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.

Experimental procedures were approved by the Cincinnati Children&#39;s Hospital Medical Center Institutional Animal Care and Use Committee and conducted in compliance with the NIH Guide for the care and use of laboratory animals.
1. Preparing for Telemetry Device Implant Surgery
<ol>
	<li>Put on personal protective equipment (i.e., scrubs, shoe covers, gown, hair net, mask, and surgical gloves). 
	NOTE: This surgery requires a sterile field.</li>
	<li>Turn on the incubator (servo-controlled humidif...

<ol>
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F., Finiels, F., Revah, F., Mallet, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuromuscular+function+impairment+is+not+caused+by+motor+neurone+loss+in+FALS+mice:+an+electromyographic+study.">Neuromuscular function impairment is not caused by motor neurone loss in FALS mice: an electromyographic study.</a> Neuroreport. 7 (8), 1427-1431 (1996).</li><li>Pinto, S., Alves, P., Pimentel, B., Swash, M., de Carvalho, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrasound+for+assessment+of+diaphragm+in+ALS.">Ultrasound for assessment of diaphragm in ALS.</a> Clin Neurophysiol. 127 (1), 892-897 (2016).</li><li>Stewart, H., Eisen, A., Road, J., Mezei, M., Weber, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electromyography+of+respiratory+muscles+in+amyotrophic+lateral+sclerosis.">Electromyography of respiratory muscles in amyotrophic lateral sclerosis.</a> J Neurol Sci. 191 (1-2), 67-73 (2001).</li><li>Arnulf, I., 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=Sleep+disorders+and+diaphragmatic+function+in+patients+with+amyotrophic+lateral+sclerosis.">Sleep disorders and diaphragmatic function in patients with amyotrophic lateral sclerosis.</a> Am J Respir Crit Care Med. 161, 849-856 (2000).</li><li>Bennett, J. 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T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spontaneous+sleep-like+brain+state+alternations+and+breathing+characteristics+in+urethane+anesthetized+mice.">Spontaneous sleep-like brain state alternations and breathing characteristics in urethane anesthetized mice.</a> PLoS One. 8 (7), 70411 (2013).</li><li>Nicaise, C., 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=Phrenic+motor+neuron+degeneration+compromises+phrenic+axonal+circuitry+and+diaphragm+activity+in+a+unilateral+cervical+contusion+model+of+spinal+cord+injury.">Phrenic motor neuron degeneration compromises phrenic axonal circuitry and diaphragm activity in a unilateral cervical contusion model of spinal cord injury.</a> Exp Neurol. 235 (2), 539-552 (2012).</li><li>Akay, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+measurement+of+muscle+denervation+and+locomotor+behavior+in+individual+wild-type+and+ALS+model+mice.">Long-term measurement of muscle denervation and locomotor behavior in individual wild-type and ALS model mice.</a> J Neurophysiol. 111 (3), 694-703 (2014).</li><li>Tysseling, V. 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=Design+and+evaluation+of+a+chronic+EMG+multichannel+detection+system+for+long-term+recordings+of+hindlimb+muscles+in+behaving+mice.">Design and evaluation of a chronic EMG multichannel detection system for long-term recordings of hindlimb muscles in behaving mice.</a> J Electromyogr Kinesiol. 23 (3), 531-539 (2013).</li><li>Weiergraber, M., Henry, M., Hescheler, J., Smyth, N., Schneider, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrocorticographic+and+deep+intracerebral+EEG+recording+in+mice+using+a+telemetry+system.">Electrocorticographic and deep intracerebral EEG recording in mice using a telemetry system.</a> Brain Res Brain Res Protoc. 14 (3), 154-164 (2005).</li><li>Pilla, R., Landon, C. S., Dean, J. 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. L., 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=Role+of+inhibitory+amino+acids+in+control+of+hypoglossal+motor+outflow+to+genioglossus+muscle+in+naturally+sleeping+rats.">Role of inhibitory amino acids in control of hypoglossal motor outflow to genioglossus muscle in naturally sleeping rats.</a> J Physiol. 552 (Pt. 3, 975-991 (2003).</li><li>Tscharner, V., Eskofier, B., Federolf, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Removal+of+the+electrocardiogram+signal+from+surface+EMG+recordings+using+non-linearly+scaled+wavelets).">Removal of the electrocardiogram signal from surface EMG recordings using non-linearly scaled wavelets).</a> J Electromyogr Kinesiol. 21 (4), 683-688 (2011).</li><li>Hof, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+method+to+remove+ECG+artifacts+from+trunk+muscle+EMG+signals.">A simple method to remove ECG artifacts from trunk muscle EMG signals.</a> J Electromyogr Kinesiol. 19 (6), e554-e555 (2009).</li><li>Lu, G., 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=Removing+ECG+noise+from+surface+EMG+signals+using+adaptive+filtering.">Removing ECG noise from surface EMG signals using adaptive filtering.</a> Neurosci Lett. 462 (1), 14-19 (2009).</li></ol>

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 per...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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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)...

Watch this Scientific Journal Video about Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease at JoVE.com

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 ...

The overall goal of this procedure is to repeatedly measure respiratory muscle activity and ventilation throughout disease progression. Following the surgical implantation of a telemetry device with electromyography leads inserted into selected muscles. This method can be used to answer key questions about how accessory respiratory muscles compensate for impaired diaphragm function following disease or injury.The main advantage of this technique is that ventilation and respiratory muscle activity can be measured on the same freely behaving mouse throughout disease progression. This method can provide insight into diagonsis and treatment of ALS, but can also be applied to other models of neuromuscular disease or spinal cord injury. Begin by preparing the EMG electroleads.Use scissors reserved for handling wires to trim the distal leads, so that there is approximately three centimeters of lead to reach the target muscle. Then use a scalpel to turn off 0.5 centimeters of the plastic covering without cutting the wire itself. Next use wire-handling tools to stretch the ends of the leads to four or five times their original length.Trim the exposed wire of the leads so that they are 0.5 centimeters in length. Then prepare lead caps to cover the wire ends by trimming off 0.25 centimeter tubes from the previously removed plastic casing. After anesthetizing the mouse, perform a toe pinch and a tail pinch to make sure that the mouse is fully anesthetized by the absence of a reaction.Then transfer the mouse to the surgery set up and place the nose cone to maintain anesthesia during surgery. Apply lubricant ophthalmic ointment to prevent the eyes from drying out during surgery. Clip the toenails ipsilateral to the surgical site to reduce the chance of the mouse pulling out the leads or causing wounds by scratching during the healing process.Shave the mouse to expose a surgical site between the ear and shoulder. Then alternate swabbing the surgical site, first with disinfectant, and then with isopropanol. To begin the implantation, pull the forelimb toward the ipsilateral foot to caudally displace the scapula, providing surgical access to the scalene and trapezius muscles.Use blunt-curved forceps to hold back the front paw ipsilateral to the surgical site, and tape the paw in place. After putting on a fresh pair of surgical gloves, use the scalpel to make a two centimeter oblique incision between the shoulder and the ear. Then with 2 laminectomy forceps in each hand, pull back the fat pad and spread apart the trapezius and platysma muscles to expose the fascia covering the sternocleidomastoid and scalene muscles.Use the pale sternocleidomastoid muscle and phrenic nerve as landmarks to identify the scalene muscles. Note that the phrenic nerve runs parallel to the scalene muscles, while the sternocleidomastoid lies inferior. The scalene muscles run obliquely from the cervical vertebrae to the ribs under the trapezius muscle.Biopotential leads will be inserted into the anterior scalene muscle which can be identified as the muscle that runs adjacent to the phrenic nerve. Next make a pocket for the transmitter by inserting the tips of tissue separating scissors beneath the skin of the mouse and spreading them until a pocket slightly larger than the width of the transmitter is formed. Flush the pocket with warm sterile saline.Then insert the transmitter with the flatter side against the muscle. Position the transmitter so that it lies flat and the wires emerge from the pocket, parallel to each other rather than twisted. Run the leads from the transmitter to the scalene and trapezius muscles, so that the two sets of biopotential leads lie flat and parallel to each other.Next use the laminectomy forceps to separate the anterior scalene from the surrounding muscles. Then insert a 25-gauge needle through the scalene muscle perpendicular to the muscle fibers. Insert one lead into the tip of the needle and then pull the needle out of the muscle leaving behind the lead inserted into the muscle up to the insulation of the wire.Record which color leads are inserted into which muscle. Apply a drop of cyanoacrylate adhesive to a 21-gauge needle to facilitate glue application to the leads. Then place the adhesive onto the exposed end of the wire close to the muscle where the lead is inserted.Quickly slide the lead cap over the wire so that no wire is exposed between the lead cap and the muscle. Trim the excess wire distal to the cap and apply a drop of cyanoacrylate adhesive to the end of the lead cap and wire. Give the glue time to polymerize before releasing.Next insert the opposite polarity lead parallel to the first in the same muscle in the same manner. This lead should be one to two millimeters away from the first lead. Repeat the procedure to insert leads into the trapezius muscle, located just anterior to the scalene muscle as shown here.Next check that the wire leads are fixed in place and that there is just enough slack in the leads for the animal to perform body movements without pulling on the leads. Then gently remove the tape holding down the forelimb. Pull the fat pad back over the muscle and use it to cover the inserted leads.Next tease the skin flaps back together so that the incision lines up. Pinch a portion of the skin flaps together with curved forceps, and apply a small line of cyanoacrylate adhesive along this line. Inject 0.1 milliliters of carprofen subcutaneously to alleviate postoperative pain while the animal is still under anesthesia.Remove the animal from the nosecone and place it in a clean cage in the pre-warmed incubator until the animal is awake and moving around the cage voluntarily. Place the mouse in the plethysmography chamber for at least one hour to acclimate it prior to recording EMG and plethysmography. Prior to recording but after calibration, turn on the transmitter by placing a strong magnet within one inch of the implanted animal.A red light on the front of the receiver will indicate when the transmitter is on. Begin acquisition using the pull-down menu labeled Acquisition and choose Start Acquisition. Although the recording duration may vary by experiment, a typical plethysmography and EMG recording lasts one to three hours.When acquisition is finished, turn off the transmitter with a magnet and return the animal to the home cage. The following are WBP and EMG traces of scalene and trapezius muscles from a presymptomatic SOD-1(G93A)mouse on postnatal day 98. Periods when the animal is at rest are used for analysis.The red box shows EMG traces at rest lacking EMG bouts, characteristic of a presymptomatic mouse. Traces outside the red box show large and irregular peaks in plethysmography traces and muscle activity and EMG traces, typical when an animal is moving. This image shows WBP and EMG traces of the trapezius muscle from a symptomatic SOD-1(G93A)mouse on postnatal day 126, when an EMG bout is present while the mouse is at rest.Once mastered, this technique can be done in 35 minutes if it is performed properly. Following this procedure, implanted animals can be used to assess the effectiveness of pharmaceuticals to enhance accessory respiratory muscle activity in models of disease or injury.

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Research

JoVE Journal

Medicine

12.7K visualizações.  University of Cincinnati. The overall goal of this procedure is to repeatedly measure respiratory muscle activity and ventilation throughout disease progression. Following the surgical implantation of a telemetry device with electromyography leads inserted into selected muscles. This method can be used to answer key questions about how accessory respiratory muscles compensate for impaired diaphragm function following disease or injury.The main advantage of this technique is that ventilation and respiratory musc...