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 &amp;amp; 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/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/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/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/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/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 ...

application-chronic-stimulation-to-study-contractile-activity-induced

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

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

An in vivo Rodent Model of Contraction-induced Injury and Non-invasive Monitoring of Recovery

Muscle strains are one of the most common complaints treated by physicians. A muscle injury is typically diagnosed from the patient history and physical exam alone, however the clinical presentation can vary greatly depending on the extent of injury, the patient's pain tolerance, etc. In patients with muscle injury or muscle disease, assessment of muscle damage is typically limited to clinical signs, such as tenderness, strength, range of motion, and more recently, imaging studies. Biological markers, such as serum creatine kinase levels, are typically elevated with muscle injury, but their levels do not always correlate with the loss of force production. This is even true of histological findings from animals, which provide a "direct measure" of damage, but do not account for all the loss of function. Some have argued that the most comprehensive measure of the overall health of the muscle in contractile force. Because muscle injury is a random event that occurs under a variety of biomechanical conditions, it is difficult to study. Here, we describe an in vivo animal model to measure torque and to produce a reliable muscle injury. We also describe our model for measurement of force from an isolated muscle in situ. Furthermore, we describe our small animal MRI procedure.

An in vivo Rodent Model of Contraction-induced Injury and Non-invasive Monitoring of Recovery

An in vivo animal model of injury is described. The method takes advantage of the subcutaneous position of the fibular nerve. Velocity, timing of muscle activation, and arc of motion are all pre-determined and synchronized using commercial software. Post injury changes are monitored in vivo using MR imaging/spectroscopy.

Muscle strains are one of the most  common complaints treated by physicians. A muscle injury is typically diagnosed from the patient history and  physical ...

Medicine

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 ...

Utilization of Microscale Silicon Cantilevers to Assess Cellular Contractile Function In Vitro

The development of more predictive and biologically relevant in vitro assays is predicated on the advancement of versatile cell culture systems which facilitate the functional assessment of the seeded cells. To that end, microscale cantilever technology offers a platform with which to measure the contractile functionality of a range of cell types, including skeletal, cardiac, and smooth muscle cells, through assessment of contraction induced substrate bending. Application of multiplexed cantilever arrays provides the means to develop moderate to high-throughput protocols for assessing drug efficacy and toxicity, disease phenotype and progression, as well as neuromuscular and other cell-cell interactions. This manuscript provides the details for fabricating reliable cantilever arrays for this purpose, and the methods required to successfully culture cells on these surfaces. Further description is provided on the steps necessary to perform functional analysis of contractile cell types maintained on such arrays using a novel laser and photo-detector system. The representative data provided highlights the precision and reproducible nature of the analysis of contractile function possible using this system, as well as the wide range of studies to which such technology can be applied. Successful widespread adoption of this system could provide investigators with the means to perform rapid, low cost functional studies in vitro, leading to more accurate predictions of tissue performance, disease development and response to novel therapeutic treatment.

Utilization of Microscale Silicon Cantilevers to Assess Cellular Contractile Function In Vitro

This protocol describes the use of microscale silicon cantilevers as pliable culture surfaces for measuring the contractility of muscle cells in vitro. Cellular contraction causes cantilever bending, which can be measured, recorded, and converted into readouts of force, providing a non-invasive and scalable system for measuring contractile function in vitro.

The development of more predictive and biologically relevant in vitro assays is predicated on the advancement of versatile cell culture systems which ...

Bioengineering

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 ...

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

Procedures for Rat in situ Skeletal Muscle Contractile Properties

There are many circumstances where it is desirable to obtain the contractile response of skeletal muscle under physiological circumstances: normal circulation, intact whole muscle, at body temperature. This includes the study of contractile responses like posttetanic potentiation, staircase and fatigue. Furthermore, the consequences of disease, disuse, injury, training and drug treatment can be of interest. This video demonstrates appropriate procedures to set up and use this valuable muscle preparation.
To set up this preparation, the animal must be anesthetized, and the medial gastrocnemius muscle is surgically isolated, with the origin intact. Care must be taken to maintain the blood and nerve supplies. A long section of the sciatic nerve is cleared of connective tissue, and severed proximally. All branches of the distal stump that do not innervate the medial gastrocnemius muscle are severed. The distal nerve stump is inserted into a cuff lined with stainless steel stimulating wires. The calcaneus is severed, leaving a small piece of bone still attached to the Achilles tendon. Sonometric crystals and/or electrodes for electromyography can be inserted. Immobilization by metal probes in the femur and tibia prevents movement of the muscle origin. The Achilles tendon is attached to the force transducer and the loosened skin is pulled up at the sides to form a container that is filled with warmed paraffin oil. The oil distributes heat evenly and minimizes evaporative heat loss. A heat lamp is directed on the muscle, and the muscle and rat are allowed to warm up to 37&deg;C. While it is warming, maximal voltage and optimal length can be determined. These are important initial conditions for any experiment on intact whole muscle. The experiment may include determination of standard contractile properties, like the force-frequency relationship, force-length relationship, and force-velocity relationship.
With care in surgical isolation, immobilization of the origin of the muscle and alignment of the muscle-tendon unit with the force transducer, and proper data analysis, high quality measurements can be obtained with this muscle preparation.

Procedures for Rat in situ Skeletal Muscle Contractile Properties

This video demonstrates the surgical preparation and procedures needed to study the contractile responses of the rat medial gastrocnemius muscle preparation in situ. This preparation allows measurement of skeletal muscle contractile properties under physiological conditions. The animal is anesthetized and the muscle is separated from surrounding tissue at its distal end. The Achilles tendon is attached to a force transducer, allowing measurement of the muscle&#x2019;s contractile response at 37 degrees C with an intact circulation.

There are many circumstances where it is desirable to obtain the contractile response of skeletal muscle under physiological circumstances: normal ...

Biology

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+ handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+ signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e., mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

We describe a method to directly measure muscle force, muscle power, contractile kinetics and fatigability of isolated skeletal muscles in an in vitro system using field stimulation. Valuable information on Ca2+ handling properties and contractile machinery of the muscle can be obtained using different stimulating protocols.

 
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, ...

In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological characteristics in the intact spinal network and holds significant advantages relative to classical in vitro or extracellular recording techniques. An advantage of in vivo intracellular recordings is that this method can be performed on adult animals with a fully mature nervous system, and therefore many observed physiological mechanisms can be translated to practical applications. In this methodological paper, we describe this procedure combined with externally applied constant current stimulation, which mimics polarization processes occurring within spinal neuronal networks. Trans-spinal direct current stimulation (tsDCS) is an innovative method increasingly used as a neuromodulatory intervention in rehabilitation after various neurological injuries as well as in sports. The influence of tsDCS on the nervous system remains poorly understood and the physiological mechanisms behind its actions are largely unknown. The application of the tsDCS simultaneously with intracellular recordings enables us to directly observe changes of motoneuron membrane properties and characteristics of rhythmic firing in response to the polarization of the spinal neuronal network, which is crucial for the understanding of tsDCS actions. Moreover, when the presented protocol includes the identification of the motoneuron with respect to an innervated muscle and its function (flexor versus extensor) as well as the physiological type (fast versus slow) it provides an opportunity to selectively investigate the influence of tsDCS on identified components of spinal circuitry, which seem to be differently affected by polarization. The presented procedure focuses on surgical preparation for intracellular recordings and stimulation with an emphasis on the steps which are necessary to achieve preparation stability and reproducibility of results. The details of the methodology of the anodal or cathodal tsDCS application are discussed while paying attention to practical and safety issues.

This protocol describes in vivo intracellular recording of rat lumbar motoneurons with simultaneous trans-spinal direct current stimulation. The method enables us to measure membrane properties and to record rhythmic firing of motoneurons before, during and after anodal or cathodal polarization of the spinal cord.

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological ...

Neuroscience

Contractility Measurements on Isolated Papillary Muscles for the Investigation of Cardiac Inotropy in Mice

Papillary muscle isolated from adult mouse hearts can be used to study cardiac contractility during different physiological/pathological conditions. The contractile characteristics can be evaluated independently of external influences such as vascular tonus or neurohumoral status. It depicts a scientific approach between single cell measurements with isolated cardiac myocytes and in vivo studies like echocardiography. Thus, papillary muscle preparations serve as an excellent model to study cardiac physiology/pathophysiology and can be used for investigations like the modulation by pharmacological agents or the exploration of transgenic animal models. Here, we describe a method of isolating the murine left anterior papillary muscle to investigate cardiac contractility in an organ bath setup. In contrast to a muscle strip preparation isolated from the ventricular wall, the papillary muscle can be prepared in toto without damaging the muscle tissue severely. The organ bath setup consists of several temperature-controlled, gassed and electrode-equipped organ bath chambers. The isolated papillary muscle is fixed in the organ bath chamber and electrically stimulated. The evoked twitch force is recorded using a pressure transducer and parameters such as twitch force amplitude and twitch kinetics are analyzed. Different experimental protocols can be performed to investigate the calcium- and frequency-dependent contractility as well as dose-response curves of contractile agents such as catecholamines or other pharmaceuticals. Additionally, pathologic conditions like acute ischemia can be simulated.

Murine left ventricular papillary muscle can be used to investigate cardiac contractility in vitro. This article describes in detail the isolation and experimental protocols to study cardiac contractile characteristics.

Papillary muscle isolated from adult mouse hearts can be used to study cardiac contractility during different physiological/pathological conditions. The ...

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury and/or remodeling. One approach for assessing these functional alterations utilizes unloaded shortening and Ca2+ transient analyses in primary adult cardiac myocytes. For this approach, adult myocytes are isolated by collagenase digestion, made Ca2+ tolerant, and then adhered to laminin-coated coverslips, followed by electrical pacing in serum-free media. The general protocol utilizes adult rat cardiac myocytes but can be readily adjusted for primary myocytes from other species. Functional alterations in myocytes from injured hearts can be compared to sham myocytes and/or to in vitro therapeutic treatments. The methodology includes the essential elements needed for myocyte pacing, along with the cell chamber and platform components. The detailed protocol for this approach incorporates the steps for measuring unloaded shortening by sarcomere length detection and cellular Ca2+ transients measured with the ratiometric indicator Fura-2 AM, as well as for raw data analysis.

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

A set of protocols are presented that describe the measurement of contractile function via sarcomere length detection along with calcium (Ca2+) transient measurement in isolated rat myocytes. The application of this approach for studies in animal models of heart failure is also included.

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury ...

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

Summary

Explore More Videos

Chapters in this video

An in vivo Rodent Model of Contraction-induced Injury and Non-invasive Monitoring of Recovery

A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation

Utilization of Microscale Silicon Cantilevers to Assess Cellular Contractile Function In Vitro

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

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

Procedures for Rat in situ Skeletal Muscle Contractile Properties

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

Contractility Measurements on Isolated Papillary Muscles for the Investigation of Cardiac Inotropy in Mice

Analysis of Cardiac Contractile Dysfunction and Ca²⁺ Transients in Rodent Myocytes