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
2. Preparing the Mouse for Surgery
3. Implanting the Telemetry Device to Record Scalene and Trapezius EMG Activity
4. Postoperative Care
5. Acquiring Simultaneous Electromyography and Plethysmography Signals
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
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 |
P-value | 0.439 | 1.000 | 0.893 | 0.410 |
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/12cm (x2) | Fine Scientific Instruments | 11223-20 | For handling wires |
Dumont #2 Laminectomy Forceps – Standard Tips/Straight/12cm (x2) | Fine Scientific Instruments | 11223-20 | For surgery |
Narrow Pattern Forceps- Serrated/Curved/12cm | Fine Scientific Instruments | 17003-12 | |
Spring Scissors – Tough Cut/Straight/Sharp/12.5cm/6mm Cutting Edge | Fine Scientific Instruments | 15124-12 | |
Tissue Separating Scissors – Straight/Blunt-Blunt/11.5cm | 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 | |
25G 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 | |
Water Blanket | JorVet | JOR784BN |