We would like to thank Rosario Maroto for technical assistance. Research reported in this publication was supported by the National Institute of Arthritis and Musculoskeletal and Skin Diseases of the National Institutes of Health under Award Number R01 AR072061 (CSF). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

All experimental procedures were approved by University of Kentucky Institutional Animal Care and Use Committee.
1. Equipment setup
<ol>
	<li>Confirm that machines are connected per manufacturer specifications.</li>
	<li>If not already in place, attach the 300D-305C-FP motor with knee extension apparatus to the 809C animal platform.</li>
	<li>Turn on the water pump to 37 &#176;C to begin heating the platform.</li>
	<li>If computer is not already on, turn the computer on, followed by the High-Power Bi-Phase Stimulator and 2 Channel Dual-Mode Lever System.</li>
	<li>Pour isoflurane into vaporizer to maximum fill line.</li>
</ol>
2. Software setup
<ol>
	<li>Open the software (details provided in Table of Materials).</li>
	<li>To use the Instant Stimulation feature in conjunction with Live Data Monitor to optimize probe placement (step 4), select Prepare Experiment followed by Configure Instant Stim (Figure 1). Set the Pulse Frequency (Hz) as 125, Pulse Width (ms) as 0.2, Number of Pulses as 1, Train Frequency (Hz) as 0.5 and Run Time (s) as 120.</li>
	<li>Select File and open Live Data Monitor.</li>
	<li>To perform twitch (step 5) and torque-frequency (Step 6) experiments, select a previously programmed study that includes appropriate twitch and knee extension torque-frequency experiments (detailed below in step 5 and step 6).
	<ol>
		<li>Select the appropriate experimental mouse or Add New Animal and input corresponding mouse information to be stored with torque data.</li>
		<li>Select Next Experiment or Previous Experiment to transition from twitch protocol to torque-frequency sequence.</li>
	</ol>
	</li>
</ol>
3. Mouse setup
<ol>
	<li>Place individual mouse into the anesthetic chamber.</li>
	<li>Release oxygen tank valve and set oxygen flow rate at 1 L/min with 2.5% isoflurane.</li>
	<li>Ensure that mouse remains in the chamber with the lid securely closed until completely unconscious. Confirm complete loss of consciousness by absent foot reflex with toe pinch.</li>
	<li>Place anesthetized mouse in a supine position with the head in the nosecone on the heated platform with oxygen flow rate at 1 L/min with 2.5% isoflurane.</li>
	<li>Shave hair from the right hind limb using electric clippers. Remove hair from the shaved area with an alcohol wipe and a small vacuum. Clean removed hair away from the hind limb and the platform.&#160;</li>
	<li>Securely clamp the upper hind limb, posterior to the knee (Figure 2). 
	NOTE: Ensure that knee range of motion is not impeded.</li>
	<li>Place the lower hind limb into the knee extension apparatus with the anterior tibia lightly touching the adjustable plastic piece (The Force In channel reading should read between 0 and -1.0 mN*m). Depending on the size of the lower hind limb of the mouse, surgical tape may be wrapped around the bottom portion of the adjustable plastic piece to allow the leg to rest securely. 
	NOTE: Detailed images and dimensions of the custom-fabricated plastic piece are shown in Supplementary Figure 1.</li>
	<li>Adjust knobs on platform to ensure the knee is bent at 60&#176;.</li>
	<li>Lightly place a piece of tape over the mouse torso onto the platform to prevent compensatory movement with maximal knee extension.</li>
</ol>
4. Electrode placement
<ol>
	<li>Place electrodes subcutaneously 2-4 mm proximal to the knee directly above the quadriceps/knee extensor muscles (Figure 2). Electrodes should be approximately 1-2 mm apart.</li>
	<li>To determine optimal placement of electrodes, utilize the Instant Stimulation function with Live Data Monitor. Set amperage/current at 50 mA for repeated twitches to confirm knee extension (the knee extensors will produce a negative twitch curve). Adjust the probes during Instant Stimulation to achieve maximal knee extension twitch torque as measured in the Live Data Monitor window. 
	NOTE: Figure 3 shows a representative Instant Stimulation output, confirming knee extension. Supplementary Video 1 and Supplementary Video 2 show real-time and slow-motion knee extensor twitches without the motor arm in place, allowing for visual confirmation of knee extension.</li>
	<li>During repeated twitches with Instant Stimulation, palpate knee flexor muscles with the index finger to confirm no activation of antagonist muscles. To maximally stimulate knee extensors, probe repositioning may be necessary depending on body composition of the mouse and slight anatomical differences in the exact location of the motor point of the femoral nerve and knee extensor muscles. 
	NOTE: A muscle motor point is the location where the motor branch of a nerve enters the muscle belly and is the point with the least resistance to electrical conductivity and subsequently the highest responsiveness to electrical stimulation14,15. In clinical applications using electrical stimulation, this point is identified by scanning with a pen electrode to find the location above the muscle at which a muscle twitch occurs with the lowest injected current14,15. Identification of the muscle motor point is essential to facilitate optimal neuromuscular electrical stimulation15. In human clinical trials, muscle motor points for quadriceps muscles have been identified in the distal half of the muscle14. To achieve optimal knee extensor stimulation in mice, this technique was recapitulated using electrode placement with Instant Stimulation to most closely approximate muscle motor point locations typically found in the distal half of knee extensors. There exists some variability in electrode placement (from relatively superficial to deep) that results in maximal torque, and the Instant Stimulation function facilitates optimal electrode placement.</li>
</ol>
5. Determination of optimal current
<ol>
	<li>Once optimal probe placement is determined, perform a series of progressive twitches to determine optimal amperage/current to be used for the torque-frequency experiment, with the goal of determining the lowest current to achieve the maximal twitch torque output. Begin with current set at 50 mA and select Run Experiment to produce a single twitch. Select Analyze Results to display torque output. Record the twitch torque displayed under Max Force with baseline subtracted. 
	NOTE: Select the option to invert the Force channel to convert measurements from negative torque to positive.</li>
	<li>Increase the current to 60-70 mA and repeat twitch experiment. Record the twitch torque displayed under Max Force with baseline subtracted.</li>
	<li>Continue with a series of twitch experiments in this manner (increasing approximately 10-20 mA with each progression) until twitch torque no longer increases (either plateaus or begins to decrease). Example of twitch series is shown in Table 1.</li>
	<li>Record the lowest current at which the highest twitch torque was achieved. This current will be used and remain constant during the upcoming force-frequency experiment. Figure 4 shows a representative peak twitch.</li>
</ol>
6. Torque-frequency experiment to determine peak isometric tetanic torque
<ol>
	<li>In the software (see Table of Materials), select the pre-programmed torque-frequency experiment for knee extension ensuring following setting. Stimulus duration: 0.35 s, Frequency sequence: 10 Hz, 40 Hz, 120 Hz, 150 Hz, 180 Hz, 200 Hz, Rest period between pulses/contractions: 120 s 
	NOTE: Sampling rate is 10,000 Hz (default setting).</li>
	<li>Run Experiment, Analyze Results, and manually record the torque displayed under Max Force with baseline subtracted (make sure the force channel is inverted, as knee extensor contraction will produce negative torque) at each frequency. Note the highest Max Force value as the peak isometric tetanic torque. Example of torque-frequency data is shown in Table 2 and Figure 5 shows a representative tetanus curve for the peak isometric tetanic torque output achieved at 120 Hz.</li>
</ol>
7. Termination of experiment
<ol>
	<li>Upon completion of the torque-frequency experiment, perform a follow up twitch and compare with the initial peak twitch at the same current to assess damage/fatigue. 
	NOTE: In some models of injury and disease, increased fatigability of skeletal muscle is expected and does not constitute a problem with the experimental setup or the mouse.</li>
	<li>When all torque measurements are completed, gently remove electrode probes, and unclamp the knee.</li>
	<li>Turn off isoflurane and remove the mouse from the nose cone.</li>
	<li>Place the mouse back into an appropriate cage placed on top of a warming pad. Monitor as the mouse recovers and regains consciousness. 
	&#8203;NOTE: The mouse should be conscious and moving within 2-3 min.</li>
</ol>
8. Data analysis
<ol>
	<li>Extract data after the experiment from analysis software (see Table of Materials).
	<ol>
		<li>Open analysis software.</li>
		<li>Select Get Data from the software.</li>
		<li>Select Date on which experiment was performed and appropriate mouse code.</li>
		<li>Select the frequency of interest (all twitch experiments and each frequency of the torque-frequency experiment will be listed).</li>
		<li>Select Muscle Analysis.</li>
		<li>Confirm that Use Baseline Correction is checked. 
		NOTE: Baseline torque is calculated by the software as the average of the first 100 points sampled and subtracted from the absolute maximum torque value.</li>
		<li>Record the torque value listed under Maximum. 
		NOTE: Data presented here are unfiltered; however, a filter may be selected in the software, if desired.</li>
	</ol>
	</li>
	<li>Alternatively, as described above in step 6.2, manually record the torque output displayed under Max Force in real time at each torque-frequency point/contraction through the Analyze Results window.
	<ol>
		<li>Confirm that the baseline is subtracted, and force channel is inverted.</li>
		<li>Input data into a spreadsheet for body weight normalization calculations (torque/body weight in grams) and graphing and statistical analyses of interest. Statistical software was used for the purpose of graphing torque-frequency curves and calculating area under the curve. 
		NOTE: Torque data are measured in mN.m (milliNewton.meters).</li>
	</ol>
	</li>
	<li>To generate tetanus curves, export complete data from each frequency from the analysis software.
	<ol>
		<li>Repeat steps 8.1.1-8.1.4 above.</li>
		<li>Select Export Data.</li>
		<li>Select Raw Filtered Data and save in location of choice. MATLAB may be used to generate tetanus curves from the exported text file and/or for further analysis. 
		&#8203;NOTE: MATLAB code to generate tetanus curve from text file is available upon request.</li>
	</ol>
	</li>
</ol>
9. Dual-mode lever system calibration
<ol>
	<li>Calibrate the system prior to the initial use to ensure accurate and reliable data, and repeat calibration periodically using the data collection software and known weights.
	<ol>
		<li>Open data collection software.</li>
		<li>Click the Setup tab and select Channel Setup.</li>
		<li>Select 305C-FP listed under My Instruments.</li>
		<li>Click Calibrate Selected to open the Calibration Editor window.</li>
		<li>To calibrate length, input a series of test voltages including both negative and positive voltages (e.g., -3, -2, -1, 0, 1, 2, 3 V).
		<ol>
			<li>Click Set for the first line.</li>
			<li>Click Read.</li>
			<li>Measure the exact length of the lever arm in millimeters and input into the corresponding box.</li>
			<li>Repeat for the next voltage.</li>
			<li>Upon recording of all voltages, click Calculate Cal Factors (recorded in mm/volt).</li>
		</ol>
		</li>
		<li>To calibrate force in, utilize a set of known weights increasing in a linear progression.
		<ol>
			<li>Adjust the motor so that it is resting on the edge of the bench or table with the lever arm parallel to the benchtop and hanging over the edge to allow the weight to hang.</li>
			<li>Hang the first weight from the lever arm using a rubber band. Under Applied Force, enter the known weight in grams accounting for the mass of the rubber band.</li>
			<li>Select Read.</li>
			<li>Repeat for at least 3 known weights.</li>
			<li>Select Calculate Cal Factor.</li>
			<li>To verify calculation, plot calibration data and curve fit by selecting Plot Cal.</li>
		</ol>
		</li>
		<li>To calibrate force out, enter calibration voltages (up to 10 volts)
		<ol>
			<li>Click Set directly next to the calibration voltage.</li>
			<li>Repeat for each voltage line.</li>
			<li>Gently apply pressure to the lever arm using a finger until the Force Out ceases to change and the motor arm begins to move.</li>
			<li>Maintain this position. Select Read.</li>
			<li>Repeat for each voltage line.</li>
			<li>Select Calculate Cal Factor.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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</ol>

Matthew Borkowski is employed by Aurora Scientific Inc., a company that may potentially benefit from the research results and is also an executive of the company.

Measurement and analysis of muscle function in rodent models is imperative to make translational and meaningful inferences regarding histological and molecular skeletal muscle adaptations observed with exercise, injury, disease, and therapeutic treatment. We demonstrate a method to assess knee extensor maximal strength reliably and repeatedly in mice using commercially available equipment, with the adjustable plastic piece to hold the lower hind limb at the anterior tibia being the only custom fabricated part that may be replicated.
Common functional assessment tools have been widely used to repeatedly evaluate physical performance within the same mouse, such as treadmill running to volitional fatigue, rotarod performance test, inverted cling test, and grip strength test. However, while informative, these assessments involve cardiopulmonary and behavioral component(s), which can obfuscate the interrogation of neuromuscular function associated with these physical performance measures. Furthermore, elements of endurance, coordination, and balance are present in many of these functional assessments to varying levels, limiting clear interpretation relative to muscle strength. The force producing ability of rodent muscle(s) can be measured in vitro, in situ, or in vivo. Each approach has relative advantages and limitations. Specifically, with in vitro assessment, the muscle is completely isolated and removed from the body of the animal so that there is no influence from perfusion or innervation19. This yields a well-controlled environment to ascertain contractile ability but limits the size of the muscle being studied through dependency on passive diffusion of oxygen and nutrients during testing. In situ testing maintains the innervation and blood supply of the muscle, but is limited to a singular terminal assessment, as with in vitro testing20. Finally, in vivo testing is the least invasive with the muscle remaining in its native environment with percutaneous electrodes inserted near the motor nerve to electrically stimulate the muscle. A strength of the in vivo approach is the potential for longitudinal testing across time21,22,23.
In vivo&#160;evaluation of peak muscle contractility optimally measures maximal strength as the normal anatomy and physiology of the mouse remains intact and the method may be repeated on the same mouse before and after an intervention or throughout the lifespan. Specifically, in vivo measurement of knee extensor strength in mice is the murine strength assessment with the greatest translational relevance to human studies, as maximum knee extension torque is commonly measured and considered the gold-standard strength test in humans with correlation to various functional and health outcomes24,25,26,27. Moreover, knee extensor pathology is observed with aging as well as a myriad of injuries and diseases1,2,4,5,6, but assessing the impact of these conditions on knee extensor strength longitudinally in mice has not been readily attainable.
Although this method offers utility to determine knee extensor peak torque in a longitudinal manner, certain limitations of the protocol should be considered. Lower frequencies between 40 Hz to 120 Hz were omitted from the torque-frequency protocol, which may limit the ability to detect left or rightward shifts in the torque-frequency curve with injury or disease. However, using this torque-frequency protocol, we have been able to detect alterations to peak tetanic torque in an ACL injury model and between C56BL/6 wild type mice and a transgenic mouse model of supraphysiological muscle mass (Figure 8). We note that it may be beneficial to secure the electrodes with helping hands or similar apparatus as muscle contractions may move electrodes slightly. We did not note any obvious displacement of electrodes with progressive contractions; however, the possibility of slight movement of the electrodes cannot be ruled out, which may impact muscle stimulation. Additionally, intramuscular electromyography (EMG) was not performed in conjunction with the stimulus protocol; however, inclusion of EMG measures may be feasible, if desired and appropriate for the experimental model of interest.
Assessment of knee extensor strength in murine models of orthopedic injury and disease will facilitate pre-clinical research with meaningful translational relevance to clinical strength measures. Our protocol enables precise and repeated assessment of maximal knee extensor strength in mice with commercially available equipment accessible to any laboratory.

Skeletal muscle is a highly adaptable tissue with compensatory alterations to mass and structure in response to a myriad of stimuli, such as exercise, nutrition, injury, disease, aging, and disuse. Many studies investigating skeletal muscle adaption in humans employ methods to measure both skeletal muscle size and impact on function, as gold-standard strength assessments are easily repeatable in human subjects.
Specifically, knee extensor and flexor strength are&#160;most assessed in clinical research. Alterations to knee extensor strength have been widely reported in human studies of aging, exercise, orthopedic injury, knee osteoarthritis, chronic disease, and disuse1,2,3,4,5,6,7. However, methods to repeatedly and non-invasively analyze knee extensor muscle (quadriceps) strength in mechanistic rodent studies have been relatively limited. A method to determine in vivo quadriceps muscle contractility in rats was previously developed8; however, extensive construction of non-commercially available equipment is required. Given the breadth of rodent models developed to study musculoskeletal outcomes following knee injury/osteoarthritis9,10,11,12,13 there exists a need for non-invasive assessment of quadriceps strength.
Furthermore, rodent studies investigating molecular mechanisms underpinning skeletal muscle adaptation often utilize mouse models due to the simplicity of genetic modification, as do many pharmacological intervention studies because of the decreased financial expense associated with lower weight-based dosing of a drug in mice compared to rats. We report a non-invasive method to repeatedly measure in vivo knee extensor function in the same mouse over time using commercially available equipment with minor modification, facilitating reproducibility among different laboratories, and providing more direct comparison to human strength outcomes.

<table><tbody><tr><td>1300A: 3-in-1 Whole Animal System- Mouse</td><td>Aurora Scientific Incorporated</td><td>300D-305C-FP: dual-mode motor with custom knee extension apparatus, 605A: Dynamic Muscle Data Acquisition and Analysis System, 701C: Electrical Stimulator, 809C: &lt;em&gt;in-situ&lt;/em&gt; Mouse Apparatus</td><td></td></tr><tr><td>6100 Dynamic Muscle Control LabBook software</td><td>Aurora Scientific Incorporated</td><td>DMC v6.000</td><td></td></tr><tr><td>611A Dynamic Muscle Analysis</td><td>Aurora Scientific Incorporated</td><td>DMA v5.501</td><td></td></tr><tr><td>BravMini hair clippers</td><td>Wahl Clipper Corporation</td><td>ASIN: B00IN24ILE</td><td></td></tr><tr><td>Eye Lube</td><td>Optixcare</td><td>Item Number: 142422</td><td></td></tr><tr><td>Isoflurane</td><td>Covetrus</td><td>NDC: 11695-6777-2</td><td></td></tr><tr><td>V-1 Tabletop Laboratory Animal Anesthesia System</td><td>VetEquip Inhalation Anesthesia Systems</td><td>Item Number: 901806</td><td></td></tr><tr><td>Prism 8</td><td>GraphPad Software, LLC</td><td>Version 8.3.0 (328)</td><td></td></tr></tbody></table>

in vivo measurement of knee extensor muscle function in mice

Skeletal muscle plasticity in response to countless conditions and stimuli mediates concurrent functional adaptation, both negative and positive. In the clinic and the research laboratory, maximal muscular strength is widely measured longitudinally in humans, with knee extensor musculature the most reported functional outcome. Pathology of the knee extensor muscle complex is well documented in aging, orthopedic injury, disease, and disuse; knee extensor strength is closely related to functional capacity and injury risk, underscoring the importance of reliable measurement of knee extensor strength. Repeatable, in vivo assessment of knee extensor strength in pre-clinical rodent studies offers valuable functional endpoints for studies exploring osteoarthritis or knee injury. We report an in vivo and non-invasive protocol to repeatedly measure isometric peak tetanic torque of the knee extensors in mice across time. We demonstrate consistency using this novel method to measure knee extensor strength with repeated assessment in multiple mice producing similar results.

The torque-frequency curve utilizes lower frequencies to produce multiple isolated isometric twitches of relatively low torque and progresses through increasingly higher frequencies, resulting in fusion of twitches for an isometric tetanus contraction at which peak tetanic torque is obtained. The presented protocol for knee extension peak tetanic torque the force-frequency curve initiates at 10 Hz which elicits 3 isolated twitches. Partial fusion of twitches occurs at 40 Hz, and peak tetanic torque is reached between 120-180 Hz (Figure 5).
Figure 6&#160;illustrates representative knee extension torque-frequency curves from female C57BL/6 mice. Three separate mice were tested at baseline, and the experiment was repeated in each mouse 2 weeks later for comparison to assess reproducibility. Torque-frequency curves are shown with raw torque values (Figure 6A), as well as raw torque values normalized to mouse body weight (Figure 6B). Repeated observations demonstrate comparable results in all 3 mice with a 2-week rest period between experiments. Body weight normalized torque data should be considered in addition to raw torque, as minor fluctuation in weight may impact functional output and is not considered with raw torque alone. Furthermore, body weight normalized torque data facilitates comparison of mice of varying sizes. Torque can also be normalized to muscle wet weight or myofiber cross sectional area, as we have previously shown16.
Figure 7A shows the area under the curve using body weight normalized isometric torque data from complete torque-frequency experiments (10 Hz, 40 Hz, 120 Hz, 150 Hz, 180 Hz, 200 Hz) for 4 separate C57BL/6 mice, highlighting similar total torque output and coefficients of variation between 5.6% to 8.8% with repeated experiments within the same mice. Data are most simply reported as peak tetanic torque (Figure 7B) which is the maximal torque value from the repeated tetanus isometric contractions from 120-200 Hz. The peak tetanic torque output is comparable in 6-8-month-old female C57BL/6 mice (Figure 7B) with coefficients of variation between 4.8% and 8.7% with longitudinal assessment within the same mice. Peak tetanic torque is most comparable to the gold-standard strength assessment in human studies: maximum isometric toque.
Furthermore, the knee extensor peak tetanic torque protocol is a useful tool to detect strength differences in multiple mouse models. Figure 8 demonstrates the stark contrast between knee extensor strength in a non-injured, healthy 6-month-old C57BL/6 female mouse (black line) and a transgenic mouse model of supraphysiological hypertrophy in which myostatin/GDF8 is knocked out (blue line). We also show a peak tetanus curve from a C57BL/6 mouse 7 days after surgical transection of the anterior cruciate ligament (ACL-T) (red line), demonstrating a nearly 50% decline in peak torque after injury which is well outside the coefficients of variation observed with repeat testing of uninjured mice. Concurrent with human data17,18, strength is markedly diminished with ACL-T. All mice are female and of similar age (6-8 months).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Twitch experiment</td>
			<td>Amperage/Current (mA)</td>
			<td>Torque (mN&#8226;m)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>50</td>
			<td>1.279</td>
		</tr>
		<tr>
			<td>2</td>
			<td>70</td>
			<td>1.341</td>
		</tr>
		<tr>
			<td>3</td>
			<td>90</td>
			<td>1.36</td>
		</tr>
		<tr>
			<td>4</td>
			<td>110</td>
			<td>1.362</td>
		</tr>
		<tr>
			<td>5</td>
			<td>*130</td>
			<td>1.449</td>
		</tr>
		<tr>
			<td>6</td>
			<td>150</td>
			<td>1.436</td>
		</tr>
		<tr>
			<td>7</td>
			<td>140</td>
			<td>1.333</td>
		</tr>
	</tbody>
</table>
Table 1: Example of twitch series. * denotes optimal amperage/current.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Frequency (Hz)</td>
			<td>Torque(mN&#8226;m)</td>
		</tr>
		<tr>
			<td>10</td>
			<td>1.385</td>
		</tr>
		<tr>
			<td>40</td>
			<td>1.869</td>
		</tr>
		<tr>
			<td>120</td>
			<td>*18.765</td>
		</tr>
		<tr>
			<td>150</td>
			<td>18.375</td>
		</tr>
		<tr>
			<td>180</td>
			<td>17.97</td>
		</tr>
		<tr>
			<td>200</td>
			<td>17.548</td>
		</tr>
	</tbody>
</table>
Table 2: Example of torque-frequency curve data. * denotes peak tetanic torque.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62211/62211fig01.jpg" /> 
Figure 1: Data collection software setup. Illustration of setup for data collection software with Live Data Monitor. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig01large.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/62211/62211fig2v2.jpg" /> 
Figure 2: Mouse setup and electrode placement. (A-B) Supine position of the mouse receiving anesthesia via a nose cone on the heated platform. Upper hind limb is securely clamped, posterior to the knee to allow for unrestricted movement at the knee joint. Motor arm is adjusted so that knee is bent at approximately 60&#176;. The femoral nerve motor point is stimulated by needle electrodes to activate contraction of knee extensors. Mouse setup is shown from a side view (A) and overhead view (B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig2v2large.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/62211/62211fig03.jpg" /> 
Figure 3: Determination of optimal electrode placement to achieve isometric knee extension. Representation of repeated negative twitches stimulated with 50 mA using the Instant Stimulation function and viewed in the Live Data Monitor. Red arrows indicate the first three knee extension twitches. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig03large.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/62211/62211fig04.jpg" /> 
Figure 4: Representative twitch to determine optimal amperage. The lowest amperage to elicit the highest twitch isometric torque must be determined for the force-frequency experiment by repeated twitch experiments with progressively increased amperage. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig04large.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/62211/62211fig05.jpg" /> 
Figure 5: Representative tetanic torque curves throughout a torque-frequency experiment for the same mouse. (A) Submaximal isometric tetanic torque produced at 10 Hz. (B) Submaximal isometric tetanic torque at 40 Hz. (C) Peak isometric tetanic torque output at 120 Hz. (D) Isometric tetanic torque at 150 Hz. (E) Isometric tetanic torque at 180 Hz. (F) Isometric tetanic torque at 200 Hz. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62211/62211fig06.jpg" /> 
Figure 6: Representative torque-frequency curve data. (A-B). Torque-frequency curve at 2 different timepoints (week 1 and 3) in 3 separate mice, presented as raw peak torque (A) and raw peak torque normalized to body weight (B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62211/62211fig07.jpg" /> 
Figure 7: Representative area under the curve (AUC) and peak tetanic torque data. (A)&#160;AUC for 4 separate mice, presented as raw torque normalized to body weight. (B) Peak tetanic torque for the same 4 mice, presented as raw peak tetanic torque normalized to body weight. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/62211/62211fig08.jpg" /> 
Figure 8: Peak tetanic torque of knee extensors in multiple mouse models. Representative peak torque tetanus curves for an overt hypertrophy transgenic mouse model (GDF8 KO), an uninjured healthy C57BL/6 mouse (mouse 2), and a C57BL/6 mouse 7 days after anterior cruciate ligament transection (ACL-T). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/62211fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Dimensions of custom fabricated plastic. Inset in red shows dimension of depth.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/Supplementary Figure 1.pdf" target="_blank">Please click here to download this File.</a>
Supplementary Video 1: Real-time knee extensor twitch without motor arm.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/Supplementary video 1.mov" target="_blank">Please click here to download this Video.</a>
Supplementary Video 2: Slow-motion knee extensor twitch without motor arm.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62211/Supplementary video 2.mov" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about In vivo Measurement of Knee Extensor Muscle Function in Mice at JoVE.com

In vivo Measurement of Knee Extensor Muscle Function in Mice

Quantification of knee extensor maximal strength is imperative to understand functional adaptations to aging, disease, injury, and rehabilitation. We present a novel method to repeatedly measure in vivo knee extension isometric peak tetanic torque.

Skeletal muscle plasticity in response to countless conditions and stimuli mediates concurrent functional adaptation, both negative and positive. In the ...

in-vivo-measurement-of-knee-extensor-muscle-function-in-mice

Nanyang Technological University

Research

JoVE Journal

Biology

4.5K Views.  University of Kentucky. Knie-extensorsterkte is een veel voorkomende functionele uitkomst die wordt beoordeeld in klinisch onderzoek. Methoden voor het herhaaldelijk beoordelen van de knie-extensorspier, met name quadricepkracht, in knaagdierstudies zijn echter relatief beperkt geweest. Dit niet-invasieve protocol kan worden gebruikt om het isometrische piektetanische koppel van knie-extensoren bij muizen te meten en kan in de lengterichting worden herhaald.Onze methoden ondersteunen de ontwikkeling van prekl...

Video: In vivo Measurement of Knee Extensor Muscle Function in Mice

In Vivo Canine Muscle Function Assay

We describe a minimally-invasive and reproducible method to measure canine pelvic limb muscle strength and muscle response to repeated eccentric 
contractions. The pelvic limb of an anesthetized dog is immobilized in a stereotactic frame to align the tibia at a right angle to the femur. Adhesive wrap affixes the 
paw to a pedal mounted on the shaft of a servomotor to measure torque. Percutaneous nerve stimulation activates pelvic limb muscles of the paw to either push (extend) or 
pull (flex) against the pedal to generate isometric torque. Percutaneous tibial nerve stimulation activates tibiotarsal extensor muscles. Repeated eccentric (lengthening) 
contractions are induced in the tibiotarsal flexor muscles by percutaneous peroneal nerve stimulation. The eccentric protocol consists of an initial isometric contraction 
followed by a forced stretch imposed by the servomotor. The rotation effectively lengthens the muscle while it contracts, e.g., an eccentric contraction. During 
stimulation flexor muscles are subjected to an 800 msec isometric and 200 msec eccentric contraction. This procedure is repeated every 5 sec. To avoid fatigue, 4 min rest 
follows every 10 contractions with a total of 30 contractions performed.

In Vivo Canine Muscle Function Assay

We describe a minimally-invasive and painless method to measure canine hindlimb muscle strength and muscle response to repeated eccentric contractions.

We describe a minimally-invasive and reproducible method to measure canine pelvic limb muscle strength and muscle response to repeated eccentric 
...

Medicine

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no curative treatment available. Recent advances in stem cell biology have generated new hopes for the development of effective cell based therapies to treat these diseases. Transplantation of various types of stem cells labeled with fluorescent proteins into muscles of dystrophic animal models has been used broadly in the field. A non-invasive technique with the capability to track the transplanted cell fate longitudinally can further our ability to evaluate muscle engraftment by transplanted cells more accurately and efficiently.
In the present study, we demonstrate that in vivo fluorescence imaging is a sensitive and reliable method for tracking transplanted GFP (Green Fluorescent Protein)-labeled cells in mouse skeletal muscles. Despite the concern about background due to the use of an external light necessary for excitation of fluorescent protein, we found that by using either nude mouse or eliminating hair with hair removal reagents much of this problem is eliminated. Using a CCD camera, the fluorescent signal can be detected in the tibialis anterior (TA) muscle after injection of 5 x 105 cells from either GFP transgenic mice or eGFP transduced myoblast culture. For more superficial muscles such as the extensor digitorum longus (EDL), injection of fewer cells produces a detectable signal. Signal intensity can be measured and quantified as the number of emitted photons per second in a region of interest (ROI). Since the acquired images show clear boundaries demarcating the engrafted area, the size of the ROI can be measured as well. If the legs are positioned consistently every time, the changes in total number of photons per second per muscle and the size of the ROI reflect the changes in the number of engrafted cells and the size of the engrafted area. Therefore the changes in the same muscle over time are quantifiable. In vivo fluorescent imaging technique has been used primarily to track the growth of tumorogenic cells, our study shows that it is a powerful tool that enables us to track the fate of transplanted stem cells.

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

We describe an in vivo fluorescence imaging protocol to monitor muscle regeneration by GFP-labeled myoblasts after transplantation into skeletal muscles of both healthy and dystrophic mice. This protocol can be adapted to study muscle regeneration by transplantation of other types of cells and in other muscular conditions as well.

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Neuroscience

Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated measurements over weeks or months of accessory respiratory muscle activity while simultaneously measuring ventilation in a non-anesthetized, freely behaving mouse. The technique includes the surgical implantation of a radio transmitter and the insertion of electrode leads into the scalene and trapezius muscles to measure the electromyogram activity of these inspiratory muscles. Ventilation is measured by whole-body plethysmography, and animal movement is assessed by video and is synchronized with electromyogram activity. Measurements of muscle activity and ventilation in a mouse model of amyotrophic lateral sclerosis are presented to show how this tool can be used to investigate how respiratory muscle activity changes over time and to assess the impact of muscle activity on ventilation. The described methods can easily be adapted to measure the activity of other muscles or to assess accessory respiratory muscle activity in additional mouse models of disease or injury.

This paper introduces a method for repeated measurements of ventilation and respiratory muscle activity in a freely behaving amyotrophic lateral sclerosis (ALS) mouse model throughout disease progression with whole-body plethysmography and electromyography via an implanted telemetry device.

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated ...

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings provide a sensitive approach to measure nerve conduction in humans and rodent models. To broaden the technical possibilities for electromyography in mice, the measurement of compound muscle action potentials (CMAPs) from the brachial plexus nerve in the forelimb using needle electrodes is described here. CMAP recordings after stimulating the sciatic nerve in hindlimbs have been previously described. The newly introduced method here allows for the evaluation of the nerve conductivity at an additional site, and thus provides a more profound overview of the neuromuscular functionality. The technique provides information on both the relative number of functional axons and the myelination level. Thereby, this method can be applied to assess both axonal diseases as well as demyelinating conditions. This minimally invasive method does not require extraction of the nerve and therefore it is suitable for repeated measurements for longitudinal follow-up in the same animal. Similar recordings are performed in clinical setups to emphasize the translational relevance of the method.

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

The measurement of nerve conduction is a useful tool to assess mouse models of neurodegeneration but it is frequently only applied to stimulate the sciatic nerve in hindlimbs. Here, we describe a technique to measure compound muscle action potential (CMAP) in vivo in the mouse forelimb muscles innervated by the brachial plexus.

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings ...

Isometric and Eccentric Force Generation Assessment of Skeletal Muscles Isolated from Murine Models of Muscular Dystrophies

Critical to the evaluation of potential therapeutics for muscular disease are sensitive and reproducible physiological assessments of muscle function. Because many pre-clinical trials rely on mouse models for these diseases, isolated muscle function has become one of the standards for Go/NoGo decisions in moving drug candidates forward into patients. We will demonstrate the preparation of the extensor digitorum longus (EDL) and diaphragm muscles for functional testing, which are the predominant muscles utilized for these studies. The EDL muscle geometry is ideal for isolated muscle preparations, with two easily accessible tendons, and a small size that can be supported by superfusion in a bath. The diaphragm exhibits profound progressive pathology in dystrophic animals, and can serve as a platform for evaluating many potential therapies countering fibrosis, and promoting myofiber stability. Protocols for routine testing, including isometric and eccentric contractions, will be shown. Isometric force provides assessment of strength, and eccentric contractions help to evaluate sarcolemma stability, which is disrupted in many types of muscular dystrophies. Comparisons of the expected results between muscles from wildtype and dystrophic muscles will also be provided. These measures can complement morphological and biochemical measurements of tissue homeostasis, as well as whole animal assessments of muscle function.

Muscle function measurements contribute to the evaluation of potential therapeutics for muscle pathology, as well as to the determination of mechanisms underlying physiology of this tissue. We will demonstrate the preparation of the extensor digitorum longus and diaphragm muscles for functional testing. Protocols for isometric and eccentric contractions will be shown, as well as differences in results between dystrophic muscles, representing a pathological state, and wildtype muscles.

Critical to the evaluation of potential  therapeutics for muscular disease are sensitive and reproducible physiological  assessments of muscle function. ...

Evaluation of Muscle Function of the Extensor Digitorum Longus Muscle Ex vivo and Tibialis Anterior Muscle In situ in Mice

Body movements are mainly provided by mechanical function of skeletal muscle. Skeletal muscle is composed of numerous bundles of myofibers that are sheathed by intramuscular connective tissues. Each myofiber contains many myofibrils that run longitudinally along the length of the myofiber. Myofibrils are the contractile apparatus of muscle and they are composed of repeated contractile units known as sarcomeres. A sarcomere unit contains actin and myosin filaments that are spaced by the Z discs and titin protein. Mechanical function of skeletal muscle is defined by the contractile and passive properties of muscle. The contractile properties are used to characterize the amount of force generated during muscle contraction, time of force generation and time of muscle relaxation. Any factor that affects muscle contraction (such as interaction between actin and myosin filaments, homeostasis of calcium, ATP/ADP ratio, etc.) influences the contractile properties. The passive properties refer to the elastic and viscous properties (stiffness and viscosity) of the muscle in the absence of contraction. These properties are determined by the extracellular and the intracellular structural components (such as titin) and connective tissues (mainly collagen) 1-2. The contractile and passive properties are two inseparable aspects of muscle function. For example, elbow flexion is accomplished by contraction of muscles in the anterior compartment of the upper arm and passive stretch of muscles in the posterior compartment of the upper arm. To truly understand muscle function, both contractile and passive properties should be studied.
The contractile and/or passive mechanical properties of muscle are often compromised in muscle diseases. A good example is Duchenne muscular dystrophy (DMD), a severe muscle wasting disease caused by dystrophin deficiency 3. Dystrophin is a cytoskeletal protein that stabilizes the muscle cell membrane (sarcolemma) during muscle contraction 4. In the absence of dystrophin, the sarcolemma is damaged by the shearing force generated during force transmission. This membrane tearing initiates a chain reaction which leads to muscle cell death and loss of contractile machinery. As a consequence, muscle force is reduced and dead myofibers are replaced by fibrotic tissues 5. This later change increases muscle stiffness 6. Accurate measurement of these changes provides important guide to evaluate disease progression and to determine therapeutic efficacy of novel gene/cell/pharmacological interventions. Here, we present two methods to evaluate both contractile and passive mechanical properties of the extensor digitorum longus (EDL) muscle and the contractile properties of the tibialis anterior (TA) muscle.

Evaluation of Muscle Function of the Extensor Digitorum Longus Muscle Ex vivo and Tibialis Anterior Muscle In situ in Mice

Changes in limb muscle contractile and passive mechanical properties are important biomarkers for muscle diseases. This manuscript describes physiological assays to measure these properties in the murine extensor digitorum longus and tibialis anterior muscles.

Body  movements are mainly provided by mechanical function of skeletal muscle. Skeletal  muscle is composed of numerous bundles of myofibers that are ...

Non-invasive Assessment of Dorsiflexor Muscle Function in Mice

Assessment of skeletal muscle contractile function is an important measurement for both clinical and research purposes. Numerous conditions can negatively affect skeletal muscle. This can result in a loss of muscle mass (atrophy) and/or loss of muscle quality (reduced force per unit of muscle mass), both of which are prevalent in chronic disease, muscle-specific disease, immobilization, and aging (sarcopenia). Skeletal muscle function in animals can be evaluated by a range of different tests. All tests have limitations related to the physiological testing environment, and the selection of a specific test often depends on the nature of the experiments. Here, we describe an in vivo, non-invasive technique involving a helpful and easy assessment of force frequency-curve (FFC) in mice that can be performed on the same animal over time. This permits monitoring of disease progression and/or efficacy&#160;of a potential therapeutic treatment.

Measurement of rodent skeletal muscle contractile function is a useful tool that can be used to track disease progression as well as efficacy of therapeutic intervention. We describe here the non-invasive, in vivo assessment of the dorsiflexor muscles that can be repeated over time in the same mouse.

Assessment of skeletal muscle contractile function is an important measurement for both clinical and research purposes. Numerous conditions can negatively ...

Measurement of Insulin- and Contraction-Stimulated Glucose Uptake in Isolated and Incubated Mature Skeletal Muscle from Mice

Skeletal muscle is an insulin-responsive tissue and typically takes up most of the glucose that enters the blood after a meal. Moreover, it has been reported that skeletal muscle may increase the extraction of glucose from the blood by up to 50-fold during exercise compared to resting conditions. The increase in muscle glucose uptake during exercise and insulin stimulation is dependent on the translocation of glucose transporter 4 (GLUT4) from intracellular compartments to the muscle cell surface membrane, as well as phosphorylation of glucose to glucose-6-phosphate by hexokinase II. Isolation and incubation of mouse muscles such as m. soleus and m. extensor digitorum longus (EDL) is an appropriate ex vivo model to study the effects of insulin and electrically-induced contraction (a model for exercise) on glucose uptake in mature skeletal muscle. Thus, the ex vivo model permits evaluation of muscle insulin sensitivity and makes it possible to match muscle force production during contraction ensuring uniform recruitment of muscle fibers during measurements of muscle glucose uptake. Moreover, the described model is suitable for pharmacological compound testing that may have an impact on muscle insulin sensitivity or may be of help when trying to delineate the regulatory complexity of skeletal muscle glucose uptake.
Here we describe and provide a detailed protocol on how to measure insulin- and contraction-stimulated glucose uptake in isolated and incubated soleus and EDL muscle preparations from mice using radiolabeled [3H]2-deoxy-D-glucose and [14C]mannitol as an extracellular marker. This allows accurate assessment of glucose uptake in mature skeletal muscle in the absence of confounding factors that may interfere in the intact animal model. In addition, we provide information on metabolic viability of incubated mouse skeletal muscle suggesting that the method&#160;applied possesses some caveats under certain conditions when studying muscle energy metabolism.

Intact regulation of muscle glucose uptake is important for maintaining whole body glucose homeostasis. This protocol presents assessment of insulin- and contraction-stimulated glucose uptake in isolated and incubated mature skeletal muscle when delineating the impact of various physiological interventions on whole body glucose metabolism.

Skeletal muscle is an insulin-responsive tissue and typically takes up most of the glucose that enters the blood after a meal. Moreover, it has been ...

In Vivo Measurement of Hindlimb Dorsiflexor Isometric Torque from Pig

Reliable assessment of skeletal muscle strength is arguably the most important outcome measure in neuromuscular and musculoskeletal disease and injury studies, particularly when evaluating regenerative therapies&#39; efficacy. Additionally, a critical aspect of translating many regenerative therapies is the demonstration of scalability and effectiveness in a large animal model. Various physiological preparations have been established to evaluate intrinsic muscle function properties in basic science studies, primarily in small animal models. The practices may be categorized as: in vitro (isolated fibers, fiber bundles, or whole muscle), in situ (muscle with intact vascularization and innervation but distal tendon attached to a force transducer), and in vivo (structures of the muscle or muscle unit remain intact). There are strengths and weaknesses to each of these preparations; however, a clear advantage of in vivo strength testing is the ability to perform repeated measurements in the same animal. Herein, the materials and methods to reliably assess isometric torque produced by the hindlimb dorsiflexor muscles in vivo in response to standard peroneal electrical stimulation in anesthetized pigs are presented.

In Vivo Measurement of Hindlimb Dorsiflexor Isometric Torque from Pig

The present protocol describes concise experimental details on the evaluation and interpretation of in vivo torque data obtained via electrical stimulation of the common peroneal nerve in anesthetized pigs.

Reliable assessment of skeletal muscle strength is arguably the most important outcome measure in neuromuscular and musculoskeletal disease and injury ...

In vivo Measurement of Knee Extensor Muscle Function in Mice

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