Massage in Rats Using a Self-Made Massager 

This protocol describes a simple device that mimics Ding's roll method, establishes a rat model of skeletal muscle injury, and uses hematoxylin-eosin staining to observe the pathology of damaged tissue and enzyme-linked immunosorbent assay to detect changes in serum damage markers.

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

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