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

Summary

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.

Abstract

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.

Introduction

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 disease^1,2. Although changes in diaphragm function have been well described in amyotrophic lateral sclerosis (ALS) patients and mouse models^3,4,5,6, 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 ALS^7,9,10.

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 procedures^11,12,13. 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 system^14,15. 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.

Protocol

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

1. Put on personal protective equipment (i.e., scrubs, shoe covers, gown, hair net, mask, and surgical gloves).
   NOTE: This surgery requires a sterile field.
2. 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.
3. 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.
4. 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).
5. 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.
6. 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.
7. 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.
8. 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.
9. 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.
10. 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

1. 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.
2. 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.
3. 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.
4. Apply lubricant ophthalmic ointment to prevent the eyes from drying out during surgery.
5. Shave the mouse to expose a surgical site between the ear and shoulder (Figure 1C).
6. 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.
7. 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

1. 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.
2. 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.
   1. 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.
3. 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).
4. 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).
5. 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.
6. 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.
   1. 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.
7. 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.
8. 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.
9. 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.
   1. 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.
10. 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 cyanoacrylate^17,18, an alternative method is to secure the lead cap in place by tying a silk suture knot around it.
11. 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).
12. 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.
13. Repeat steps 3.10 - 3.12 to insert leads into the trapezius muscle, located just anterior to the scalene muscle (Figure 1L and M).
14. 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.
15. 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.
16. 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.
17. 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

1. House animals separately following surgery. Provide healing animals with diet gel and a water bottle.
2. 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.
3. 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

1. Turn on all acquisition equipment, including bias flow.
   NOTE: The flow rate for mice is typically set at 1.0 L/min.
2. 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.
3. Input transmitter calibration as specified by the manufacturer.
4. 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).
5. 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.
6. 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.
7. When acquisition is finished, turn off the transmitter with a magnet and remove the animal from the chamber.
8. 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

1. 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.
2. 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.
3. Identify EMG bouts independently for each muscle.
   1. 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.
   2. 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).
   3. 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.
   4. 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.
   5. Repeat steps 6.3.1 - 6.3.4 for the second muscle.
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.
5. Determine if changes in ventilation are associated with the activation of the recorded muscles.
   1. 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."
   2. Identify the breaths that occur during EMG bout activity and the breaths that occur during EMG baseline activity (Figure 4).
   3. 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."
   4. 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.
   5. 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."
   6. 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.

Results

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)...

Discussion

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
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
Water Blanket	JorVet	JOR784BN