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 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.
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.
Experimental procedures were approved by the Cincinnati Children'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
- Put on personal protective equipment (i.e., scrubs, shoe covers, gown, hair net, mask, and surgical gloves).
NOTE: This surgery requires a sterile field.
- Turn on the incubator (servo-controlled humidifier/infant incubator set to 29 °C) and line it with dry, white towels to allow proper warming for recovery.
- Prior to surgery, sterilize all surgical instruments with ethylene oxide and sterilize the transmitter with an enzymatic detergent and chemical sterilant (or as specified by the manufacturer) before use.
NOTE: Surgical instruments should include #2 laminectomy forceps (standard tips/straight/12 cm) (x4), narrow pattern forceps (serrated/curved/12 cm), tissue-separating scissors (straight/blunt-blunt/11.5 cm), and a scalpel holder and blade. It is recommended to have a separate set of sterilized instruments (two #2 forceps and scissors) reserved exclusively for handling the wires of the transmitter to keep the surgical tools in good condition.
- Sterilize all surfaces within the surgical field with an acceptable disinfectant. Position a stereo dissection microscope, isoflurane anesthetic machine, surgical tools, and clippers in the surgical field (see the Table of Materials).
- To maintain the mouse body temperature while under anesthesia, place a heating pad or water blanket under the sterile towel in front of the stereo dissection microscope.
- Ensure that the transmitter is fully functional prior to use.
NOTE: A small magnet placed within 2 in of the transmitter will turn the device on and off. When the transmitter is on and held close to a radio set at 500 Hz AM frequency, it will emit a continuous, high-pitched buzzing noise. To save battery life, turn the battery off before implanting it in the animal.
- Prepare the electrode leads prior to surgery by trimming the distal leads with the scissors reserved for handling wires so that there is approximately 3 cm of lead (enough to reach the target muscle) (Figure 1A). Alternatively, coil the wires proximal to the device and tie them together with sutures so that there is approximately 3 cm of uncoiled lead.
NOTE: Save the trimmed-off portion of the electrode leads in order to prepare "lead caps" (plastic insulated casing), as described in step 1.9. In step 3, the transmitter leads will be inserted through the muscle, so and the distal, exposed part of the wires must be held in place and insulated using lead caps.
- Use a scalpel to trim off 0.5 cm of the plastic covering without cutting the wire itself. Use the tools reserved for wire handling to stretch the ends of the leads 4 - 5x their original length so that they fit easily inside a 25-gauge needle (Figure 1B-B''). Trim the exposed wire of the leads so that they are 0.5 cm in length.
- Prepare lead caps (plastic tubes to cover the wire ends) prior to surgery. Use a scalpel to trim off 0.25 cm-long tubes from the plastic casing surrounding the segments of electrode leads saved from step 1.7.
NOTE: Four lead caps are required for each mouse that will undergo implantation, but it is best to prepare twice the required amount of caps. Having additional sterile prepared lead caps is helpful in case securing a lead cap is unsuccessful on the first attempt.
- Record the serial number of the inserted transmitter and save the original packaging with the calibrated information. Each transmitter has different frequency calibrations for the EMG recording; enter these into the acquisition software to obtain acceptable EMG recordings.
2. Preparing the Mouse for Surgery
- Choose the desired mouse for implantation (i.e., SOD1(G93A) or control) and weigh the animal.
NOTE: The recommended age and weight for a mouse (male or female) undergoing this surgery is P56 - P120 and ≥ 24 g, respectively.
- Anesthetize the mouse under 3.5% isoflurane with a 2 L/min oxygen flow rate. Perform a toe pinch and a tail pinch to make sure that the mouse is fully anesthetized.
- Once anesthetized, remove the mouse from the drop box and maintain anesthesia via a nose cone by setting the isoflurane level to 1.5% and the oxygen flow rate to 1 L/min. Perform a toe pinch and/or tail pinch to make sure that the anesthesia is maintained.
- Apply lubricant ophthalmic ointment to prevent the eyes from drying out during surgery.
- Shave the mouse to expose a surgical site between the ear and shoulder (Figure 1C).
- 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.
- Alternate swabbing the surgical site, first with disinfectant and then with isopropanol. Repeat 2 more times.
3. Implanting the Telemetry Device to Record Scalene and Trapezius EMG Activity
- Place the animal underneath a dissection microscope, on its side on top of a sterile pad covering a heating pad, and secure the nose cone in place with tape. Note that it is best to implant the right side to reduce ECG signal originating from the heart.
NOTE: Monitor breathing and adjust the isoflurane levels, if necessary, to maintain a regular respiration rate and appropriate surgical plane of anesthesia.
- Pull the forelimb toward the ipsilateral foot along the torso.
NOTE: This position displaces the scapula caudally, providing surgical access to the scalene and trapezius muscles.
- Take the blunt curved forceps, hold back the front paw ipsilateral to the surgical site, and tape the paw in place (Figure 1C). Use strong adhesive surgical tape to make sure that the paw is secure for the duration of the procedure.
- Put on a fresh pair of surgical gloves. Use the scalpel to make an oblique incision, approximately 2 cm-long, between the shoulder and the ear (red line in Figure 1C).
- Using two #2 laminectomy forceps, one 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 (Figure 1D and E).
- 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 (Figure 1F and G).
NOTE: Caution. This area is highly vascularized, and care must be taken to avoid cutting the subclavian artery. Avoid damaging the phrenic nerve and brachial plexus.
- Once the scalene and trapezius muscles have been identified (Figure 1G), make a subcutaneous pocket for the transmitter on the back of the animal, between the scapulas.
- Use the tissue-separating scissors, insert the blunt tips of the scissors just beneath the skin and spread them until a pocket opening that is about 1.25x the width of the transmitter is formed (Figure 1H).
NOTE: The transmitter should be inserted with minimal resistance, but the pocket should not be so big that the transmitter can move on its own. If the pocket is too small, the transmitter could rub against the skin, causing irritation that may prompt the animal to scratch the skin and/or pull the leads out. If the pocket is too large, seromas could form, or the transmitter could migrate to an unfavorable position.
- Use the tissue-separating scissors, insert the blunt tips of the scissors just beneath the skin and spread them until a pocket opening that is about 1.25x the width of the transmitter is formed (Figure 1H).
- Flush with warm, sterile saline and 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 (Figure 1I). Curl any excess length of wire under the device and lay it flat.
- Run the leads from the transmitter to the scalene and trapezius muscles so that the two sets of bipotential leads lie flat and parallel to each other.
- Use the laminectomy forceps to separate the anterior scalene from the surrounding muscles and 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 (Figure 1J and K). Record which colored leads are inserted into which muscle.
- Place a small drop of cyanoacrylate adhesive onto the exposed end of the wire, close to the muscle where the lead is inserted, and quickly slide the lead cap over the wire so that no wire is exposed between the lead cap and the muscle (Figure 2A and B).
NOTE: Although it is an accepted practice to secure EMG leads with cyanoacrylate17,18, an alternative method is to secure the lead cap in place by tying a silk suture knot around it.
- Trim the excess wire distal to the cap and apply a drop of cyanoacrylate adhesive to the end of the lead cap/wire. Give the glue time to polymerize before releasing (Figure 2C and D).
- Follow the same steps (steps 3.10-3.12) to insert the opposite polarity lead parallel to the first in the same muscle, 1 - 2 mm away from the first lead.
- Repeat steps 3.10 - 3.12 to insert leads into the trapezius muscle, located just anterior to the scalene muscle (Figure 1L and M).
- Ensure 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. Ensure that any excess length of lead does not push against the skin, as this could cause irritation that may prompt the animal to scratch or pull the leads out. Reposition the leads, if necessary, to prevent any potential discomfort.
- Gently remove the tape holding down the forelimb. Pull the fat pad back over the muscle and use it to cover the inserted leads. Close the incision with cyanoacrylate adhesive by teasing the skin flaps back together so that the incision lines up. Pinch a portion of the skin flaps together with the curved forceps and apply a small line of cyanoacrylate adhesive along this line.
- Inject 0.1 mL of carprofen subcutaneously to alleviate post-operative pain while the animal is still under anesthesia.
NOTE: Continue to administer 0.1 mL of carprofen once a day for 1 - 2 days post-surgery, and then as needed after that.
- 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. Keep the animal in the incubator for at least 15 min afterward, monitoring its movements and alertness.
4. Postoperative Care
- House animals separately following surgery. Provide healing animals with diet gel and a water bottle.
- Monitor the animal for the first 30 min after surgery. Check on the animal at least every hour for 5 h post-surgery. In the days following surgery, check at least twice daily.
- Watch for necrosis, infection along the incision and within the body cavity containing the implant (i.e., heat, swelling, and redness), and seroma formation.
NOTE: These signs occur within the first week after surgery. A healthy healed animal one month after surgery is shown in Figure 1N. Although EMG recordings can be made immediately after implantation, the animals are given at least one week to heal before recording EMG and plethysmography, as ECG signals can be high immediately after implantation.
5. Acquiring Simultaneous Electromyography and Plethysmography Signals
- Turn on all acquisition equipment, including bias flow.
NOTE: The flow rate for mice is typically set at 1.0 L/min.
- Calibrate the plethysmography chamber(s) using a flow meter.
NOTE: Periodically check the plethysmography chambers to ensure that the seals are not cracked or broken. Coat the rubber seals with a lubricant such as vacuum grease once a week to maintain their good condition.
- Input transmitter calibration as specified by the manufacturer.
- Place the mouse in the plethysmography chamber for at least 1 h to acclimate it prior to recording EMG and plethysmography. Using multiple chambers, it is possible to record from one mouse while the next mouse is acclimating in a second chamber. Do not turn on the transmitter during the acclimation period in order to conserve battery power (Figure 1O).
- Prior to recording (but after calibration), turn on the transmitter by placing a strong magnet within 1 in of the implanted animal; a red light on the front of the receiver will indicate when the transmitter is on.
- Begin acquisition using the pulldown menu labeled "Acquisition" and choose "Start Acquisition." Although the recording duration may vary by experiment, a typical plethysmography and EMG recording lasts 1 - 3 h.
NOTE: The transmitter has an intrinsic sampling rate of 240 Hz. A faster rate of 500 Hz is set in the software to interpolate between points and to provide a smoother waveform. The low pass filter (that serves as an anti-alias filter) and the high pass filter in the implant specify the 1- to 50-Hz bandwidth for this telemetry device. 60-Hz A/C interference does not contribute to excess noise in the EMG signal because the implants are battery powered and the animal shields the implant and leads from electric fields. Plethysmography, EMG, and video are automatically synchronized in real time via acquisition software.
- When acquisition is finished, turn off the transmitter with a magnet and remove the animal from the chamber.
- If starting another recording, clean the chamber, enter in the new transmitter calibrations from the next animal, and begin the second recording. If finished with acquisition for the day, turn off the transmitter, clean the plethysmography chamber, and turn off all acquisition equipment and the bias flow.
Figure 1. Implantation of Telemetry Device to Measure Respiratory Muscle EMG. (A) Telemetry transmitters with two pairs of biopotential leads to measure EMG. Leads can be trimmed to the desired length (bottom) or coiled and tucked underneath the transmitter (top). (B) Transmitter leads. (B') Leads with trimmed-off plastic insulation to expose the wires and to make lead caps (inset). (B'') Leads with wires stretched 4 - 5x their original length. Leads should be trimmed so that they are 0.5 cm long (not shown). (C) Mouse prepared for surgery, with the shaved surgical site and correctly positioned forepaw. The red dotted line indicates the incision site. (D) Superficial muscles located underneath the fat pad and fascia, seen following the initial incision. T = trapezius. S = sternocleidomastoid. P = platysma. Yellow arrow = phrenic nerve. (E) Cartoon diagram of the muscles and phrenic nerve shown in (D). Forceps should be used to spread apart the trapezius and platysma muscles to reach the deeper scalene muscle, shown in (F) and (G). (F) Landmarks used to identify the location of the scalene and the trapezius. This image shows the subclavian artery (white arrow), the phrenic nerve/brachial plexus (black arrow), and the pale sternocleidomastoid muscle (yellow arrow). (G) Cartoon depicting the location of the deeper muscles (i.e., middle scalene, anterior scalene, and SCM), subclavian artery, and phrenic nerve. The posterior scalene is not visible. These can be accessed only when the superficial muscles (in D and E) are spread apart. (H) Making a pocket for the transmitter using the blunt-tip scissors. (I) Inserted transmitter in the subcutaneous pocket, with the parallel-positioned leads emerging from the pocket. (J) Insertion of the 25-gauge needle into the scalene, perpendicular to the muscle fibers, to make a tunnel for the wire lead. (K) Both leads inserted into the scalene muscle. Lead caps are positioned on the end and glued into place. (L) Insertion of the 25-gauge needle into the trapezius, perpendicular to the muscle fibers, to make a tunnel for the wire lead. (M) All four leads inserted into the trapezius and scalene muscles and lying flat prior to the closure of the incision. (N) Fully recovered mouse, with the transmitter positioned subcutaneously on the back. (O) Simultaneously recording plethysmography, muscle EMG activity, and video using a plethysmography chamber (yellow arrow), telemetry receiving pad (red arrow), and camera (black arrow), respectively. A multifunction bias flow is connected to the plethysmography chamber via a plastic tube (blue arrow) to supply oxygen to the mouse. Please click here to view a larger version of this figure.
Figure 2. Securing Lead Caps with Cyanoacrylate Adhesive. (A) Apply a small drop of cyanoacrylate (purple circle) to the exposed wire of the electrode lead (E) wire proximal to the muscle. (B) Quickly slide the prepared lead cap (LC) onto the exposed wire over the cyanoacrylate adhesive so that the lead cap is positioned directly adjacent to the muscle. (C) Trim off a small portion of the distal end of the lead cap and wire so that there is no exposed electrode present that is not insulated with plastic. (D) Apply one small drop of cyanoacrylate adhesive to the end of the lead cap. Remove the trimmed-off distal end of the lead cap from the animal. Please click here to view a larger version of this figure.
6. Analysis of ARM EMG and Plethysmography
- Open the analysis software and review the file of interest (go to "File" and choose "Open review file"). Filter the EMG signals using a 30-Hz high pass filter by right-clicking on the EMG trace, choosing "Analyze Attributes," highlighting the "Advanced Attributes 1" tab, and changing the high pass filter to 30 Hz.
NOTE: This filtering step removes non-discriminate, low-frequency information.
- Locate areas of mouse inactivity by visual inspection based on the lack of movement in the synchronized video file and the lack of large, irregular pressure changes due to movement in the plethysmography trace (red box in Figure 3A); inactivity occurs when the mouse is asleep or awake but still.
- Identify EMG bouts independently for each muscle.
- Rectify and integrate the filtered EMG signal over 30 ms (Figure 4).
NOTE: Because mice breathe at a rate of 3 Hz, each breath is represented by approximately 11 integrated values.
- Determine the baseline EMG amplitude by averaging the rectified and integrated values associated with the EMG signal for a period of 3 s when the mouse is inactive and the plethysmography trace shows eupnea (normal breathing) (Figure 4).
- Identify "bouts" of activity defined by at least 3 consecutive rectified and integrated values that are at least a 50% increase above the baseline EMG signal (determined in step 6.3.2).
NOTE: Three consecutive values represent a 90-ms window, but some bouts will contain more than 3 values above the threshold and will last longer than 90 ms.
- Use the synchronized video and plethysmography trace to exclude bouts that occur during sighs (Figure 3B); sniffing (Figure 3C); or volitional mouse movements, such as head turning or grooming.
- Repeat steps 6.3.1 - 6.3.4 for the second muscle.
- Rectify and integrate the filtered EMG signal over 30 ms (Figure 4).
- Calculate the bout frequency for each muscle. Record a) the beginning time and end time of each inactive period and b) the time each bout occurred using the above criteria. Sum the total inactive time. Divide the total number of bouts by the total min of inactive time over the course of the recording session to calculate the bout frequency.
- Determine if changes in ventilation are associated with the activation of the recorded muscles.
- Select the respiration parameters to be measured (e.g., peak inspiratory flow, tidal volume, minute volume, and breaths per min).
NOTE: All possible selections can be found in the P3 Setup pulldown menu under "Derived Parameters."
- Identify the breaths that occur during EMG bout activity and the breaths that occur during EMG baseline activity (Figure 4).
- Create parser segments spanning plethysmography breaths that are associated with EMG bout activity and create independent parser segments that are associated with baseline EMG activity. Make sure to set the type of analysis to "Parser Seg."
NOTE: This selection is found in the P3 Setup pulldown menu under "Data Reduction Setup."
- Mark the beginning of each parser segment with an event by right-clicking on the plethysmography trace. Specify a bout containing parser segments as "Event 1" in the dropdown menu and specify the baseline parser segments as "Event 2" to distinguish the two classes of segments.
- Under the "Functions" menu, save the "Marks Section" and "Marks Derived Data." Under the Data Parser menu, save the "Parsed Review File" and "Parsed Derived Data."
NOTE: The selected respiration parameters for each individual breath are found in the Marks Derived Data sheet in the tab labeled "Derivations."
- Compare the respiratory parameters for breaths that occur during ARM bouts (marked as Event 1) versus breaths that occur during baseline activity (marked as Event 2) to determine if muscle activity is associated with changes in ventilation.
- Select the respiration parameters to be measured (e.g., peak inspiratory flow, tidal volume, minute volume, and breaths per min).
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.
|Stage||State||Stage Onset||Hindlimb Presentation|
|0||Pre-symptomatic||< P100||No notable differences compared to wildtypes.|
|1||Disease onset||~ P100||Hindlimb collapse when mouse is suspended from tail.|
|2||Paresis||~ P120||Full or partial hindlimb collapse with appearance of tremor.|
|3||Paralysis onset||~ P140||Difficulty walking, toe curling and/or foot dragging.|
|4||Advanced paralysis||~ P150||Minimal joint movement, hindlimb not being used for forward motion.|
|5||Endstage||~ P160||Mouse unable to right itself from side within 30 seconds.|
Table 1. Neurological Scoring of ALS-like Disease Progression in SOD1(G93A) Mice.
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). Please click here to view a larger version of this figure.
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. Please click here to view a larger version of this figure.
|PIF (mL/s)||TV (mL)||MV (mL/min)||Breathing Frequency (Breaths/min)|
|Naive (n=5)||4.4 ± 0.7||0.27 ± 0.04||58 ± 13||223 ± 41|
|Implanted (n=4)||4.1 ± 0.2||0.27 ± 0.11||56 ± 29||201 ± 32|
|Values shown reflect mean ± SD. P-values were calculated with a Student's t-test.|
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 ± SD. P-values were calculated with a Student'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).
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.
The authors have nothing to disclose.
Support for this work was provided by a Cincinnati Children's Hospital Medical Center Trustee Award to S.A.C. and an NIH training grant (T32NS007453) to V.N.J.
|B6.Cg-Tg (SOD1*G93A)1 Gur/J||Jackson Laboratory||4435|
|Plethysmography Chamber||Buxco Respiratory Products/ Data Sciences International||601-1425-001|
|Telemetry Receivers (Model RPC-1)||Data Sciences International||272-6001-001|
|Bias Flow Pump (Model BFL0500)||Data Sciences International||601-2201-001|
|ACQ-7000 USB||Data Sciences International||PNM-P3P-7002XS|
|Dataquest A.R.T. Data Exchange Matrix||Data Sciences International||271-0117-001|
|New Ponemah Analysis System||Data Sciences International||PNM-POST-CFG|
|Ponemah Physiology Platform Acqusition software v5.20||Data Sciences International||PNM-P3P-520|
|Ponemah Unrestrained Whole Breath Plethysmography analysis package v5.20||Data Sciences International||PNM-URP100W|
|Configured Ponemah Software System||Data Sciences International||PNM-P3P-CFG|
|Analysis Module (URP)||Data Sciences International||PNM-URP100W|
|Universal Amplifier||Data Sciences International||13-7715-59|
|Sync Board||Data Sciences International||271-0401-001|
|Sync Cable||Data Sciences International||274-0030-001|
|Transducer-Pressure Buxco||Data Sciences International||600-1114-001|
|Flow Meter||Data Sciences International||600-1260-001|
|Magnet and Radio included in F20-EET Starter Kit||Data Sciences International||276-0400-001|
|Axis P1363 Video Camera||Data Sciences International||275-0201-001|
|Terg-A-Zyme||Fisher Scientific||50-821-785||Enzyme Detergent|
|Actril||Minntech Corporation||78337-000||Chemical Sterilant|
|Stereo Dissecting Microscope (Model MEB126)||Leica||10-450-508|
|Servo-Controlled Humidifier/Infant Incubator||OHMEDA Ohio Care Plus||6600-0506-803|
|TL11M2-F20-EET Transmitters||Data Sciences International||270-0124-001|
|Dumont #2 Laminectomy Forceps - Standard Tips/Straight/12 cm (x2)||Fine Scientific Instruments||11223-20||For handling wires|
|Dumont #2 Laminectomy Forceps - Standard Tips/Straight/12 cm (x2)||Fine Scientific Instruments||11223-20||For surgery|
|Narrow Pattern Forceps- Serrated/Curved/12 cm||Fine Scientific Instruments||17003-12|
|Spring Scissors - Tough Cut/Straight/Sharp/12.5 cm/6 mm Cutting Edge||Fine Scientific Instruments||15124-12|
|Tissue Separating Scissors - Straight/Blunt-Blunt/11.5 cm||Fine Scientific Instruments||14072-10|
|Fine Scissors - Tough Cut/Curved/Sharp-Sharp/9 cm||Fine Scientific Instruments||14058-11||For cutting wires and clipping nails|
|Scalpel Handle #3||World Precision Instruments||500236|
|Scalpel Blade||Fine Scientific Instruments||10010-00||For preparing lead caps|
|Polysorb Braided Absorbable suture||Coviden||D4G1532X||For coiling transmitter leads|
|Gluture||Zoetis Inc.||6606-65-1||Cyanoacrylate adhesive|
|3 mL Syring Slip Tip - Soft||Vitality Medical||118030055|
|25 G Needle (x2)||Becton Dickinson and Co.||305-145|
|Cotton Tipped Applicators||Henry Schein Animal Health||100-9175|
|Andis Easy Cut Hair Clipper Set||Andis||049-06-0271||Electrical Razor sold at Target|
|Isoflurane||Henry Schein Animal Health||29404||Anesthetic|
|Isopropyl Alcohol 70%||Priority Care 1||MS070PC|
|Dermachlor 2% Medical Scrub (chlorohexidine 2%)||Butler Schein||55482|
|Artificial Tears||Henry Schein Animal Health||48272||Lubricant Opthalmic Ointment|
|Vacuum grease||Dow Corning Corporation||1597418|
- Johnson, R. A., Mitchell, G. S. Common mechanisms of compensatory respiratory plasticity in spinal neurological disorders. Respir Physiol Neurobiol. 189, (2), 419-428 (2013).
- Sieck, G. C., Gransee, H. M. Respiratory Muscles: Structure, Function & Regulation. Morgan & Claypool Life Sciences. Lecture #34 (2012).
- Rizzuto, E., Pisu, S., Musaro, A., Del Prete, Z. Measuring Neuromuscular Junction Functionality in the SOD1(G93A) Animal Model of Amyotrophic Lateral Sclerosis. Ann Biomed Eng. 43, (9), 2196-2206 (2015).
- Kennel, P. F., Finiels, F., Revah, F., Mallet, J. Neuromuscular function impairment is not caused by motor neurone loss in FALS mice: an electromyographic study. Neuroreport. 7, (8), 1427-1431 (1996).
- Pinto, S., Alves, P., Pimentel, B., Swash, M., de Carvalho, M. Ultrasound for assessment of diaphragm in ALS. Clin Neurophysiol. 127, (1), 892-897 (2016).
- Stewart, H., Eisen, A., Road, J., Mezei, M., Weber, M. Electromyography of respiratory muscles in amyotrophic lateral sclerosis. J Neurol Sci. 191, (1-2), 67-73 (2001).
- Arnulf, I., et al. Sleep disorders and diaphragmatic function in patients with amyotrophic lateral sclerosis. Am J Respir Crit Care Med. 161, 849-856 (2000).
- Bennett, J. R., et al. Respiratory muscle activity during REM sleep in patients with diaphragm paralysis. Neurology. 62, (1), 134-137 (2004).
- Pinto, S., de Carvalho, M. Motor responses of the sternocleidomastoid muscle in patients with amyotrophic lateral sclerosis. Muscle Nerve. 38, (4), 1312-1317 (2008).
- Romer, S. H., et al. Accessory respiratory muscles enhance ventilation in ALS model mice and are activated by excitatory V2a neurons. Exp Neurol. 287 (Pt. 2, 192-204 (2017).
- Moldovan, M., et al. Nerve excitability changes related to axonal degeneration in amyotrophic lateral sclerosis: Insights from the transgenic SOD1(G127X) mouse model. Exp Neurol. 233, (1), 408-420 (2012).
- Pagliardini, S., Gosgnach, S., Dickson, C. T. Spontaneous sleep-like brain state alternations and breathing characteristics in urethane anesthetized mice. PLoS One. 8, (7), 70411 (2013).
- Nicaise, C., et al. Phrenic motor neuron degeneration compromises phrenic axonal circuitry and diaphragm activity in a unilateral cervical contusion model of spinal cord injury. Exp Neurol. 235, (2), 539-552 (2012).
- Akay, T. Long-term measurement of muscle denervation and locomotor behavior in individual wild-type and ALS model mice. J Neurophysiol. 111, (3), 694-703 (2014).
- Tysseling, V. M., et al. Design and evaluation of a chronic EMG multichannel detection system for long-term recordings of hindlimb muscles in behaving mice. J Electromyogr Kinesiol. 23, (3), 531-539 (2013).
- Weiergraber, M., Henry, M., Hescheler, J., Smyth, N., Schneider, T. Electrocorticographic and deep intracerebral EEG recording in mice using a telemetry system. Brain Res Brain Res Protoc. 14, (3), 154-164 (2005).
- Pilla, R., Landon, C. S., Dean, J. B. A potential early physiological marker for CNS oxygen toxicity: hyperoxic hyperpnea precedes seizure in unanesthetized rats breathing hyperbaric oxygen. J Appl Physiol. 114, (1985), 1009-1020 (1985).
- Morrison, J. L., et al. Role of inhibitory amino acids in control of hypoglossal motor outflow to genioglossus muscle in naturally sleeping rats. J Physiol. 552 (Pt. 3, 975-991 (2003).
- Tscharner, V., Eskofier, B., Federolf, P. Removal of the electrocardiogram signal from surface EMG recordings using non-linearly scaled wavelets). J Electromyogr Kinesiol. 21, (4), 683-688 (2011).
- Hof, A. L. A simple method to remove ECG artifacts from trunk muscle EMG signals. J Electromyogr Kinesiol. 19, (6), e554-e555 (2009).
- Lu, G., et al. Removing ECG noise from surface EMG signals using adaptive filtering. Neurosci Lett. 462, (1), 14-19 (2009).