Method Article

Electrophysiological Methods to Assess Peripheral Pain Block in an Anesthetized Rat

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DOI:

10.3791/68740

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November 21st, 2025

 ,  ,  ,  ,  ,  , 
1Department of Physical Medicine and Rehabiliation, The MetroHealth System, 2Department of Physical Medicine and Rehabilitation, Case Western Reserve University School of Medicine

In This Article

Summary

Here, we present a method for assessing the activation and block of sensory axons in peripheral nerves in acute, anesthetized, in vivo rat experiments using two types of spinal electrophysiological measurements that can discriminate the activity of nerve fiber subtypes: Local Field Potentials and Wide Dynamic Range single neuron recordings.

Abstract

In vivo electrophysiological recordings from the lumbar region of the rat spine can be used to study the activation of sensory neural signals in the hindlimbs, as well as the efficacy of therapeutic interventions that interrupt ascending pain signals.

Here, we describe the surgical procedures (laminectomy and sciatic nerve exposure), data collection, and analysis of two complementary electrophysiological signals: local field potential recordings and wide dynamic range single neuron recordings. Both differentiate activation of nerve fiber subclasses based on conduction velocity relative to an electrical stimulation pulse.

Local field potentials (LFP) can differentiate between A α/β and C nerve fibers but cannot detect Aδ activity. They provide information about the global activation of the whole sciatic. Following electrical direct stimulation of the sciatic nerve using a J-cuff style bipolar electrode, area under the curve measurements of the two troughs corresponding to each component are calculated.

Wide dynamic range (WDR) single neuron recordings assess the activity of a subset of sciatic activation, and can differentiate Aα/β, Aδ, and C fiber types. Electrical stimulation can be applied directly to the sciatic as with LFP, or via needle electrodes placed in the plantar surface of the hindpaw. This higher-frequency signal yields high signal-to-noise action potential spikes. Spike counts within predefined latency windows reflect the level of activity of the various fiber types.

To date, these techniques have been used to assess the temporal properties of three different pain block modalities: Direct current electrical nerve block using a custom-made Carbon Separated Nerve Interface Electrode (CSINE), Kilohertz Frequency Alternating Current, and photobiomodulation using an 830 nm laser. This surgical preparation has been used for nerve block interventions for up to 5 h.

Introduction

Directly blocking pain signals in peripheral nerves prior to reaching the spinal cord has potential for several pain therapy applications, such as knee osteoarthritis and post-amputation phantom limb syndrome1,2,3,4. In general, such pain therapy interventions would reduce/eliminate the systemic side effects noted in current pharmaceutical anesthetics and analgesics1,2.

Two types of complementary electrophysiological measurements can be recorded from the lumbar spinal cord to assess peripheral sensory activation: local field potentials (LFP)3,4 and wide dynamic range (WDR)5,6,7 single neuron recordings. LFP can be recorded from the spinal dorsal columns. They are used to measure the activation of the whole sciatic nerve; the LFP represents the responses of many cells, including excitatory postsynaptic potentials in the dendrites and action potentials recorded from the soma and axon terminals. They have been used to study spinal cord neuroplasticity3 and the pain-mitigating effects of electrical spinal cord stimulation8. From the LFP signals, one can measure the amplitude of large, myelinated Aα/β as well as the amplitude of unmyelinated C-fibers3. WDR neurons are present in the dorsal horn of the L4 and L5 spinal cord. These neurons are multi-receptive; they receive, and faithfully transmit, input from all subclasses of sensory neuron: nociceptive Aδ- and C-fibers, and non-nociceptive Aα/β-fibers7,9,10,11. Therefore, based on the latency with respect to the sensory stimulus, the activity of WDR neurons provides detailed information about peripheral sensory activity5. Such recordings have been used previously to assess sensory block using direct current (DC)9 and kilohertz frequency alternating current (KHFAC)10. Both techniques use the latency of the recorded signals to discriminate between the various subclasses of sensory peripheral nerve fibers; this is because of the different conduction velocities of the subclasses of nerve fibers12. The signals are complementary because LFP recordings reflect the activity of the whole sciatic nerve but can only distinguish Aα/β from C-fibers and don't detect Aδ13,14. At the same time, WDR recordings reflect the activity of a subpopulation of sciatic nerve fibers but can discriminate Aα/β, Aδ, and C-fibers7,9,10,11. This disadvantage can be overcome by multicontact electrode arrays to simultaneously record multiple WDR neurons15.

Different peripheral nerve fiber subtypes convey different types of sensory or motor signals. For example, large, myelinated sensory fibers transmit tactile signals whilst smaller, unmyelinated fibers transmit pain signals16,17,18. When assessing nerve block technologies, it is important to understand which fiber subtypes are primarily affected because it would be clinically advantageous to block pain signals while preserving tactile sensation. Furthermore, complex temporal properties of peripheral nerve block have been observed. For example, the local anesthetic lidocaine initially blocks only unmyelinated C-fibers, then proceeds to block myelinated fibers later19. Following a complete block of all nerve fibers with lidocaine, fiber-type differences have also been observed in recovery from block20. The electrophysiological technique described here allows researchers to elucidate the temporal properties of peripheral nerve block methodologies on the various subpopulations of sensory nerve fibers. To date, we have used this method to assess block using direct current electrical block21,22, kilohertz frequency alternating current electrical block23, and photobiomodulation using near infrared wavelengths24,25. These techniques would also be suitable to investigate other interventions that act locally on a peripheral nerve, such as cold, heat, and chemical injections26,27,28.

Assessment of peripheral nerve therapies often employs disease models of inflammatory29 or neuropathic pain30 combined with behavioral tests of mechanical and thermal hypersensitivity31. The electrophysiological methods described here can quantify the neural activity changes that would contribute to a reduction in pain sensation. Furthermore, they can provide useful data on any undesirable effects of peripheral nerve block, such as the block of large sensory fibers involved in tactile sensation.

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Protocol

This protocol involving animal experiments was approved by the Institutional Animal Care and Use Committee (IACUC) of Case Western Reserve University (CWRU). Case Western Reserve University is an AAALAC-accredited institution and conforms to appropriate federal, state, and local laws and regulations, as well as institutional policies. Our research group was trained and certified by CWRU staff. When working on IACUC-approved protocols, we adhere to Public Health Service policy and the Guide for the Care and Use of Laboratory Animals (8th edition)32. This procudure is terminal and euthanasia would be triggered (through IACUC approved protocol) upon completion of planned trials or degradation of the spinal electrophysiological signals.

NOTE: The parameters for ventilation, anesthesia, and other drug administration described herein have been successfully used for adult rats of either sex in the weight range of 350-500 g. Positive pressure ventilation and thermal support allow maintaining animals for up to 6 h with good heart rate and peripheral perfusion. This amount of time is necessary for the collection of spinal electrophysiological data during multiple randomized trials. With practice, the surgical procedure can be completed in 90 min.

1. Anesthesia and ventilation

  1. Induce anesthesia in a sealed acrylic box with 5% isoflurane. Once the rat is completely unconscious, move the animal to a nose cone and titrate the isoflurane between 1-3% to maintain an appropriate level of anesthesia. Apply ophthalmic ointment to both eyes to prevent corneal drying. For shaving, use an electric clipper and remove fur from the back and the left hind leg from the iliac crest to the knee joint. Confirm the depth of anesthesia via a toe-pinch and eye blink reflexes and monitor the anesthetic plane every 15 min.
    NOTE: The rat is ready for intubation when spontaneous respiration occurs at a rate of 15-30 breaths/min. Provide and maintain thermal support throughout the procedure.
  2. Intubate the rat in a supine position with the sheath from a 14 G angiocath IV catheter (a 16 G catheter may be more appropriate for smaller animals) and a laryngoscope with a Miller 0 or 00 size blade.
  3. To improve visibility of the vocal cords, gently prop the tongue up using non-crushing forceps. Validate the intubation by chest excursion and ventilatory rate, which coincides with the rate of chest elevation.
  4. Use positive pressure ventilation at a stroke rate of 60/min and a stroke volume of 3-5 cc/stroke, while keeping the isoflurane concentration at 1%-3%.
  5. Catheterize a lateral caudal tail vein with a 22 G intravenous catheter. To improve visibility, cleanse the tail vein on the lateral side of the tail with lukewarm water and 70% isopropyl alcohol.
  6. Inject approximately 1 mL of sterile saline while carefully monitoring flow through the catheter. Any edema indicates incorrect placement of the catheter.
  7. Using polyethylene tubing, connect the inserted catheter to a syringe of rocuronium solution (2 mg/mL) mounted in a syringe driver. Do not start the infusion until ready to perform spinal recordings.

2. Surgery

NOTE: All surgical procedures should be performed on the baseplate of a small animal stereotaxic apparatus. The rest of the stereotaxic equipment can be assembled in stages as the surgery progresses.

  1. Spinal surgery
    1. For a right-handed surgeon, position the rat with the tail to the right and the head to the left. A left-handed surgeon may prefer to position the rat with the head to the right.
    2. Feel for the caudal-most rib on the left flank of the rat and follow it up to the spine. Using a scalpel, make a midline incision approximately 60 mm centered on this point.
    3. Use the scalpel to scrape the tissue from the spinous processes. Use scissors to expose the ribs and locate the caudal-most rib.
    4. Using the blunt dissection technique (insert scissors closed, then open scissors to separate tissue), confirm the caudal rib visually and tactilely. Use a surgical microscope at this step (3.5x-45x magnification). The caudal rib joins the rostral aspect of the T13 vertebra. Mark the T13 spinous process using a permanent marker.
    5. Stabilize the spine using spinal clamps at the T11 and L4 vertebrae. To do this, use blunt dissection to create a pocket on both sides of each vertebra.
    6. Slide the two vertical posts with base mounts into the T-track in the stereotax platform, align with T11 and L4 vertebrae, but do not tighten. Insert the cylindrical shafts of the vertebrae clamps into the post clamps, but do not tighten.
    7. Starting with T11, hold the spine using toothed forceps, insert the jaws of the spine clamps on either side of the vertebrae, at approximately 45° away from the T13 vertebra, and tighten the jaws of the spine clamp. Then tighten the shaft of the spine clamp in the post clamp. Repeat this process for L4 vertebra.
    8. Elevate both post clamps to lift the spine such that the torso of the rat is supported via the spine clamps (Figure 1A). This reduces respiration-related movement artifacts in the electrophysiological recordings.
    9. Mark the rostro-caudal position of the T13 and L1 spinous processes using single sutures in the muscle surface lateral to the spine.
      NOTE: The spinous process landmarks will be removed during the laminectomy.
  2. Laminectomy
    1. Temporarily position the U-shaped stereotaxic frame so that the horizontal rail can be used to rest the heels of the hands whilst performing this delicate procedure (Figure 1B). It should be removed prior to the sciatic nerve surgery.
    2. Clear muscle and connective tissue from the surface of the vertebral laminae between T12 and L3 using toothed forceps, scissors, and scraping with a scalpel. Under a surgical microscope, use Friedman-Pearson Rongeurs to remove the vertebral laminae of L2 to T13.
    3. To start, position the rongeurs close to horizontal and take small bites from the caudal portion of L2 where it overlaps L3. Extend the laminectomy rostrally to expose the midline of the spinal cord, taking care to minimize pressure on the spinal cord, and regularly clearing bone fragments from rangers.
    4. Finally, extend the laminectomy laterally 2 mm on each side of the midline. Remove the dura mater from the exposed spinal cord using fine forceps and spring scissors.
    5. Tent the dura mater prior to making any cuts. A small amount of cerebrospinal fluid will flow from the dura mater after the first cut. Gently soak this up with a twisted piece of lint-free tissue before removing the remaining section of dura mater.
    6. Cover the laminectomy with a piece of saline-dampened tissue until the electrophysiology recording period.
  3. Sciatic nerve surgery
    1. Remove the stereotaxic U-shaped frame to access the animal's leg. Expose the sciatic nerve at the mid-thigh level by making an incision with a scalpel (Figure 1C). Clear connective tissue from the nerve using blunt dissection. The amount of exposed nerve will depend on whether the nerve is being electrically stimulated.
    2. Place the device for a nerve block adjacent to the exposed nerve. Ensure the stimulation electrode and nerve block device are both proximal to the branch point at which the sciatic divides into the tibial, sural, and peroneal nerves33. Replace the stereotax frame and add the stereotax arm to prepare for electrophysiological recordings (Figure 1D).
    3. Perform stimulation for WDR neuron recordings either on the plantar surface of the hindpaw or via direct stimulation of the sciatic using a nerve cuff electrode. For LFP recordings, perform stimulation via direct stimulation of the nerve because activation via foot stimulation causes the C-fiber notch to become too diffuse to detect.
    4. If the sensory stimulation is being applied directly to the sciatic nerve, place a custom-made bipolar platinum J-cuff style electrode around the nerve34, exposed contact size 1 mm x 3 mm (comparable nerve cuff electrodes can be custom-made by MicroProbes for Life Science, Gaithersburg, MD, or World Precision Instruments, Sarasota, FL).
    5. For the most effective direct nerve stimulation, ensure the electrode is placed proximal to the branch point at which the sciatic divides into the tibial, sural, and peroneal nerves33.
    6. For plantar stimulation, insert two 13 mm stainless steel needle electrodes, inside of the 5th and outside of the 4th digits, close to the plantar surface.

3. Electrical stimulation

  1. If using nerve cuff stimulation, start systemic infusion of rocuronium solution (2 mg/mL at a rate of 1-1.4 mL/h). Systemic infusion of rocuronium should take effect within 30-60 s. Look for stimulation-related movement in the hindpaw and adjust the infusion rate as necessary to ensure that all movement is eliminated.
  2. Apply current-controlled electrical stimulation to either the plantar surface of the foot or the sciatic nerve using a biphasic rectangular isolated pulse stimulator, using stimulation parameters: pulse width 1-5 ms, pulse rate 0.1-0.2 Hz, amplitude 1-10 mA peak-to-peak.
  3. Adjust the stimulation parameters during the electrophysiological recordings, described in the following section, to ensure maximal stimulation of all fiber subtypes.
  4. The stimulator has a trigger output signal, separate from the current-controlled stimulation pulse. Connect this to the data acquisition device to synchronize the electrophysiological data with the stimulation pulse. See the next section for further details.

4. Electrophysiological recordings

  1. Insert hypodermic needles (21 G, 1 ½ inch) into the muscle on each side of the spine, parallel to the spine. Connect the grounding needle to the metal housing of each of the head stages.
  2. Connect the reference needle to each of the reference inputs on the head stages. Each of the 8 channels on the amplifier has its own BNC output port, connected to a separate analog-to-digital channel on the data acquisition device.
  3. Digitize the signals continuously (20 kHz sampling frequency) and display (Spike2 software). Using a BNC cable, connect the trigger pulse output (labeled 'TRIG OUT') from the isolated pulse stimulator to one of the digital input channels on the data acquisition device.
  4. In the software, set up the trigger channel to record Events. This will allow analysis of the LFP or WDR signals relative to the time of each electrical stimulation.
    NOTE: LFP and WDR recordings can be made with the same custom microelectrode array (1 x 8 multielectrode array, 500 µm spacing between shafts, 1 MΩ impedance, custom-made). Penetration depth and amplifier bandpass filter settings are the key differences between signal types. The array was held and connected to the stereotax using a custom 3D printed connector (Figure 2A). However, a commercially available alternative is available (Figure 2B, Standard Electrode Holder [Table of Materials]).
  5. For LFPs, insert the array oriented in the rostrocaudal direction, with the rostral-most electrode level with the suture marking the T13 spinous process, as close to midline without damaging the midline blood vessel. The strongest signals are between the cord surface and 300 µm depth.
  6. Set bandpass filters on the amplifier to 10-100 Hz, with 1,000x gain. Add a 60 Hz notch filter (or 50 Hz, depending on geographical location) to remove mains power noise. Signal strength quality will differ across the array. Select the best signal to use for further analysis.
  7. WDR neurons are distributed throughout the lumbar enlargement of the spinal cord, in the dorsal horn from 600-1200 µm deep and 200-800 µm lateral to midline, ipsilateral to the stimulation. Set bandpass filters on the amplifier to 300-10,000 Hz, with 10,000x gain. Use a notch filter for these recordings as well, although mains noise is typically removed by the bandpass filter.
  8. Adjust the stimulation parameters while observing the electrophysiological signal. Adjust the stimulation amplitude and pulse width to ensure maximal activation of all nerve fiber subtypes.
    NOTE: Large myelinated fibers (Aα/β) have the lowest activation thresholds, followed by small myelinated fibers (Aδ). Unmyelinated C-fibers require higher stimulation pulse widths and amplitudes to generate action potentials via electrical stimulation.
  9. Starting at 1 ms pulse width, increase the amplitude until all components of the signal are seen. If the higher threshold C-fibers cannot be seen, increase the stimulation pulse width.
  10. For LFP recording, observe the short latency, low threshold A-fiber response between 40-100 ms, and the longer latency, high threshold C-fiber response between 150-350 ms (Figure 3B)3,8.
  11. For WDR recordings, use the following post-stimulus latencies to identify nerve fiber subtypes: Aα/β: 0 - 25 ms, Aδ: 25 - 100 ms, and C: 100-500 ms (Figure 3C)5,6,7.

5. Peripheral nerve block

  1. Using a carbon separated nerve interface electrode (CSINE)21, apply 0.1-5 mA cathodic direct current to the sciatic nerve, proximal to the stimulating cuff (if applicable). Example block trials of LFP and WDR recordings are shown in Figure 4 and Figure 5.

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Results

The data generated using this protocol allow one to assess the relative amount of block for subclasses of sensory nerve fibers. It also allows the temporal properties of the block to be assessed; both the rate at which block in induced, and the rate of recovery following the cessation of the blocking stimulus35. For the LFP measurements the activation of fibers is determined by calculating the area under the curve for the specific time period corresponding to the fiber type of interest. In

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Discussion

This method describes the collection of two complementary electrophysiological signals that can be used to measure the efficacy of potentially therapeutic interventions to block sensory signals in peripheral nerves.

Local field potentials (LFP) can be recorded from the spinal dorsal columns using high impedance (1 MΩ) microelectrodes. They are used to measure the activation of the whole sciatic nerve; the LFP represents the responses of many cells, including excitatory postsynaptic poten...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

This work was supported by NIH NINDS R01NS116009.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1 x 8 multielectrode array, 500 µm spacing between shafts MicroprobesCustom productCustom microelectrode array 
14 G Angio catheterBencton DickinsonSKU: 381164, GTIN: 382903811649
8-channel bioamplifierWorld Precision InstrumentsISO-DAM8AHead stage
AmScope SM-6T series trinocular zoom stereo microscope 3.5-45x Magnification with articulating arm with ring lightAmscopeSM-8TW2-144SSurgical microscope
Data acquisition device Cambridge Electronic DesignPower1401-3A
Dumont #5 forcepsFine Science Tools11251-20
Friedman-Pearson Rongeurs (1mm cup size)Fine Science Tools16021-14
Harvard apparatus ventilatorHarvard Apparatus Model 683
Isoflurane, USPPiramel Critical Care66794-017-25
LaryngoscopeSarnova HC301-B802
Miller 00 laryngoscope bladeSarnova HC301-B5100-20
Paired subdermal needlesRhythmLinkRLSND121-1.013 mm stainless steel needle electrodes 
Rocuronium bromium, USPSolco43547-530-10
Spike2Cambridge Electronic Design
Square wave isolated pulse stimulator AM-SystemsModel 2100Biphasic rectangular isolated pulse stimulator
Stereotaxic platform  with adjustable base mounts with post and clampDavid Kopf InstrumentsModel 982
SureSite IV 22G x 1" safety catheter MedlineDYNA2210022 G intravenous catheter 
Syringe driverCole ParmerModel 74900
Ultra precise micro manipulator
10 micron resolution		David Kopf InstrumentsModel 961
Universal holderDavid Kopf InstrumentsModel 1772 Alternate 3D printed connector 
U-shaped stereotaxic frame David Kopf InstrumentsModel 1430
Vannas spring scissors - 2 mm cutting edgeFine Science Tools15000-03 
Vertebrae clamps David Kopf InstrumentsModel 986C 

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Peripheral Nerve BlockElectrophysiological RecordingSciatic Nerve StimulationLocal Field PotentialWide Dynamic RangeLaminectomy ProcedurePain Signal TransmissionCarbon Nerve ElectrodeDirect Current BlockRat Spinal Cord

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