We are grateful to Liam Tyron for his expert reading of the manuscript. This work was supported by funding from the Natural Sciences and Engineering Research Council of Canada (NSERC) to D. A. Hood. D. A. Hood is also the holder of a Canada Research Chair in Cell Physiology.

All animal-related procedures were reviewed and approved by the York University Animal Care Committee. Upon arrival at the animal facility at York University, all rats were given a minimum of five days to acclimatize to their environment prior to the surgical procedure, with food provided ad libitum. Although this protocol has been previously applied to other species15,17,22, the current paper builds on the pioneering work of Pette and colleagues23 and focuses on the rat model, specifically.
1. Preparation of Chronic Contractile Activity Unit
<ol>
	<li>Using a voltmeter, check the potential of the coin lithium batteries. 
	NOTE: The potential of each battery should be 3.0 &#177; 0.10 V.</li>
	<li>Insert two or three batteries into the slot of the unit so that the total potential is 6 - 9 V. 
	NOTE: It is up to the discretion of the researchers to consider how much potential (6 or 9 V) to maintain during the whole experimental procedure. Depending on the study design, and the desired intensity of the stimulus, either 2 or 3 batteries can be used.</li>
	<li>Verify that the unit is working properly via the pulse indicator, by exposing the unit to 1 pulse emitted by a portable infrared strobe light.</li>
</ol>
2. Surgical Procedure of Chronic Contractile Activity
NOTE: Sterilize all surgical tools before the surgical procedure. Both during and immediately after the surgery, the body temperature of the rats is maintained by a heating pad.&#160;It is desirable to perform the surgical procedure on a surgical drape. The surgeon should wear sterile surgical gloves as well as a clean lab coat. If needed, it is recommended to wear a disposable respirator mask.
<ol>
	<li>Anesthetize rats under 1 - 3% isoflurane inhalation with oxygen, which is operated by a gas vaporizer system. Confirm animal is fully sedated by checking hindlimb toe pinch and by observing respiratory depth and rate. Apply ocular lubricant on eyes to avoid dryness.&#160;Apply a subcutaneous injection of Meloxicam (0.5 mg/mL) at 2 mg/kg. 
	NOTE: It is also recommended to have a multi-modal application of analgesia (e.g., Meloxicam plus lidocaine) to minimize any pain during and after the surgery.</li>
	<li>Gently shave the left hindlimb, as well as a strip around the torso from the back of the neck, around behind of the forelimbs and across the anterior thorax. Gently wipe shaved areas with iodine and ethyl alcohol to disinfect.</li>
	<li>With the animal laying on its stomach, make a small incision (~0.5 cm) on the back of the neck in the center of the shaved region (the palpable area between shoulder blades) using a scalpel (No. 10 blade).</li>
	<li>Locate the common peroneal nerve.
	<ol>
		<li>Roll animal onto its right side and make a ~2 - 3 cm-long cut in the skin of the left hindlimb. Target the incision area around the upper thigh muscle groups that is landmarked between the dimple of the knee joint and the back close to the origin of the tail. 
		NOTE: Be careful not to contaminate the first incision area when changing the position of body.</li>
		<li>Using blunt-tipped curved surgical scissors, dissect the subcutaneous area widely ~3.5 - 4 cm, separating the skin from underlying muscle in order to make a pocket between the opened skin and underlying muscle. Open the skin from underlying tissue around the entire circumference of the incision (~1.5 cm2).</li>
		<li>Make a small incision (less than 0.5 cm) on the biceps femoris muscle with surgical scissors, ensuring that the tips of the scissors are cutting directly down through the muscle.</li>
		<li>Gently open the cut area until the internal muscle groups and the common peroneal nerve are visible (the depth of external muscle tissues (i.e., biceps femoris) is around ~0.5 cm). Using forceps, gently touch/pinch the visible nerve and observe responses of the target muscle groups (e.g., TA muscle) and toes (visible dorsiflexion) to confirm that the common peroneal nerve is isolated. 
		NOTE: This step should be done with extreme caution to avoid cutting or damaging the nerve.</li>
		<li>Fix the window by pulling it open with metal retractors such that the size of the window is ~1.5 cm2 with the peroneal nerve lying in the center of the window. Use small metal hooks attached to straps and/or rubber bands that are fastened down to the table surface (or surgery board) (Figure 2A).</li>
	</ol>
	</li>
	<li>Attach the wire to both sides of the nerve.
	<ol>
		<li>Prepare ~50 - 60 cm of polytetrafluoroethylene (PTFE)-coated fine stainless-steel wire and fold it in half.&#160; 
		NOTE: It may be helpful to expose PTFA-coated wire under the UV light before the surgery.</li>
		<li>Hook the folded part of the wire into the slit of a 30 cm-long stainless-steel rod. Pass the rod, along with the wire, subcutaneously from the open pocket of the hindlimb towards the small incision area at the back of the neck, in an L-shape pattern up the leg and along the center of the back.</li>
		<li>Find the two ends of the wire at the hindlimb. Strip the wires since all wires are PTFE-insulated. Using a scalpel, carefully strip the ends of the wire by ~1.5 cm. If the wires become frayed, cut them out and re-strip. Wrap the stripped wire ends around a blunted 21-G needle (5 times), making a coil. Once the coils are properly made, release the needle from them.</li>
		<li>Using a size 6-0 surgical silk, secure each of the coils on either side of the common peroneal nerve (Figure 2A).
		<ol>
			<li>Make a knot at the very end of the coil, and suture it on the left-hand side of the nerve. Ensure that the coil is at 1.5 - 2.5 mm from the nerve.</li>
			<li>To secure the coil, apply two or three additional sutures along the coil.</li>
			<li>Repeat these steps on the right-hand side of the nerve.</li>
		</ol>
		</li>
		<li>Apply 2 - 3 drops of antibiotic solution (ampicillin in saline; 132 mg/mL), and then carefully suture the window (i.e., biceps femoris muscle tissue) using size 5-0 silk.</li>
	</ol>
	</li>
	<li>Loosely wind the remaining slack of wire (about the diameter of the index finger) and push into the subcutaneous pocket above the sutured incision of the biceps femoris muscle (approximately above the hip).</li>
	<li>Apply 2 - 3 drops of antibiotic solution again (ampicillin in saline; 132 mg/mL). Close the opened skin by stapling.</li>
	<li>Connect the wires (coming out of the incision area of the neck) to the CCA stimulator.
	<ol>
		<li>Connect pin sockets with the wires.
		<ol>
			<li>Cut the wire loop coming out of the incision at the top of the neck to create 2 wire ends (these lead to the coils sutured to either side of the peroneal nerve).</li>
			<li>Using a scalpel, strip off the ends of the wires by ~0.5 cm. Cut off frayed wires, if any.</li>
			<li>Slowly push the stripped parts of the wires into the hole of the pin sockets, and using a soldering iron, solder the wires on the pin sockets.</li>
		</ol>
		</li>
		<li>Optionally, check the connection of wires.
		<ol>
			<li>Connect the pins to a large benchtop stimulator unit via alligator clips.</li>
			<li>Deliver a single pulse of 9 V (0.1 ms, 10 Hz) to confirm that TA muscle contracts and the left foot dorsiflexes.</li>
		</ol>
		</li>
		<li>Pass the pin connected-wire ends through the sterile gauze pad that is ~4 cm x 4 cm.</li>
		<li>Connect the pins to the CCA stimulator unit.
		<ol>
			<li>Pass the wires through the hole in the base of the stimulator box. 
			NOTE: This box is a home-made chamber for the CCA stimulator unit and is 3.5 cm &#215; 3.5 cm &#215; 2.5 cm13.</li>
			<li>Insert the pins into the connection sockets on the CCA unit. Gently put the CCA unit into the chamber. Use a sticky tack to secure the CCA unit at the bottom of the chamber.</li>
		</ol>
		</li>
		<li>Using an athletic tape or the porous surgical tape, fix the chamber with tape around the shaved torso. Close the top of the chamber with three layers of taping and finish by wrapping a tape around the sides of the stimulator box to secure the box (Figure 2B).</li>
	</ol>
	</li>
	<li>Check if the CCA is working by exposing the unit to a single pulse of infrared light (wavelength spectrum &#62;770 nm) that is emitted by a portable infrared strobe light. 
	NOTE: If the CCA is properly working, researchers will be able to see the hindlimb muscles (i.e., TA) are contracting in response to the infrared light.</li>
	<li>Observe the rat and monitor its temperature until it has fully regained consciousness. House it in a single-occupant cage to prevent any harm from other animals and do not leave any tunnels or plastic objects in the cage for the remainder of the study to mitigate the risk of damage to the stimulator unit or injury to the animal. Supply with an Amoxicillin-included water bottle (0.5 mg/mL).</li>
	<li>Apply 1 mg/kg dose of Meloxicam subcutaneously every 24 h following the surgery, which continues at least for 72 h.</li>
</ol>
3. Chronic Contractile Activity
<ol>
	<li>After the surgery, allow at least 5 - 7 days for a full recovery of incision and suture areas in/around the skeletal muscles. 
	NOTE: During and after CCA procedure, thoroughly check each animal&#39;s condition by observing their behaviors (e.g., eating, drinking, and/or moving). In addition, determine any severe stress or adverse effects by checking a change of body weight before and after the CCA procedure.</li>
	<li>On the day of CCA stimulation, turn on CCA by exposing the stimulator unit to a single pulse of infrared light (wavelength range &#62;770 nm) by a portable infrared strobe light.</li>
	<li>Apply 3 or 6 h of 10 Hz CCA stimulation. 
	NOTE: The time frame for stimulation is up to the researcher. The frequency of stimulation has never been altered in these experiments and only very modest enhancements in the mitochondrial adaptation have been observed by extending the stimulation from 3 to 6 h/day, in our experience. If possible, check the stimulation and animal every 30 - 60 min.</li>
	<li>Following the desired CCA period, turn off the CCA unit via infrared light exposure (same procedure as turning on the unit).</li>
	<li>If applying multiple days, repeat the step 3.2. to 3.4.</li>
	<li>Determine the timing of tissue collection. For example, collect tissues 24 h after the onset of the final bout of CCA (i.e., 18 h after the last CCA stimulation of 6 h), which is conducted under the anesthesia as done during the CCA surgical procedure. Immediately after collecting all tissues, euthanize animals by excising the heart while the animal is still under anesthetic.</li>
</ol>

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The authors have nothing to disclose.

The chronic contractile activity (CCA) model of exercise, through low-frequency muscle stimulation in vivo, is an excellent model for studying muscle phenotypic adaptations to exercise13,24,25,26. As shown in previous studies20,27, CCA is an effective tool by which researchers can control training volumes and frequencies (i.e., time and days) and investigate various biochemical and/or molecular events over the course of repeated bouts of contractile activity. One of the best features of this model is that muscles of the contralateral hindlimb are used as an internal control, which helps to minimize variability between animals. Furthermore, our numerous experiments over a decade have shown that the external CCA unit does not limit an animal&#39;s ordinary behavioral patterns (e.g., roaming, eating, and sleeping), and allows for the use of additional experimental treatments such as drug injections. Thus, researchers can apply the CCA protocol in their training studies with their own experimental modifications.
This CCA protocol has several critical steps, although all steps may demand a high-level concentration due to a nature of survival surgery. It is especially important to connect wires at consistent locations, and it is highly recommended to have the same skillful researcher perform the surgery to maintain consistency in landmarking the same spot of the nerve, suturing wires the same distance from the nerve, etc. In addition, it is necessary to confirm the security of the taping before and during the CCA application, because loosened or worn out tape may lead to a failure of the procedure.
This CCA model has some minor limitations. Since the CCA stimulation unit is fixed by taping, some animals exhibit minor skin irritations around the taping area. This could be improved in future studies through the use of a wearable jacket that would replace the CCA chamber without taping, however, we have not had success with such jackets. Alternatively, the stimulator can be implanted in the intraperitoneal space to avoid the taping procedure28, though this procedure is more invasive. In addition, the described external stimulator unit is designed to be controlled by exposing to infrared light. However, if a unit fails to respond to infrared light, this may indicate changes in the durability or sensitivity of the unit after long-time usage. The implementation of a microchip may ultimately avoid the use of the infrared light and would allow all CCA parameters to be programmed and stored, enabling CCA to be applied in a more controlled and accurate fashion. All study designs should also consider the use of contralateral limb sham surgery to exclude any possible outcomes resulting from the surgical procedure itself.
It is worthwhile to continue investigating&#160;how CCA can regulate muscle mass and phenotype following periods of chronic muscle disuse, or in the context of other muscular diseases. As shown in recent clinical studies29,30, CCA can also be applied to examine its efficacy on anabolic signaling mechanisms in aging populations. In conclusion, we encourage researchers to take advantage of using this CCA protocol in their studies, whereby they can investigate the various cellular and molecular mechanisms underlying skeletal muscle phenotypic adaptations to chronic exercise.

Skeletal muscle can adapt to exercise training through changes in its bioenergetics and physical structure1. One of the major alterations brought about by endurance training is mitochondrial biogenesis, which can be evaluated by an increase in the expression of mitochondrial components (e.g., cytochrome c oxidase [COX] subunits), as well as the expression of the transcriptional coactivator, PGC-1&#945;2. A growing number of studies have indicated that numerous other factors, including mitochondrial turnover and mitophagy, are also important for muscle adaptations. However, the mechanisms by which acute or chronic exercise regulate these processes in skeletal muscle are&#160;still unclear.
To delineate the pathways which regulate exercise-induced muscle adaptations, various exercise models have been commonly used in rodent studies, including treadmill, running wheel, and swimming exercise. However, these protocols have some limitations in that ~4 - 12 weeks are needed to observe these phenotypic changes3,4,5. As an alternative experimental method, low-frequency stimulation-induced chronic contractile activity (CCA) has been effectively used, as it can lead to muscle adaptations in a substantially shorter period (i.e., up to 7 days) and its effects appear to be comparable to, or even greater than other exercise protocols. Furthermore, the presence of hormonal6, temperature7, and neurological effects8 may make it difficult to understand muscle-specific responses to chronic exercise. For example, thyroid hormone9,10 and insulin-like growth factor (IGF)-111 have been identified to mediate training-induced&#160;muscle adaptations, which may also regulate other signaling pathways in skeletal muscle. Notably, CCA-induced effects are minimally regulated by systemic factors, allowing focus to be placed on the direct response of skeletal muscle to contractile activity.
The external unit for CCA was first introduced by Tyler and Wright12, and has been developed with modifications12. In short, the unit is composed of three main parts: an infrared detector that can be turned on and off by exposure to infrared light, a pulse generator, and a pulse indicator (Figure 1). The detailed circuit design of the stimulator unit has been described previously13. The detailed and specific features of CCA can be found in greater depth in a number of review articles14,15,16,17. In brief, the stimulation protocol is designed to activate the common peroneal nerve at low frequency (i.e., 10 Hz), and the innervated muscles (tibialis anterior [TA] and extensor digitorum longus [EDL] muscle) are forced to contract for a predetermined length of time (e.g., 3 - 6 h). Over time, this shifts the aforementioned muscles to a more aerobic phenotype, demonstrated by an increase in both capillary density18 and mitochondrial content19,20,21. Thus, this method is a proven model to mimic some of the major endurance training adaptations within skeletal muscle of rats.
This paper presents a detailed procedure of the electrode implantation surgery to induce CCA so that the researchers can apply this model in their exercise studies. CCA is an excellent model for studying the time course of muscle adaptations, thus providing an effective tool for the investigation of various molecular and signaling events at both early and later time points following the onset of exercise training.

<table>
	<tbody>
		<tr>
			<td>Sprague Dawley Rat</td>
			<td>Charles River</td>
			<td>Strain 400</td>
			<td></td>
		</tr>
		<tr>
			<td>Chronic contractile activity unit</td>
			<td>Home-made</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>CCA unit protective box (3.5 x 3.5 x 2.5 cm)</td>
			<td>Home-made</td>
			<td>n/a</td>
			<td>Box should be made of opaque material or covered in an opague tape</td>
		</tr>
		<tr>
			<td>Coin lithium ion batteries (3V)</td>
			<td>Panasonic</td>
			<td>CR2016</td>
			<td></td>
		</tr>
		<tr>
			<td>Medwire</td>
			<td>Leico Industries</td>
			<td>316SS7/44T</td>
			<td></td>
		</tr>
		<tr>
			<td>Solder pin (socket)</td>
			<td>Digi-Key</td>
			<td>ED6218-ND</td>
			<td></td>
		</tr>
		<tr>
			<td>Zonas porous tape</td>
			<td>Johnson &#38; Johnson</td>
			<td>5104</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture silk (Size 5)</td>
			<td>Ethicon</td>
			<td>640G</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture silk (Size 6)</td>
			<td>Ethicon</td>
			<td>706G</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved blunt scissor (11.5 cm Length)</td>
			<td>F.S.T.</td>
			<td>14075-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved blunt scissor (15 cm Length)</td>
			<td>F.S.T.</td>
			<td>14111-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Delicate haemostatic forceps (16 cm Length)</td>
			<td>Lawton</td>
			<td>06-0230</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Feather</td>
			<td>3</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved forceps</td>
			<td>F.S.T.</td>
			<td>11052-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless-steel rod (30 cm; 7mm diameter)</td>
			<td>Home-made</td>
			<td>n/a</td>
			<td>Rod should have 5 mm slit in one end to hold the wire for tunneling under the skin</td>
		</tr>
		<tr>
			<td>Clip applying forceps</td>
			<td>KLS Martin</td>
			<td>20-916-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Staples (clips)</td>
			<td>Bbraun</td>
			<td>BN507R</td>
			<td></td>
		</tr>
		<tr>
			<td>Metal hooks/retractor</td>
			<td>Home-made</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone-iodine (500 mL)</td>
			<td>Rougier</td>
			<td>#NPN00172944</td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin sodium</td>
			<td>Novopharm</td>
			<td>#DIN00872644</td>
			<td></td>
		</tr>
		<tr>
			<td>Metacam</td>
			<td>Boehringer</td>
			<td>#DIN02240463</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital multimeter (voltmeter)</td>
			<td>Soar Corporation</td>
			<td>ME-501</td>
			<td></td>
		</tr>
		<tr>
			<td>LED digital stroboscope</td>
			<td>Lutron Electronic Enterprise</td>
			<td>DT-2269</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

We have shown that chronic contractile activity (CCA) is an effective tool to induce favorable mitochondrial adaptations within skeletal muscle. Rats subjected to 7 days of CCA (6 h per day) display enhanced mitochondrial biogenesis in the stimulated muscle as compared to the unstimulated contralateral (control) hindlimb. This increase in mitochondrial biogenesis is indicated by increased protein expression of PGC-1&#945; (Figure 3A), considered the master regulator of mitochondrial biogenesis, along with elevations of other key mitochondrial proteins COX-I and COX-IV, which are important components of the electron transport chain. Moreover, Cytochrome c Oxidase (COX) enzyme activity, a useful indicator of mitochondrial content, increased approximately 30% following 7 days of CCA (Figure 3B). In addition, we assessed changes in mitochondrial function using permeabilized muscle fibers to measure mitochondrial respiratory capacity and found that CCA resulted in an increase in the maximal respiratory (i.e., state 3) capacity of muscle subjected to 7 days of stimulation relative to control muscle (Figure 3C). Furthermore, both mitochondrial populations, subsarcolemmal (SS) and intermyofibrillar (IMF), increased following 7 days of CCA (Figure 4A and B), and we found the IMF fractions from muscle subjected to 7 days of contractile activity to be observably more red in color than those derived from the control (CON) muscle using differential centrifugation, presumably indicating higher levels of myoglobin, heme, and other oxygen-related elements (Figure 4C).
Adaptations to the autophagy and lysosomal systems can also be brought about by CCA. In particular, we have observed an increase in the protein abundance of transcription factor EB (TFEB; Figure 5A), the principal regulator of lysosomal biogenesis, by CCA at all time points (i.e., 1, 3, and 7 days), as well as other lysosomal markers such as lysosomal-associated membrane protein 1 (LAMP1; Figure 5B). Interestingly, our study has shown that alterations to the autophagy, mitophagy and lysosomal systems occur prior to mitochondrial biogenesis.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56827/56827fig1.jpg" /> 
Figure 1. A schematic diagram shows major components of a portable CCA stimulator unit. This Figure has been modified from Takahashi et al.13. IR, infrared. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56827/56827fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56827/56827fig2.jpg" /> 
Figure 2. Electrode and stimulator implantation. (A) A window for connecting wires at both sides of common peroneal nerve. (B) An exemplary image for demonstrating an assembly of CCA unit. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56827/56827fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56827/56827fig3.jpg" /> 
Figure 3. PGC-1alpha, mitochondrial respiration and enzyme activity with CCA. (A) CCA induces mitochondrial adaptations in skeletal muscle, as shown by increases in protein levels of PGC-1&#945;. A representative blot image is shown along with &#223;-Actin as the loading control. CON refers to Control, i.e., contralateral hindlimb not subjected to CCA, and fold change data was obtained by normalizing results relative to CON at Day 1. (B) COX activity and (C) permeabilized muscle respirations were also increased following 7 days of CCA. All data is shown as Mean &#177; SEM (N = 6-8). &#8224;P &#60;0.05, interaction effect between CCA and time; &#167;P &#60;0.05, main effect of CCA; *P &#60;0.05, significant difference vs. control at Day 1. Figure 3A and 3B have been modified from Kim and Hood19. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56827/56827fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56827/56827fig4.jpg" /> 
Figure 4. Evidence of mitochondrial biogenesis in muscle with CCA. (A and B) Electron microscopic images indicate enlarged volumes of both subsarcolemmal (SS) and intermyofibrillar (IMF) mitochondria in skeletal muscle of rats exposed to CCA for 7 days. This image has been modified from Ljubicic et al.21, and scale bars indicate 1 &#181;m. (C) A comparison of muscle mitochondrial fractionations between control (CON) and stimulated (CCA) muscle after 7 days of CCA stimulation. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56827/56827fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56827/56827fig5.jpg" /> 
Figure 5. The lysosomal system is upregulated by 7 days of CCA. This is indicated by increase in protein abundances in both (A) TFEB and (B) LAMP1. Results are shown as fold changes by normalizing data relative to CON at Day 1. (C) Representative blot images are shown and GAPDH is a loading control. &#167;P &#60; 0.05, main effect of CCA. All data is shown as Mean &#177; SEM (N = 8); &#182;P &#60; 0.05, main effect of time; *P &#60; 0.05, significant difference vs. control (CON) at Day 1.&#160;This figure has been modified from Kim and Hood19.&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56827/56827fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Application of Chronic Stimulation to Study Contractile Activity-induced Rat Skeletal Muscle Phenotypic Adaptations at JoVE.com

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

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 ...

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Nanyang Technological University

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6.7K Views.  York University. This protocol describes the use of the chronic contractile activity model of exercise to observe stimulation-induced skeletal muscle adaptations in the rat hindlimb. 

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

Ex Vivo Culture of Pharyngeal Arches to Study Heart and Muscle Progenitors and Their Niche

The pharyngeal mesoderm of developing embryos contributes to broad regions of head and heart musculature. We have developed a novel method to study head and heart progenitor cell development with pharyngeal arches (also known as branchial arches) ex vivo. Using this method, we have recently described that the second pharyngeal arch contains self-renewing heart progenitors and serves as a microenvironment for expansion of the progenitors during mouse heart development. The progenitor cells remain undifferentiated and expansive inside the arch, but quickly become functional cardiomyocytes as they migrate out of the arch. We also reported that first pharyngeal arch contains muscle progenitors giving rise to myotubes after leaving the arch. Here, we demonstrate the procedure for the dissection and ex vivo culture of first and second pharyngeal arches from developing mouse embryos. The method enables one to study head and heart progenitor/muscle development, including cardiomyocyte and myotube formation in detail ex vivo.

Ex Vivo Culture of Pharyngeal Arches to Study Heart and Muscle Progenitors and Their Niche

Here, we present a protocol to culture pharyngeal arches to study the biology of heart and muscle progenitor cells and their microenvironment.

The pharyngeal mesoderm of developing embryos contributes to broad regions of head and heart musculature. We have developed a novel method to study head ...

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 ...

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 ...

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 Vitro Generation of Somite Derivatives from Human Induced Pluripotent Stem Cells

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give rise to multiple cell types, including the myotome (MYO), sclerotome (SCL), dermatome (D), and syndetome (SYN), which in turn develop into skeletal muscle, axial skeleton, dorsal dermis, and axial tendon/ligament, respectively. Therefore, the generation of SMs and their derivatives from human induced pluripotent stem cells (iPSCs) is critical to obtain pluripotent stem cells (PSCs) for application in regenerative medicine and for disease research in the field of orthopedic surgery. Although the induction protocols for MYO and SCL from PSCs have been previously reported by several researchers, no study has yet demonstrated the induction of SYN and D from iPSCs. Therefore, efficient induction of fully competent SMs remains a major challenge. Here, we recapitulate human SM patterning with human iPSCs in vitro by mimicking the signaling environment during chick/mouse SM development, and report on methods of systematic induction of SM derivatives (MYO, SCL, D, and SYN) from human iPSCs under chemically defined conditions through the presomitic mesoderm (PSM) and SM states. Knowledge regarding chick/mouse&#160;SM development was successfully applied to the induction of SMs with human iPSCs. This method could be a novel tool for studying human somitogenesis and patterning without the use of embryos and for cell-based therapy and disease modeling.

We present here a protocol for the differentiation of human induced pluripotent stem cells into each somite derivative (myotome, sclerotome, dermatome, and syndetome) in chemically defined conditions, which has applications in future disease modeling and cell-based therapies in orthopedic surgery.

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give ...

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 ...

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

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

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

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

Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice

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

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

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