Name: repeated measurement of respiratory muscle activity and ventilation in mouse models of neuromuscular disease
Uploaded: 2017-04-17T14:00:00.000+00:00
Duration: 9 min 24 s
Description: Watch this Scientific Journal Video about Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease at JoVE.com

Support for this work was provided by a Cincinnati Children&#39;s Hospital Medical Center Trustee Award to S.A.C. and an NIH training grant (T32NS007453) to V.N.J.

<ol>
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H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Accessory+respiratory+muscles+enhance+ventilation+in+ALS+model+mice+and+are+activated+by+excitatory+V2a+neurons.">Accessory respiratory muscles enhance ventilation in ALS model mice and are activated by excitatory V2a neurons.</a> Exp Neurol. 287 (Pt. 2, 192-204 (2017).</li><li>Moldovan, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nerve+excitability+changes+related+to+axonal+degeneration+in+amyotrophic+lateral+sclerosis:+Insights+from+the+transgenic+SOD1(G127X)+mouse+model.">Nerve excitability changes related to axonal degeneration in amyotrophic lateral sclerosis: Insights from the transgenic SOD1(G127X) mouse model.</a> Exp Neurol. 233 (1), 408-420 (2012).</li><li>Pagliardini, S., Gosgnach, S., Dickson, C. 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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+potential+early+physiological+marker+for+CNS+oxygen+toxicity:+hyperoxic+hyperpnea+precedes+seizure+in+unanesthetized+rats+breathing+hyperbaric+oxygen.">A potential early physiological marker for CNS oxygen toxicity: hyperoxic hyperpnea precedes seizure in unanesthetized rats breathing hyperbaric oxygen.</a> J Appl Physiol. 114 (1985), 1009-1020 (1985).</li><li>Morrison, J. 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The authors have nothing to disclose.

The procedure demonstrated here allows for the noninvasive (after initial surgical implantation of the transmitter) measurement of respiratory muscle activity and ventilation over many months in the same animal. This technique has several advantages over standard EMG techniques in anesthetized mice: 1) the experiments require fewer mice and provide the ability to record data from the same site in a single mouse across disease stages (instead of using multiple mice at different disease stages); 2) data analysis can be performed with more powerful statistical tests (i.e., using repeated measures instead of comparing separate experimental groups); 3) the simultaneous recording of EMG and WBP allows for the direct assessment of the effects of ARM activity on ventilation; and 4) the experiments can be performed on mice in different states of sleep/wakefulness. Moreover, because healthy mice have a very low frequency of ARM bouts at rest, this technique is capable of detecting even small changes in the frequency of ARM activity in ALS model mice at early symptomatic stages of disease10. However, because this technique measures the activity of a large but unknown number of muscle fibers during natural behavior, rather than following nerve stimulation at experimentally controlled intensities, it is not suitable for estimating motor unit size or number. Another limitation is that telemetry-based transmitters suitable for implantation in mice are currently limited to two sets of biopotential leads; thus, only two sites can be recorded from the same mouse. For experiments requiring the simultaneous recording of more than two muscles in the same mouse, multiple EMG leads may be implanted and connected to an acquisition system using a wire tether, as previously described14,15. However, modifications to the plethysmography chamber or seals would be required to allow for the simultaneous recording of muscle activity and ventilation if a mouse is tethered.
When applying this technique, certain steps of the protocol must be carried out with care. Biopotential leads must be placed so that they do not impede movement or irritate the overlying skin. In addition, the transmitter must be positioned so that it does not affect the normal movement or posture of the mouse. It is recommended that the proper placement of leads (i.e., fully embedded in the correct muscle and not contacting adjacent muscles) and the lack of muscle damage or infection are verified by autopsy after the experiment has been terminated. Further, it is imperative that the transmitter is turned off after every recording session to conserve battery life.
An unavoidable consequence of measuring EMG from muscles in the chest and neck area is the high likelihood of recording electrocardiogram (ECG) signals, which appear as regular spikes within the EMG trace (Figure 3B, arrowhead). ECG signals can be minimized by careful placement of the leads such that all metal is embedded fully in the muscle and by avoiding placement near major blood vessels. Implanting leads into muscles on the right side of the body rather than the left, which is closer to the heart, can also reduce ECG signals. Although the ECG signal may be filtered out of EMG traces using computational algorithms or by subtracting an independently recorded ECG signal19,20,21, it is not typically necessary. The ECG signal can be readily distinguished from the EMG signal by its regular shape, frequency, and amplitude.
The described technique has been used to measure changes in ARM activity at rest in the SOD1(G93A) mouse model of ALS10. Accessory respiratory muscles are also recruited in other neuromuscular diseases (e.g., muscular dystrophy, spinal muscular atrophy, peripheral neuropathies, etc.) and following nerve or spinal cord injuries. ARM activity may therefore serve as a proxy to measure functional impairment of the diaphragm and gauge the severity of disease, monitor recovery from injury, or assess potential treatment benefits to improve breathing in a variety of animal disease or injury models.

Accessory respiratory muscles (ARMs) increase ventilation during times of high demand (e.g., exercise) and help to maintain ventilation when diaphragm function is compromised following injury or disease1,2. Although changes in diaphragm function have been well described in amyotrophic lateral sclerosis (ALS) patients and mouse models3,4,5,6, much less is known about the activity or function of ARMs in ALS. However, one study suggested that ALS patients that recruit ARMs have a better prognosis than those with similar diaphragm dysfunction that do not7. Furthermore, ARM activity is sufficient for respiration in cases of diaphragm paralysis8. These studies indicate that strategies to augment ARM function may improve breathing in patients suffering from neuromuscular disease, spinal cord injury, or other conditions in which diaphragm function is impaired. However, the mechanisms controlling ARM recruitment for breathing are largely unknown. Methods to measure respiratory function and changes in ARM activity over time in animal models of disease or injury are needed to study how ARMs are recruited, as well as to evaluate therapies to improve ARM recruitment and ventilation. Moreover, the increased activity of ARMs coinciding with the progressive loss of diaphragm function may be a useful biomarker for disease progression in neuromuscular diseases such as ALS7,9,10.
This protocol describes a method to non-invasively (following the initial surgery) and repeatedly measure the activity of respiratory muscles and ventilation in awake, behaving mice. Synchronized recordings of electromyography (EMG), whole-body plethysmography (WBP), and video allow the investigator to assess how changes in ARM activity impact ventilation and to determine when the subject is at rest or moving. A major advantage of this method is that it can be performed in awake, behaving mice, whereas some alternative methods to measure EMG require anesthesia and/or are terminal procedures11,12,13. The recording of EMG activity in awake mice over time may also be accomplished through the chronic implantation of EMG leads, where the mouse is tethered by wires to the acquisition system14,15. Because tethering a mouse could interfere with normal movement or behavior and may not be compatible with a standard plethysmography chamber, the described method uses telemetry devices to wirelessly transmit the EMG signal to the acquisition system. The transmitter can be turned on or off with a magnet to conserve battery power and allows repeated measurements of EMG activity over several months. This protocol can be easily adapted to measure the activity of additional respiratory or non-respiratory muscles by inserting the EMG leads into different muscles. Alternatively, one of the two leads may be used to measure EEG activity to assess sleep state or to identify seizure activity16. This technique has successfully been used to measure changes in ARM activity at rest throughout disease progression in a mouse model of ALS and to identify key neurons driving ARM activity in healthy mice10.

<table><tbody><tr><td>B6.Cg-Tg (SOD1*G93A)1 Gur/J</td><td>Jackson Laboratory</td><td>4435</td><td></td></tr><tr><td>Plethysmography Chamber</td><td>Buxco Respiratory Products/ Data Sciences International</td><td>601-1425-001</td><td></td></tr><tr><td>Telemetry Receivers (Model RPC-1)</td><td>Data Sciences International</td><td>272-6001-001</td><td></td></tr><tr><td>Bias Flow Pump (Model BFL0500)</td><td>Data Sciences International</td><td>601-2201-001</td><td></td></tr><tr><td>ACQ-7000 USB</td><td>Data Sciences International</td><td>PNM-P3P-7002XS</td><td></td></tr><tr><td>Dataquest A.R.T. Data Exchange Matrix</td><td>Data Sciences International</td><td>271-0117-001</td><td></td></tr><tr><td>New Ponemah Analysis System</td><td>Data Sciences International</td><td>PNM-POST-CFG</td><td></td></tr><tr><td>Ponemah Physiology Platform Acqusition software v5.20</td><td>Data Sciences International</td><td>PNM-P3P-520</td><td></td></tr><tr><td>Ponemah Unrestrained Whole Breath Plethysmography analysis package v5.20</td><td>Data Sciences International</td><td>PNM-URP100W</td><td></td></tr><tr><td>Configured Ponemah Software System</td><td>Data Sciences International</td><td>PNM-P3P-CFG</td><td></td></tr><tr><td>Analysis Module (URP)</td><td>Data Sciences International</td><td>PNM-URP100W</td><td></td></tr><tr><td>Universal Amplifier</td><td>Data Sciences International</td><td>13-7715-59</td><td></td></tr><tr><td>Sync Board</td><td>Data Sciences International</td><td>271-0401-001</td><td></td></tr><tr><td>Sync Cable</td><td>Data Sciences International</td><td>274-0030-001</td><td></td></tr><tr><td>Transducer-Pressure Buxco</td><td>Data Sciences International</td><td>600-1114-001</td><td></td></tr><tr><td>Flow Meter</td><td>Data Sciences International</td><td>600-1260-001</td><td></td></tr><tr><td>Magnet and Radio included in F20-EET Starter Kit</td><td>Data Sciences International</td><td>276-0400-001</td><td></td></tr><tr><td>Axis P1363 Video Camera&amp;nbsp;&amp;nbsp;</td><td>Data Sciences International</td><td>275-0201-001</td><td></td></tr><tr><td>Terg-A-Zyme</td><td>Fisher Scientific</td><td>50-821-785</td><td>Enzyme Detergent</td></tr><tr><td>Actril</td><td>Minntech Corporation</td><td>78337-000</td><td>Chemical Sterilant</td></tr><tr><td>Stereo Dissecting Microscope (Model MEB126)</td><td>Leica</td><td>10-450-508</td><td></td></tr><tr><td>Servo-Controlled Humidifier/Infant Incubator</td><td>OHMEDA Ohio Care Plus</td><td>6600-0506-803</td><td></td></tr><tr><td>TL11M2-F20-EET Transmitters</td><td>Data Sciences International</td><td>270-0124-001</td><td></td></tr><tr><td>Dumont #2 Laminectomy Forceps - Standard Tips/Straight/12 cm (x2)&amp;nbsp;</td><td>Fine Scientific Instruments</td><td>11223-20</td><td>For handling wires</td></tr><tr><td>Dumont #2 Laminectomy Forceps - Standard Tips/Straight/12 cm (x2)</td><td>Fine Scientific Instruments</td><td>11223-20</td><td>For surgery</td></tr><tr><td>Narrow Pattern Forceps- Serrated/Curved/12 cm</td><td>Fine Scientific Instruments</td><td>17003-12</td><td></td></tr><tr><td>Spring Scissors - Tough Cut/Straight/Sharp/12.5 cm/6 mm Cutting Edge</td><td>Fine Scientific Instruments</td><td>15124-12</td><td></td></tr><tr><td>Tissue Separating Scissors - Straight/Blunt-Blunt/11.5 cm</td><td>Fine Scientific Instruments</td><td>14072-10</td><td></td></tr><tr><td>Fine Scissors - Tough Cut/Curved/Sharp-Sharp/9 cm&amp;nbsp;</td><td>Fine Scientific Instruments</td><td>14058-11</td><td>For cutting wires and clipping nails</td></tr><tr><td>Scalpel Handle #3</td><td>World Precision Instruments</td><td>500236</td><td></td></tr><tr><td>Scalpel Blade</td><td>Fine Scientific Instruments</td><td>10010-00</td><td>For preparing lead caps</td></tr><tr><td>Polysorb Braided Absorbable suture</td><td>Coviden</td><td>D4G1532X</td><td>For coiling transmitter leads</td></tr><tr><td>Gluture&amp;nbsp;</td><td>Zoetis Inc.</td><td>6606-65-1</td><td>Cyanoacrylate adhesive</td></tr><tr><td>3 mL Syring Slip Tip - Soft</td><td>Vitality Medical</td><td>118030055</td><td></td></tr><tr><td>25 G Needle (x2)</td><td>Becton Dickinson and Co.</td><td>305-145</td><td></td></tr><tr><td>Cotton Tipped Applicators</td><td>Henry Schein Animal Health</td><td>100-9175</td><td></td></tr><tr><td>Andis Easy Cut Hair Clipper Set</td><td>Andis</td><td>049-06-0271</td><td>Electrical Razor sold at Target</td></tr><tr><td>Isoflurane</td><td>Henry Schein Animal Health</td><td>29404</td><td>Anesthetic&amp;nbsp;</td></tr><tr><td>Isopropyl Alcohol 70%</td><td>Priority Care 1</td><td>MS070PC</td><td></td></tr><tr><td>Dermachlor 2% Medical Scrub (chlorohexidine 2%)</td><td>Butler Schein</td><td>55482</td><td></td></tr><tr><td>Artificial Tears</td><td>Henry Schein Animal Health</td><td>48272</td><td>Lubricant Opthalmic Ointment</td></tr><tr><td>Vacuum grease</td><td>Dow Corning Corporation</td><td>1597418</td><td></td></tr><tr><td>Water Blanket</td><td>JorVet</td><td>JOR784BN</td><td></td></tr></tbody></table>

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.

The described protocol was used to implant a telemetry device and to record scalene and trapezius EMG, WBP, and video of a SOD1(G93A) ALS model mouse. Periods in which the animal is inactive (e.g., does not move) were identified using the video recording and confirmed by the lack of movement-related activity in the WBP trace (Figure 3A). Inactive periods include time spent in REM or non-REM sleep, as well as time spent awake but still (Figure 3A). EMG activity during this inactive time was scored as a bout when at least 3 consecutive rectified and integrated (over 30 ms) values had amplitudes with at least a 50% increase over baseline EMG levels (Figure 4). Bouts of activity that occurred during sighing or sniffing (determined by plethysmography), or volitional movements (assessed by video) were excluded from analysis (Figure 3B-C). SOD1(G93A) mice at early- to mid-symptomatic stages (Table 1) exhibit bouts of increased ARM activity at rest that last for one to several breaths (Figure 4). Bouts of ARM activity are rare in pre-symptomatic SOD1(G93A) (Figure 3A) or wildtype mice10.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Stage</td>
			<td>State</td>
			<td>Stage Onset</td>
			<td>Hindlimb Presentation</td>
		</tr>
		<tr>
			<td>0</td>
			<td>Pre-symptomatic</td>
			<td>&#60; P100</td>
			<td>No notable differences compared to wildtypes.</td>
		</tr>
		<tr>
			<td>1</td>
			<td>Disease onset</td>
			<td>~ P100</td>
			<td>Hindlimb collapse when mouse is suspended from tail.</td>
		</tr>
		<tr>
			<td>2</td>
			<td>Paresis</td>
			<td>~ P120</td>
			<td>Full or partial hindlimb collapse with appearance of tremor.</td>
		</tr>
		<tr>
			<td>3</td>
			<td>Paralysis onset</td>
			<td>~ P140</td>
			<td>Difficulty walking, toe curling and/or foot dragging.</td>
		</tr>
		<tr>
			<td>4</td>
			<td>Advanced paralysis</td>
			<td>~ P150</td>
			<td>&#160;Minimal joint movement, hindlimb not being used for forward motion.</td>
		</tr>
		<tr>
			<td>5</td>
			<td>Endstage</td>
			<td>~ P160</td>
			<td>Mouse unable to right itself from side within 30 seconds.</td>
		</tr>
	</tbody>
</table>
Table 1. Neurological Scoring of ALS-like Disease Progression in SOD1(G93A) Mice.
<img alt="Figure 3" src="/files/ftp_upload/55599/55599fig3.jpg" /> 
Figure 3. Representative WBP and EMG Traces. (A-C) WBP and EMG of scalene and trapezius muscles from a pre-symptomatic SOD1(G93A) mouse (age P98). (A) Periods when the animal is at rest (red box) are used for analysis. Traces outside the red box show large and irregular peaks in the plethysmography traces and muscle activity in the EMG traces, typical when an animal is moving, as determined by synchronized video recordings (not shown). The red box shows EMG traces lacking EMG bouts, characteristic of a pre-symptomatic mouse. (B) Bouts of EMG activity frequently occur directly preceding a sigh (as shown in the plethysmography trace). Sighs are characterized by high-amplitude inspiration followed by dramatic expiration. The black arrowhead points to a characteristic ECG signal. (C) Bouts of EMG activity frequently occur while the mouse is sniffing. Sniffing is reflected in the plethysmography trace by a prolonged increase in both frequency and amplitude over multiple breaths (co-occurring with bursts of EMG activity). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55599/55599fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55599/55599fig4.jpg" /> 
Figure 4. Scoring Bouts of EMG Activity. (A and B) Two examples of WBP, filtered trapezius EMG traces, and rectified and integrated trapezius EMG signals from a symptomatic SOD1(G93A) mouse (age P126). Blue dotted lines indicate baseline EMG level, determined by averaging rectified and integrated signals over a time period of 3 s. Red dotted lines indicate a 50% increase in amplitude over baseline EMG activity. A bout of activity is scored when at least 3 consecutive rectified and integrated values exceed the 50% baseline threshold. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55599/55599fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>PIF (mL/s)</td>
			<td>TV (mL)</td>
			<td>MV (mL/min)</td>
			<td>Breathing Frequency (Breaths/min)</td>
		</tr>
		<tr>
			<td>Naive (n=5)</td>
			<td>4.4 &#177; 0.7</td>
			<td>0.27 &#177; 0.04</td>
			<td>58 &#177; 13</td>
			<td>223 &#177; 41</td>
		</tr>
		<tr>
			<td>Implanted (n=4)</td>
			<td>4.1 &#177; 0.2</td>
			<td>0.27 &#177; 0.11</td>
			<td>56 &#177; 29</td>
			<td>201 &#177; 32</td>
		</tr>
		<tr>
			<td>P-value</td>
			<td>0.439</td>
			<td>1.000</td>
			<td>0.893</td>
			<td>0.410</td>
		</tr>
		<tr>
			<td colspan="5">Values shown reflect mean &#177; SD. P-values were calculated with a Student&#39;s t-test.</td>
		</tr>
	</tbody>
</table>
Table 2. Comparison of Respiration Between Naive (not Implanted) and Implanted Stage 4 SOD1(G93A) Mice. No significant differences were found in peak inspiratory flow (PIF), tidal volume (TV), minute volume (MV), or breaths per min between the two groups. The values shown reflect the mean &#177; SD. P-values were calculated with a Student&#39;s t-test.
Repeated measurements of EMG and/or WBP can be made in the same mouse over several months, with very little change in EMG signal or baseline after a 1- to 2-week recovery period following surgery. The time course is typically limited by the battery life and thus will be determined by the frequency and duration of the individual recordings. Researchers should be aware that adverse events due to the implanted device may occasionally occur. The mouse may pull out the wires from the implanted muscle or scratch/chew at the skin if the leads or transmitter are improperly placed. In most cases, ethical considerations dictate that these animals be sacrificed. The transmitter may be removed, sterilized, and re-implanted in another mouse.
To verify that device implantation does not affect breathing, plethysmography measurements between naive SOD1(G93A) mice (not implanted) at ALS stage 4 and implanted SOD1(G93A) mice at ALS stage 4 were compared. No significant differences were found in peak inspiratory flow (PIF), tidal volume (TV), minute volume (MV), or breaths per minute between the two groups (Table 2). The scalene and trapezius are adjacent to each other and are directly in contact with one another. Although simultaneous EMG bouts are sometimes observed in both muscles, strong EMG bouts are also detected in the trapezius when EMG bouts are absent in the scalene (and vice versa), demonstrating that there is minimal cross-talk between electrodes implanted in each muscle. Independent bouts of EMG activity are also observed when leads are placed in the trapezius and sternocleidomastoid muscles (data not shown).

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

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 ...

repeated-measurement-respiratory-muscle-activity-ventilation-mouse

Nanyang Technological University

Research

JoVE Journal

Medicine

12.9K Views.  University of Cincinnati.  Dit artikel beschrijft een werkwijze voor herhaalde metingen van ventilatie en respiratoire spieractiviteit in vrij gedragen amyotrofische laterale sclerose (ALS) muismodel gedurende ziekteprogressie met hele-lichaamsplethysmografie en elektromyografie via een geïmplanteerde apparaat telemetrie. 

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

Electrophysiological Motor Unit Number Estimation (MUNE) Measuring Compound Muscle Action Potential (CMAP) in Mouse Hindlimb Muscles

Compound muscle action potential (CMAP) and motor unit number estimation (MUNE) are electrophysiological techniques that can be used to monitor the functional status of a motor unit pool in vivo. These measures can provide insight into the normal development and degeneration of the neuromuscular system. These measures have clear translational potential because they are routinely applied in diagnostic and clinical human studies. We present electrophysiological techniques similar to those employed in humans to allow recordings of mouse sciatic nerve function. The CMAP response represents the electrophysiological output from a muscle or group of muscles following supramaximal stimulation of a peripheral nerve. MUNE is an electrophysiological technique that is based on modifications of the CMAP response. MUNE is a calculated value that represents the estimated number of motor neurons or axons (motor control input) supplying the muscle or group of muscles being tested. We present methods for recording CMAP responses from the proximal leg muscles using surface recording electrodes following the stimulation of the sciatic nerve in mice. An incremental MUNE technique is described using submaximal stimuli to determine the average single motor unit potential (SMUP) size. MUNE is calculated by dividing the CMAP amplitude (peak-to-peak) by the SMUP amplitude (peak-to-peak). These electrophysiological techniques allow repeated measures in both neonatal and adult mice in such a manner that facilitates rapid analysis and data collection while reducing the number of animals required for experimental testing. Furthermore, these measures are similar to those recorded in human studies allowing more direct comparisons.

Electrophysiological Motor Unit Number Estimation MUNE Measuring Compound Muscle Action Potential CMAP in Mouse Hindlimb Muscles

We present refined protocols that allow in vivo monitoring of motor unit function in the mouse. Techniques to measure compound muscle action potential (CMAP) and motor unit number estimation (MUNE) in the mouse hind limb muscles innervated by the sciatic nerve are described.

Compound muscle action potential (CMAP) and motor unit number estimation (MUNE) are electrophysiological techniques that can be used to monitor the ...

Behavior

A Simple and Low-cost Assay for Measuring Ambulation in Mouse Models of Muscular Dystrophy

Measuring functional outcomes in the treatment of muscular dystrophy is an essential aspect of preclinical testing. The assessment of voluntary ambulation in mouse models is a non-invasive and reproducible activity assay that is directly analogous to measures of patient ambulation such as the 6-minute walk test and related mobility scores. Many common methods for testing mouse ambulation speed and distance are based on the open field test, where an animal&#39;s free movement within an arena is measured over time. One major downside to this approach is that commercial software and equipment for high-resolution motion tracking is expensive and may require transferring mice to specialized facilities for testing. Here, we describe a low-cost, video-based system for measuring mouse ambulation that utilizes free and open-source software. Using this protocol, we demonstrate that voluntary ambulation in the dystrophin-null mdx mouse model for Duchenne muscular dystrophy (DMD) is decreased relative to wild-type mouse activity. In mdx mice expressing the utrophin transgene, these activity deficits are not observed and the total distance traveled is indistinguishable from wild-type mice. This method is effective for measuring changes in voluntary ambulation associated with dystrophic pathology, and provides a versatile platform that can be readily adapted to diverse research settings.

This protocol describes a flexible, low-cost system for measuring mouse ambulation in an open field activity assay. We show that a 6-minute ambulation assay based on this system detects a decrease in voluntary movement in mdx mice, and accurately distinguishes improvement in a muscle-specific rescue of these animals.

Measuring functional outcomes in the treatment of muscular dystrophy is an essential aspect of preclinical testing. The assessment of voluntary ambulation ...

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment (RMCA) in Individuals with Chronic Spinal Cord Injury

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination is disrupted by spinal cord injury (SCI), control of respiratory muscles innervated below the injury level is compromised1,2 leading to respiratory muscle dysfunction and pulmonary complications. These conditions are among the leading causes of death in patients with SCI3. Standard pulmonary function tests that assess respiratory motor function include spirometrical and maximum airway pressure outcomes: Forced Vital Capacity (FVC), Forced Expiratory Volume in one second (FEV1), Maximal Inspiratory Pressure (PImax) and Maximal Expiratory Pressure (PEmax)4,5. These values provide indirect measurements of respiratory muscle performance6. In clinical practice and research, a surface electromyography (sEMG) recorded from respiratory muscles can be used to assess respiratory motor function and help to diagnose neuromuscular pathology. However, variability in the sEMG amplitude inhibits efforts to develop objective and direct measures of respiratory motor function6. Based on a multi-muscle sEMG approach to characterize motor control of limb muscles7, known as the voluntary response index (VRI)8, we developed an analytical tool to characterize respiratory motor control directly from sEMG data recorded from multiple respiratory muscles during the voluntary respiratory tasks. We have termed this the Respiratory Motor Control Assessment (RMCA)9. This vector analysis method quantifies the amount and distribution of activity across muscles and presents it in the form of an index that relates the degree to which sEMG output within a test-subject resembles that from a group of healthy (non-injured) controls. The resulting index value has been shown to have high face validity, sensitivity and specificity9-11. We showed previously9 that the RMCA outcomes significantly correlate with levels of SCI and pulmonary function measures. We are presenting here the method to quantitatively compare post-spinal cord injury respiratory multi-muscle activation patterns to those of healthy individuals.

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment RMCA in Individuals with Chronic Spinal Cord Injury

The purpose of this publication is to present our original work on a multi-muscle surface electromyographic approach to quantitatively characterize respiratory muscle activation patterns in individuals with chronic spinal cord injury using vector-based analysis.

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination ...

A Reversible, Non-invasive Method for Airway Resistance Measurements and Bronchoalveolar Lavage Fluid Sampling in Mice

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated procedures to obtain such measurements in the same animal are generally not feasible.  Here, we demonstrate protocols for obtaining from mice repeated measurements of AHR and bronchoalveolar lavage fluid samples.  Mice were challenged intranasally seven times over 14 days with a potent allergen or sham treated.  Prior to the initial challenge, and within 24 hours following each intranasal challenge, the same animals were anesthetized, orally intubated and mechanically ventilated.  AHR, assessed by comparing dose response curves of respiratory system resistance (RRS) induced by increasing intravenous doses of acetylcholine (Ach) chloride between sham and allergen-challenged animals, were determined.  Afterwards, and via the same intubation, the left lung was lavaged so that differential enumeration of airway cells could be performed.  These studies reveal that repeated measurements of AHR and BAL fluid collection are possible from the same animals and that maximal airway hyperresponsiveness and airway eosinophilia are achieved within 7-10 days of initiating allergen challenge.  This novel technique significantly reduces the number of mice required for longitudinal experimentation and is applicable to diverse rodent species, disease models and airway physiology instruments.  

Repeated measurements of rodent respiratory physiology and sampling of airway inflammatory cells are desirable, but generally not feasible. Here we describe a repeatable method for orally intubating mice that permits repeated measurements of airway hyperreactivity and sampling of airway inflammatory cells.

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated ...

Biology

Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique

The forced oscillation technique (FOT) is a powerful, integrative and translational tool permitting the experimental assessment of lung function in mice in a comprehensive, detailed, precise and reproducible manner. It provides measurements of respiratory system mechanics through the analysis of pressure and volume signals acquired in reaction to predefined, small amplitude, oscillatory airflow waveforms, which are typically applied at the subject's airway opening. The present protocol details the steps required to adequately execute forced oscillation measurements in mice using a computer-controlled piston ventilator (flexiVent; SCIREQ Inc, Montreal, Qc, Canada). The description is divided into four parts: preparatory steps, mechanical ventilation, lung function measurements, and data analysis. It also includes details of how to assess airway responsiveness to inhaled methacholine in anesthetized mice, a common application of this technique which also extends to other outcomes and various lung pathologies. Measurements obtained in na&iuml;ve mice as well as from an oxidative-stress driven model of airway damage are presented to illustrate how this tool can contribute to a better characterization and understanding of studied physiological changes or disease models as well as to applications in new research areas.

The present protocol provides a detailed step-by-step description of the procedures required to execute measurements of respiratory system mechanics as well as the assessment of airway responsiveness to inhaled methacholine in mice using the forced oscillation technique (flexiVent; SCIREQ Inc, Montreal, Qc, Canada).

The  forced oscillation technique (FOT) is a powerful, integrative and translational  tool permitting the experimental assessment of lung function in mice ...

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to non-invasively assess some features of the respiratory function in humans as well as in laboratory animals. The present article focuses on the application of the double-chamber plethysmography (DCP) in small animals. It provides background information on the methodology as well as a detailed step-by-step procedure to successfully assess respiratory function in conscious, spontaneously breathing animals in a non-invasive manner. The DCP can be used to monitor the respiratory function of multiple animals in parallel, as well as to identify changes induced by aerosolized substances over a chosen time period and in a repeated manner. Experiments on control and allergic mice are used herein to demonstrate the utility of the technique, explain the associated outcome parameters, as well as to discuss the related advantages and shortcomings. Overall, the DCP provides valid and theoretically sound readouts that can be trusted to evaluate the respiratory function of conscious small animals both at baseline and after challenges with aerosolized substances.

The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to ...

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative assessment of lung function in rodent models is an important tool in the development of future therapies. Commonly used techniques for assessing respiratory function including invasive plethysmography and forced oscillation. While these techniques provide valuable information, data collection can be fraught with artefacts and experimental variability due to the need for anesthesia and/or invasive instrumentation of the animal. In contrast, unrestrained whole-body plethysmography (UWBP) offers a precise, non-invasive, quantitative way by which to analyze respiratory parameters. This technique avoids the use of anesthesia and restraints, which is common to traditional plethysmography techniques. This video will demonstrate the UWBP procedure including the equipment set up, calibration and lung function recording. It will explain how to analyze the collected data, as well as identify experimental outliers and artefacts that results from animal movement. The respiratory parameters obtained using this technique include tidal volume, minute volume, inspiratory duty cycle, inspiratory flow rate and the ratio of inspiration time to expiration time. UWBP does not rely on specialized skills and is inexpensive to perform. A key feature of UWBP, and most appealing to potential users, is the ability to perform repeated measures of lung function on the same animal.

The assessment of respiratory physiology has traditionally relied upon techniques, which require restraint or sedation of the animal. Unrestrained whole-body plethysmography, however, provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models. In addition, the technique allows repeated respiratory assessment of mice allowing for longitudinal studies.

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative ...

Quantifying Phrenic Motor Unit Function in Rodent Models of Neurodegeneration

Assessing Rat Diaphragm Motor Unit Connectivity Outcome Measures as Quantitative Biomarkers of Phrenic Motor Neuron Degeneration and Compensation

Loss of ventilatory muscle function is a consequence of motor neuron injury and neurodegeneration (e.g., cervical spinal cord injury and amyotrophic lateral sclerosis, respectively). Phrenic motor neurons are the final link between the central nervous system and muscle, and their respective motor units (groups of muscle fibers innervated by a single motor neuron) represent the smallest functional unit of the neuromuscular ventilatory system. Compound muscle action potential (CMAP), single motor unit potential (SMUP), and motor unit number estimation (MUNE) are established electrophysiological approaches that enable the longitudinal assessment of motor unit integrity in animal models over time but have mostly been applied to limb muscles. Therefore, the objectives of this study are to describe an approach in preclinical rodent studies that can be used longitudinally to quantify the phrenic MUNE, motor unit size (represented as SMUP), and CMAP, and then to demonstrate the utility of these approaches in a motor neuron loss model. Sensitive, objective, and translationally relevant biomarkers for neuronal injury, degeneration, and regeneration in motor neuron injury and diseases can significantly aid and accelerate experimental research discoveries to clinical testing.

Author Spotlight: Studying Neuromuscular Responses and Motor Neuron Plasticity in Neurodegenerative Diseases

In this study, we present an in vivo method for estimating motor unit number and size to quantify rat diaphragm motor unit connectivity. A step-by-step approach to these techniques is described.

Loss of ventilatory muscle function is a consequence of motor neuron injury and neurodegeneration (e.g., cervical spinal cord injury and amyotrophic ...

Neuroscience

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.

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 ...

Real-Time Quantification of Mitochondrial Respiration in Muscle Fibers

Measurement of Mitochondrial Respiration in Human and Mouse Skeletal Muscle Fibers by High-Resolution Respirometry

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle is linked to prevalent metabolic disorders (e.g., diabetes and obesity), muscular dystrophies, and sarcopenia. While there are many techniques to evaluate mitochondrial content and morphology, the hallmark method to assess mitochondrial function is the measurement of mitochondrial oxidative phosphorylation (OXPHOS) by respirometry. Quantification of mitochondrial OXPHOS provides insight into the efficiency of mitochondrial oxidative energy production and cellular bioenergetics. A high-resolution respirometer provides highly sensitive, robust measurements of mitochondrial OXPHOS in permeabilized muscle fibers by measuring real-time changes in mitochondrial oxygen consumption rate. The use of permeabilized muscle fibers, as opposed to isolated mitochondria, preserves mitochondrial networks, maintains mitochondrial membrane integrity, and ultimately allows for more physiologically relevant measurements. This system also allows for the measurement of fuel preference and metabolic flexibility - dynamic aspects of muscle energy metabolism. Here, we provide a comprehensive guide for mitochondrial OXPHOS measurements in human and mouse skeletal muscle fibers using a high-resolution respirometer. Skeletal muscle groups are composed of different fiber types that vary in their mitochondrial fuel preference and bioenergetics. Using a high-resolution respirometer, we describe methods for evaluating both aerobic glycolytic and fatty acid substrates to assess fuel preference and metabolic flexibility in a fiber-type-dependent manner. The protocol is versatile and applicable to both human and rodent muscle fibers. The goal is to enhance the reproducibility and accuracy of mitochondrial function assessments, which will improve our understanding of an organelle important to muscle health.

Author Spotlight: Unveiling Mitochondrial Function and Cellular Metabolic Adaptation in Metabolic Diseases

Here, we describe a comprehensive method for measuring mitochondrial oxidative phosphorylation in fresh permeabilized skeletal muscle fibers from either human or mouse muscle. This method allows for the real-time quantification of mitochondrial respiration and the assessment of fuel preference and metabolic flexibility while preserving existing mitochondrial networks and membrane integrity.

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle ...

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

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