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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

Respiratory complications are the leading cause of death in individuals with cervical spinal cord injury (cSCI). Animal models of cSCI are essential for mechanistic evaluations and pre-clinical studies. Here, we introduce a reproducible method to assess functional recovery of diaphragm muscle (DIAm) activity following unilateral C2 spinal hemisection (C2SH) in rats.

Abstract

Following cSCI, activation of the DIAm can be impacted depending on the extent of the injury. The present manuscript describes a unilateral C2 hemisection (C2SH) model of cSCI that disrupts eupneic ipsilateral diaphragm (iDIAm) electromyographic (EMG) activity during breathing in rats. To evaluate recovery of DIAm motor control, the extent of deficit due to C2SH must first be clearly established. By verifying a complete initial loss of iDIAm EMG during breathing, subsequent recovery can be classified as either absent or present, and the extent of recovery can be estimated using the EMG amplitude. Additionally, by measuring the continued absence of iDIAm EMG activity during breathing after the acute spinal shock period following C2SH, the success of the initial C2SH may be validated. Measuring contralateral diaphragm (cDIAm) EMG activity can provide information about the compensatory effects of C2SH, which also reflects neuroplasticity. Moreover, DIAm EMG recordings from awake animals can provide vital physiological information about the motor control of the DIAm after C2SH. This article describes a method for a rigorous, reproducible, and reliable C2SH model of cSCI in rats, which is an excellent platform for studying respiratory neuroplasticity, compensatory cDIAm activity, and therapeutic strategies and pharmaceuticals.

Introduction

There are more than 300,000 individuals with spinal cord injury (SCI) in the United States, approximately half of whom have cervical injuries1. These injuries result in significant loss of well-being and place a financial strain on individuals, their families, and the healthcare system. Fortunately, the majority of SCIs are incomplete—providing the potential for strengthening of spared pathways1. This neuroplasticity may allow recovery of at least some function, including DIAm activity, which is important for ventilatory and non-ventilatory behaviors. Thus, promoting neuroplasticity is a promising avenue of research to help individuals with SCI2.

Rodent models of SCI have the potential to contribute substantially to the discovery of treatments to improve human health. One of the classic models of SCI used to study neuroplasticity is a unilateral transection (hemisection) of the spinal cord at C2 (C2SH), which leaves the contralateral side intact3,4,5,6,7,8,9,10,11,12,13. The effect of C2SH on phrenic output and the importance of spared contralateral pathways was first revealed over a hundred years ago by Porter12, whose seminal article laid the foundation for modern-day studies of respiratory neuroplasticity. The C2SH model interrupts descending inputs from the rostral ventral respiratory group (rVRG) in the medulla, which contains premotor neurons responsible for transmitting the output of respiratory rhythm generation14. These rVRG premotor neurons also transmit excitatory neural drive to phrenic motor neurons (Figure 1). Several investigators have taken different approaches to the C2SH model10,11,15,16, which may partly explain some of the variability in recovery across studies. Briefly, approaches vary in terms of sparing the dorsal funiculi, performing a complete hemisection, or performing a lateral partial transection that does not completely interrupt descending inputs from the ipsilateral rVRG. Generally, C2SH models are particularly useful for studying respiratory neuroplasticity due to the rates of spontaneous recovery of eupneic iDIAm electromyographic (EMG) activity over time, which can be improved by several factors, including neurotrophic signaling17,18,19,20,21. However, an initial loss of function—defined as the silencing of eupneic iDIAm EMG activity—must be first established before recovery can be clearly classified. This validation of inactivity at the time of C2SH is not done in several studies3,4,6,7,11,22,23.

Histological assessments of the excised spinal cord only provide evidence of damage to the appropriate location of ipsilateral excitatory bulbospinal pathways innervating phrenic motor neurons in the spinal cord, but histology does not substitute for physiological evidence (e.g., DIAm EMG). Furthermore, histological assessments are performed in ex vivo at terminal time points (often several weeks to months post-injury) and thus do not provide "real-time" information. Some investigators have noted that the magnitude of the lesion relates to the amount of functional deficit or lack thereof5,24,25,26. It is important to note that the validity of such claims is likely highly dependent on how "function" is classified (i.e., what the functional tasks are and how they are quantified), and the variability across studies highlights the difficulty of producing functionally identical lesions across animals. Indeed, investigators have emphasized that the relationship between the extent of injury and limb muscle locomotor function (quantified by the Basso, Beattie, and Bresnahan (BBB) score24) is not linear27,28. In previous studies, we have found no relationship between the extent of the C2SH and the extent of recovery of eupneic iDIAm EMG activity post-injury10,29,30,31, although other investigators have reported a relationship between ventilatory function and the extent of white matter sparing5. Thus, in the case of the C2SH model, an approach for functional validation of iDIAm inactivity at the time of the surgery and preferably early in the time course of chronic spinal cord injury experiments is both beneficial and necessary.

The present article underscores the use of DIAm EMG for real-time confirmation of the initial loss of DIAm EMG during breathing after the C2SH as well as subsequent confirmatory assessments at 3 days (Day 3) after the injury18,21,31,32,33. In earlier work with the C2SH model, repeated laparotomies were performed to record DIAm EMG10,13,30,34. However, more recent work has used chronic EMG electrodes, which allow the recording of EMG in anesthetized and awake rats. Additionally, chronic electrodes reduce the risk of pneumothorax and don't require repeated laparotomies, which can cause inhibition of the DIAm35,36. Although versions of the C2SH model have been used by many investigators, confirmation of the silencing of iDIAm activity was not made at the time of surgery3,4,6,7,11,22,23. Without such a confirmation of inactivity, it is difficult to know what portion of subsequent recovery to attribute to the neuroplasticity of ipsilateral versus contralateral pathways, which may have differential impacts. This is an important consideration because the inspiratory neural drive from the rVRG to phrenic motoneurons is primarily ipsilateral, with a loss of about 50% of excitatory glutamatergic inputs to phrenic motor neurons after C2SH33. However, there are remaining inspiratory excitatory inputs from the contralateral rVRG that decussate below the site of the lesion to innervate ipsilateral phrenic motor neurons and can be strengthened via neuroplasticity to promote functional recovery. By removing the predominant ipsilateral excitatory input to phrenic motor neurons, eupneic iDIAm EMG activity is lost (at least under anesthesia), while the activity of the cDIAm continues and is even enhanced. The loss of iDIAm EMG activity during breathing is thus a measure of a successful C2SH (Figure 2).

Some level of iDIAm EMG activity is present as early as 1-4 days following C2SH in awake animals23,37. Additionally, in decerebrate animals, iDIAm activity is present within minutes to hours after upper cervical hemisection and is suppressed by anesthesia38. Additionally, the success of the C2SH is validated by confirming the absence of iDIAm EMG activity during breathing (eupnea) in anesthetized rats on Day 3 post-injury. Confocal imaging studies confirmed the loss of glutamatergic synaptic inputs on phrenic motor neurons during this initial stage of injury37. At Day 3 post-injury, if there is any residual eupneic iDIAm EMG activity, this is interpreted as evidence of incomplete removal of ipsilateral descending inspiratory drive from the rVRG. The present article is divided into three sections: (1) chronic DIAm EMG recordings, (2) C2SH, and (3) EMG data acquisition in awake and anesthetized animals. This protocol describes a rigorous, reproducible, and reliable C2SH model of cSCI in rats, which is an excellent platform for studying respiratory neuroplasticity, compensatory cDIAm activity, and therapeutic strategies and pharmaceuticals.

Protocol

This protocol was approved by the Mayo Clinic Institutional Animal Care and Use Committee (Protocol Number: A00003105-17-R23). The animals in the present study were a mix of male and female Sprague-Dawley rats approximately 3 months old and weighing between 200 g to 350 g. The details of the reagents and the equipment used in the study are listed in the Table of Materials.

1. Electrode implantation

  1. Preparing the electrodes
    1. Ensure that the mentioned equipment is available: (1) Multistranded, stainless steel wire, (2) scalpel with #11 blade, (3) scissors, 4) dissection microscope, (5) ruler, (6) forceps, (7) 25 g needles, (8) 2 pliers (at least one with a crimping zone).
    2. Details of the process for making electrodes are shown in Figure 3. Repeat this process at least twice for paired differential recording electrodes.
    3. First, measure and cut 18 cm of the stainless steel wire.
    4. Tie a basic overhand knot at 4 cm from one end of the wire, as shown in Figure 3; this will serve as an anchor when the short side of the wire is implanted and sutured into the DIAm.
    5. Using a scalpel and a surgical dissection scope, carefully strip 2 mm of insulation immediately adjacent to the knot on the shorter side of the wire. Exercise great care not to cut strands of the multistranded wire.
      NOTE: This uninsulated portion serves as the conducting surface of the electrode. Generally, signal quality begins to degrade if three or more strands are inadvertently cut, and if so, the electrode is replaced.
    6. Next, strip ~5 mm from the end of the short side of the wire, which is then inserted into the lumen of a 25 G needle to make a suture thread-like tool that will be used to implant the electrode into the DIAm.
      NOTE: Higher gauge needles (e.g., up to 28 G) may also be used depending on personal preference. Due to the thinner diameter of higher-gauge needles, the risk of pneumothorax may potentially be reduced. However, it may also be more difficult to insert the multistranded wires into the smaller lumen of the needle.
    7. To accomplish this, remove a 25 G needle from its packaging, and using both pliers, carefully bend the needle at ~1 cm from the sharp beveled end from side to side until it breaks.
      NOTE: This process must not be rushed; otherwise, the shearing forces can cause the lumen of the needle to shut, requiring the process to be repeated.
    8. Insert the stripped end of the wire into the lumen of the broken-off 1 cm needle, ensuring that the needle rests immediately adjacent to the remaining insulated portion of the wire.
    9. Ensuring that there is no uninsulated portion between the needle and the remaining shorter side of the wire, crimp the needle to attach it to the wire.
      NOTE: Crimping at this point ensures that the microscopic rough edges of the broken-off needle are flattened.
    10. Next, use the two pliers to carefully bend the attached needle to give it a curvature similar to that of a standard surgical suturing needle.
    11. Finally, strip away ~5 mm from the end of the long side of the wire (Figure 3). After the electrodes are sutured into the DIAm, the long side of the wire will be tunneled for external access to allow for DIAm EMG recordings.
    12. Repeat the process to make a second electrode for differential recordings.
  2. Diaphragm EMG electrode placement
    1. Prior to surgery, sterilize or autoclave all surgical equipment, including (1) small animal hair clippers, (2) scalpel with #11 blade, (3) surgical scissors, (4) forceps, and (5) micro-needle holder.
    2. Acquire a clean cage and provide food, water, bedding, and environmental enrichment. Set aside on a heating pad. This cage will be used to house the animal after surgery.
    3. Sterilize the surgical field with appropriate disinfectant and put on personal protective equipment (scrubs, caps, mask, and surgical gloves). Place the previously prepared electrodes in a pool of ethylene oxide.
    4. Place a heating pad in the surgical area to maintain the rat's body temperature during anesthesia and surgery. Cover the heating pad with a sterilized surgical towel.
    5. Weigh the rat and calculate the appropriate anesthetic dosage based on institutional animal care and use guidelines.
    6. Anesthetize the rat with an intraperitoneal injection of ketamine (80 mg/kg) and xylazine (10 mg/kg) or other appropriate anesthetic (following institutionally approved protocols). Inject extended-release buprenorphine (1 mg/kg) and carprofen (5 mg/kg) preoperatively. Proceed once the animal no longer responds to toe pinch.
    7. Use clippers to shave the rat from the xiphoid process to approximately halfway to the hind limbs. Also, shave the back of the rat where electrodes will be externalized beyond the reach of the animal (usually around the shoulder blades). Clean off excess hair with a piece of gauze.
    8. Prepare the skin for incision by cleaning it with sterile gauze dipped in chlorohexidine gluconate (4% weight by volume). Clean off with 70%isopropyl alcohol. Alternate with chlorhexidine gluconate and isopropyl alcohol three times.
    9. Place the animal in a supine position on the sterilized heating pad. Starting from the xiphoid process, use the scalpel and a sterile #11 blade to make a 4-5 cm rostrocaudal incision. This should expose the underlying abdominal muscle layers.
    10. Make another incision (3-4 cm) along the midline in the muscle layers to expose the abdominal contents. Take care not to cut through into the abdominal organs.
    11. Ensure that the abdominal (i.e., inferior) surface of the DIAm is visible, although the animal's right side will be obscured by the liver.
    12. Using a scalpel handle or other blunt instrument, push down the liver and other organs to expose the abdominal side of the DIAm.
    13. Prepare the electrode for use by removing it from the pool of isopropyl alcohol and washing it with sterile saline. Then, using a micro-needle holder, orient one of the previously prepared electrodes perpendicular to muscle fibers in the mid-costal region of the DIAm. Muscle fibers spread out radially across the DIAm.
      NOTE: The DIAm is thin (~2 mm) and must not be punctured. Rather, the goal is to embed the uninsulated portion of the electrode wire (Figure 3) within the DIAm.
    14. Use the surgical microscope to carefully insert the electrode suture wire (needle end) into the DIAm. Allow the curvature of the needle to guide the electrode into and then out of the abdominal surface of the DIAm, ensuring that the entire uninsulated portion of the wire will be embedded fully in the muscle.
    15. Carefully pull the needle through until the knot in the wire anchors itself in the DIAm. Ensure that the uninsulated portion (2 mm) of the electrode wire is completely within the DIAm, with a knot at the point where the needle first entered the DIAm.
    16. Next, cut off the needle from the electrode wire and dispose of it in a sharps container.
    17. Using two forceps, tie a knot at the point where the suture (3-0) needle exited the DIAm. To do this, tie a loop in the short side of the wire, slide the loop down to the abdominal surface of the DIAm, and carefully tighten the loop into a knot. The electrode (uninsulated portion) should now be anchored at both its entrance and exit point.
    18. Cut off the excess part of the shorter side of the wire.
    19. Follow steps 1.2.14-1.2.18 to suture in another electrode on the same side of the DIAm, ~3 mm away from the first.
      NOTE: The secured electrode wire is safe to leave in the rats for chronic EMG measurements for multiple weeks.
    20. Perform electrode pair implantation in the opposite hemidiaphragm in the same manner.
    21. Next, ensure that the longer side of the electrode wires are all outside of the abdominal cavity and begin to suture (3-0) the muscle layers to close the abdomen. For each pair of electrodes, make a loop and anchor it.
      NOTE: Ensure that 2-3 cm of wire is left within the abdominal cavity to allow the rat to stretch without causing tension on the DIAm.
    22. Clean off excess blood with saline and then suture (3-0) the muscle layer in a continuous pattern to close the peritoneum.
    23. Sterilize the point of exit in the medial dorsum for the wire alternating with chloroxidine gluconate and alcohol three times. Tunnel a 16 G catheter from the medial dorsal skin of the rat through to the open abdominal skin and pull a pair of electrode wires through to the back of the rat.
    24. Repeat the process with a second catheter on the other side and pull the second pair of electrode wires through.
    25. Suture the abdominal skin with a 3-0 suture in an interrupted suture pattern.
    26. Next, for each pair, tie the electrode wires into a knot and suture them in place.
    27. Provide the animal with a subcutaneous saline injection (~1 ml per 50 g of animal mass). Put the animal in a clean cage over a heating pad to recover.
      NOTE: Laparotomy can cause inhibition of DIAm activity35,36. Allow at least 72 h before quantitative analysis of any EMG measurements.

2. Cervical spinal hemisection

  1. Prepare sterile surgical area as before.
  2. Prior to surgery, autoclave or sterilize all surgical equipment. Equipment required: (1) small animal hair clippers, (2) scalpel with #11 blade, (3) surgical scissors, (4) forceps, (5) rongeurs, (6) retractors, (7) angled dissecting knife.
  3. Approximately 72 h after the DIAm EMG electrode implantation, weigh the rat and calculate the appropriate anesthetic dosage based on institutional animal care and use guidelines.
  4. Prior to anesthetizing the rat, place the rat in a Bowman-style cage, and record bilateral DIAm EMG as outlined in step 3.2.
  5. Anesthetize the rat with an intraperitoneal injection of ketamine (80 mg/kg) and xylazine (10 mg/kg) or other appropriate anesthetic as per the local Institutional Animal Care and Use Committee protocol. Preoperatively inject extended-release buprenorphine (1 mg/kg) subcutaneously and carprofen (5 mg/kg) intraperitoneally, or other appropriate analgesics as per the local institutional protocol.
  6. Proceed once the animal no longer responds to toe pinch. Check the depth of anesthesia with a toe pinch approximately every 15 min.
  7. Prior to beginning surgery, place the rat in a prone position and record EMG in the anesthetized rat as outlined in step 3.3.
    NOTE: Ensure that all recordings are performed with the rat in the same position.
  8. Shave the hair from the neck at about the ear level and down to the scapulae and remove it with wet gauze.
  9. Clean the skin with chlorohexidine gluconate (4% weight by volume) and isopropyl alcohol (70%), alternating three times. Cover the animal with sterile surgical covers (except for the upper dorsum).
  10. Using a scalpel, make a 4 cm rostro-caudal incision. Retract the skin and cut the acromiotrapezius muscle. Then dissect the rhomboid muscle to expose the spinalis muscles.
    NOTE: The acromiotrapezius is the middle trapezius muscle, and its muscle fibers run approximately parallel to the spine. The rhomboid muscle is exposed in the plane below the acromiotrapezius and its muscle fibers run diagonally.
  11. Remove the spinalis muscles from C1 to C3.
    NOTE: The spinalis muscles run immediately adjacent to the spine.
  12. Carefully perform a laminectomy at C2 using a rongeur while looking through a surgical microscope to ensure that no major arteries or nerves are damaged.
  13. Cut and retract the dura mater at C2.
  14. Connect exposed electrode wires from the rat's back to the amplifier while performing the spinal hemisection. Refer to step 3 for more details.
  15. While monitoring DIAm EMG, insert the angled dissecting knife just below the point where the dorsal root enters the spinal cord and section all the way to the midpoint of the ventral surface in the manner shown in Figure 2A.
    NOTE: If using an electronic heat pad, it may be necessary to turn it off while monitoring EMG to avoid noise.
  16. Within seconds after this sectioning, ensure that the eupneic DIAm EMG activity ipsilateral to the injury ceases and a compensatory increase in the cDIAm EMG activity is evident (Figure 2C).
  17. If eupneic iDIAm EMG activity persists, perform another cut with the dissecting knife, as highlighted in Figure 2C. However, do not cut across the midline; cutting to the contralateral side will most likely lead to the death of the animal.
  18. Suture the muscles and then the skin with sterile sutures (3-0).
  19. To maintain hydration, subcutaneously inject 1 mL of saline per 50 g of animal mass and place the animal in a clean cage with a heating pad for recovery. Monitor the animal as it recovers from the surgery.
  20. Approximately 72 h later (Day 3 Post-C2SH), record the EMG activity in awake rats as per the protocol step 3.2. Be careful not to damage the stitches on the dorsum of the rat.
  21. Next, anesthetize the rat once again (following institutionally approved protocols), using a lighter dose of anesthetic than would be required for a surgical plane (between ⅓ to ½ of a normal dose). Additional doses may be given as long as the rats display an intact corneal reflex and respond to toe pinch.
  22. Monitor DIAm EMG activity following protocol step 3.
    NOTE: Place the rat in the same position as the one used for the initial assessment of iDIAm EMG activity.
  23. If eupneic iDIAm EMG activity (i.e., activity in phase with inspiratory cDIAm EMG activity) is absent on Day 3, conclude that the initial C2SH was successful and include the same animal for further analyses.

3. Data acquisition and analysis

  1. General settings and data extraction
    1. Set band-pass filters of the preamplifier to 100 Hz (high-pass) and 1000 Hz (low-pass).
      NOTE: These filter settings eliminate movement artifacts, and markedly reduce 60 Hz noise and ECG contamination (which has peak power at ~75 Hz) while retaining the vast majority of the frequency content of the intramuscular DIAm EMG signal. Based on the Nyquist theory, digitize with a sampling frequency of 2000 Hz to avoid aliasing and allow adequate resolution to note motor unit activity in the compound EMG. If performing specialized frequency content analyses of the sub-100 Hz frequency range, it is important to set the preamplifier filter settings to ensure that the frequencies of interest are not filtered.
    2. Monitor two channels (left and right hemidiaphragm EMG) in visualization software on a computer and save the data in an appropriate format for further analyses.
    3. Collect at least 1-2 min of eupneic recordings to have enough data for analyses.
    4. Ensure that the data is collected at three or more time points:
      1. Prior to C2SH, establish a baseline eupneic recording in awake and anesthetized conditions.
      2. During the C2SH surgery to establish silencing of eupneic iDIAm EMG activity in anesthetized conditions.
      3. At Day 3 post-C2SH to confirm that the absence of eupneic iDIAm EMG activity persists and establish an initial point of the extent of the deficit in awake animals that is not as influenced by spinal shock2,39 or administration of respiratory depressants like buprenorphine.
    5. Analyze the DIAm EMG recordings using any software that allows calculations to be performed on digitized signals (e.g., Matlab, Python, R). The basic process is as follows:
      1. Load the raw EMG files. Depending on the filtering settings, it may be necessary to account for a DC offset in the signal.
        NOTE: An easy fix is to subtract the mean of the raw EMG signal from itself.
      2. Calculate the root-mean-square of the EMG signal with a moving window of 50 ms. This can be done by moving a 50 ms calculation window forward by one sample point at a time until the end of the signal is reached.
        NOTE: Analyses of individual DIAm EMG events provide the most statistical power and allow clustering of eupneic and non-eupneic activity. Thus, detection of the onset and offset of individual breaths in the DIAm EMG is paramount, as highlighted in a previous report40.
    6. Perform the filtering of eupneic activity by visualizing the histograms of the instantaneous respiratory rate and burst duration, as shown in a previous report41.
  2. Awake recordings.
    1. Perform awake recordings by placing rats in a Bowman-style rodent cage. A cylindrical Bowman-style cage can accommodate adult rats up to 750 g.
    2. Unscrew metal rods on one side of the Bowman-style cage and gently coax the rat inside. Screw metal rods back on, trapping the rat in the cylindrical cage.
      NOTE: Acclimate the rats to the cage for at least 30 min before proceeding with recording. If possible, it is best to acclimate rats to the Bowman-style cage for multiple days before any measurements. It is good practice to provide treats while the rats are in the cage during this acclimatization phase.
    3. To minimize external visual stimuli, use a paper towel or similar object to make a tent around the front of the cage where the rat's head is. In addition, minimize loud noises and ensure the physical comfort of the rat at all points.
      NOTE: Obvious signs of discomfort include fidgeting, rapid breathing, and struggling against the cage. If any of these signs are observed, it may be necessary to return the rat to the home cage and attempt recordings when the rat is calmer.
    4. For smaller rats (<300 g), insert filler materials (e.g., paper towel, cotton balls) into extra spaces in the cage to discourage the rat from turning 180 degrees.
    5. After the rat is acclimated, carefully connect the wires exiting the rat dorsum to a preamplifier, which feeds the analog signals into an analog-to-digital converter, and follow the procedures outlined in step 3.1.
  3. Anesthetized recordings
    1. Place rats in the same position each time, preferably prone to more closely mimic the awake rat posture.
    2. Connect the wires exiting the rat dorsum to a preamplifier, which feeds the analog signals into an analog-to-digital converter, and follow the procedures outlined in step 3.1.
  4. Animal recovery
    1. Place rats inside an empty, clean rat cage with no bedding for recovery.
    2. Place the cage on a heating pad.
    3. Monitor the rat at regular intervals (<15 min) for the first 3 h or until the rat begins to move. Monitor at less regular intervals (30 min) until the animal is awake again.
    4. After the rat is ambulatory, place them in a clean cage with bedding, access to food and water, and environmental enrichment.
    5. Continue to monitor the rat at least once daily throughout the time course of the experiment. Pay special attention to changes in weight/ability to feed as well as the integrity of the incision sites and sutures. Re-dose buprenorphine ER and carprofen or appropriate analgesics for 72 h after surgery.

Results

The approach presented in this article minimizes inter-operator variability by setting clear criteria for evaluating DIAm EMG in a rat model of C2SH. First, the cessation of eupneic iDIAm EMG activity immediately after C2SH must be observed, as shown in Figure 2. If not, a secondary transection can be performed until eupneic iDIAm activity disappears. Second, on Day 3 post-C2SH, the continued absence of eupneic iDIAm EMG must be verified while animals are ane...

Discussion

C2 spinal hemisection
The procedure described in this article emphasizes assessments of DIAm EMG activity that serve as a validation of a C2 spinal lesion that transects the lateral and ventral funiculi while sparing the dorsal funiculi (Figure 2A). The proposed surgical approach has two major benefits. First, it spares the dorsal funiculi, which preserves ambulatory function in rats, while still severing ipsilateral inputs to phrenic motor neuron...

Disclosures

The authors have no conflicts of interest to declare.

Acknowledgements

The authors acknowledge the NIH funding source (NIH R01HL146114).

Materials

NameCompanyCatalog NumberComments
25 G NeedleCardinal Health1188825100Covidien Monoject Hypdermic Standard Needles: 25 G x 1" (0.508 mm x 2.5 cm) A
3-0 Vicryl Violet BraidedEthiconJ774D3-0 Suture
Adson-Brown ForcepsFine Science Tools11627-12Tip Shape: Straight, Tips: Shark Teeth, Tip Width: 1.4mm, Tip Dimensions: 2 x 1.4 m, Alloy / Material: Stainless Steel, Length: 12 cm
Bowman Style CageBraintree ScientificPOR-530Weight range: 250 up to 750 g; Maximum length: 9" (228 mm); Basic unit is constructed of .5" (123 mm) jeweled acrylic.
Castroviejo Needle HolderFine Science Tools12565-14Tip Shape: Straight, Tip Width: 1.5 mm, Clamping Length: 10 mm, Lock: Yes, Scissors: No, Alloy / Material: Stainless Steel, Length: 14 cm, Serrated:
Yes, Feature: Tungsten Carbide
Clip Lead 1m TP ShieldedBiopac Systems, IncLEAD110SShielded lead wires for EMG
Data Acquisition SoftwareLabChartLabChart 7.3.8Data recording, visualization, and analysis software for multi-channel recordings and real-time assessments
Data Analysis Software - Matlab 2023bMathworks, Inc.Version 23.2General purpose programming language for post hoc analysis
Dissecting KnifeFine Science Tools10056-12Cutting Edge: 4 mm, Thickness: 0.5 mm, Alloy / Material: Stainless Steel, Length: 12.5 cm, Blade Shape: Angled 30°
Dumont #3 ForcepsFine Science Tools11293-00Style: #3, Tip Shape: Straight, Tips: Standard, Tip Dimensions: 0.17 x 0.1 mm, Length: 12 cm, Alloy / Material: Dumostar
Electromyogram AmplifierBiopac Systems, IncEMG100CEMG amplifier
Friedman RongeurFine Science Tools16000-14Tip Shape: Curved, Cup Size: 2.5mm, Alloy / Material: Stainless Steel, Length: 13cm, Joint Action: Single
Friedman-Pearson RongeursFine Science Tools16021-14Alloy / Material: Stainless Steel, Length: 14cm, Joint Action: Single, Cup Size: 1mm, Tip Shape: Curved
Isolated Power Supply ModuleBiopac Systems, IncIPS100COperates 100-series amplifier modules indepdent of the Biopac Systems, Inc.'s MP series Data Acquisition System
Kelly HemostatsFine Science Tools13019-14Tips: Serrated, Tip Width: 1.5mm, Clamping Length: 22mm, Alloy / Material: Stainless Steel, Length: 14cm, Tip Shape: Curved
Knife CuretteV. MuellerVM101-4414Tip: Sharp, Tip Diameter: 2 mm
Micro Dissecting ScissorsBiomedical Research Instruments, Inc.11-2420Length: 4", Angle: Straight, Blade Length: 23 mm
Multistranded stainless steel wireCooner Wire, Inc.AS 631AWG 40; Overall diameter: 0.011 mm (with insulation), 0.008 mm (without insulation).
PowerLab 8/35ADInstrumentsPL3508Data acquisition system
Scalpel Blade #11Fine Science Tools10011-00Blade Shape: Angled, Cutting Edge: 20 mm, Thickness: 0.4 mm, Alloy / Material: Carbon Steel
Scalpel Handle #3Fine Science Tools10003-12Alloy / Material: Stainless Steel, Length: 12 cm
Sprague Dawley RatInotivOrder code: 002Sprague Dawley outbred rats (female and male)
Surgical MicroscopeOlympusSZ61Surgical microscope 
Suture Cutting ScissorsGeorge Tiemann & Co.110-1250SBAlloy / Material: Stainless Steel, Tip Shape: Straight, Tips: Sharp/Blunt, Length: 4.5"
Vannas Spring ScissorsFine Science Tools15000-08Tips: Sharp, Cutting Edge: 2.5 mm, Tip Diameter: 0.05 mm, Length: 8 cm, Alloy / Material: Stainless Steel, Serrated: No, Tip Shape: Straight
Weitlaner RetractorCodman50-5647Prongs: 2 x 3 Blunt, Length: 4.5"

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Functional RecoveryEupneic Diaphragm ActivityUnilateral Cervical Spinal Cord HemisectionC2 Hemisection ModelDIAm Motor ControlEMG ActivitySpinal Cord InjuryNeuroplasticityCompensatory EffectsRespiratory NeuroplasticityCDIAm ActivityTherapeutic Strategies

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