The project was supported by Lundbeck Foundation, the Novo Nordisk Foundation, the Danish Medical Research Council, the Ludvig and Sara Elsass Foundation, the Foundation for Research in Neurology and Jytte and Kaj Dahlboms Foundation. R.A is supported by an Early Career Post-Doctoral Fellowship of the National Health and Medical Research Council of Australia (#1091006)

All experimental procedures described here complied with the Animal Care and Ethics Committee of UNSW Sydney and were performed in accordance with the National Health and Medical Research Council (NHMRC) of Australia regulations for animal experimentation.
1. Experimental Set Up
NOTE: 12 week old, female Long-Evans rats were used in this procedure.
<ol>
	<li>Anesthetize the rat in an induction chamber using 4% isoflurane and 1 L per min O2 flow rate. Confirm sufficient anesthesia by testing for the righting reflex and ensure its absence before removing the animal from the induction chamber. Note that various anesthetic agents have differential effects on nerve excitability10.</li>
	<li>Securely place the animal&#39;s snout in the nose cone attachment and deliver a maintenance dose of 2.5% isoflurane and 1 L per min O2 flow rate.</li>
	<li>Confirm adequate anesthesia by checking for both pedal withdrawal and corneal reflexes by pinching the animal&#39;s toes and gently touching its eye. 
	NOTE: Application of vet ointment on eyes, to prevent dryness while under anesthesia, is recommended but not essential as the procedure typically takes 30 min per animal.</li>
	<li>Maintain the body temperature of the rat at 37 &#176;C with the use of a feedback controlled heating mat and a rectal thermometer probe. Set the heating mat and integrated body temperature sensor to 40 &#176;C to prevent any skin damage that occurs with higher temperatures. 
	NOTE: It is recommended to monitor and record physiological measures (heart rate, oxygen saturation, body temperature and respiratory rate) every 10s with the use of an animal physiological monitoring system. Optimal recording procedures should include local measurement of the limb temperature as the peripheral temperature can be colder than that of the core temperature11 (Figure 1).</li>
	<li>Wear antiseptic gloves and always use instruments that have been cleaned with 70% ethanol, to maintain a pathogen-free environment throughout the procedure.</li>
</ol>
2. Electrophysiological Set Up
<ol>
	<li>Use low impedance platinum electroencephalogram (EEG) needle electrodes for this procedure.</li>
	<li>Prepare the recording needle electrodes by inserting the recording needle electrode (Figure 1; indicated in purple) through the hypothenar muscle and the reference electrode through the dorsal aspect of the 4th digit (Figure 1; indicated in orange) to record compound muscle action potentials (CMAPs).</li>
	<li>Place the ground electrode through the skin on the superior aspect of the forearm between stimulating and recording electrodes (Figure 1; indicated in green). Take care when inserting the electrodes to avoid muscle tissue.</li>
	<li>Prepare the percutaneous stimulating needle electrodes by inserting the cathode (Figure 1; labelled in blue) approximately 4 mm distal to the cubital tunnel at the elbow. Insert the anode (Figure 1; labelled in red) approximately 1 cm proximally through the skin of the axillar region.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56102/56102fig1.jpg" /> 
Figure 1: A schematic diagram of the needle electrode placements in the rat forelimb. The cathode (blue) is inserted approximately 4 mm distal to the cubital tunnel at the elbow and the anode (red) is inserted approximately 1 cm proximally through the skin of the axillar region. The ground needle electrode (green) is inserted through the skin on the superior aspect of the forearm between stimulating and recording electrodes. The recording (purple) and reference (orange) needle electrodes are inserted through the hypothenar muscle and through the dorsal aspect of the 4th digit. The temperature probe (grey) is placed on the superior aspect of the thenar muscle. <a href="//cloudfront.jove.com/files/ftp_upload/56102/56102fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Axonal Excitability Procedures
<ol>
	<li>Perform the rodent motor nerve TRONDNF protocol using a semi-automated, computer controlled axonal excitability program (see the Table of Materials) linked to a constant current stimulator and an amplifier. Remove excess 50 Hz electrical noise using a 50/60 Hz noise eliminator.
	<ol>
		<li>Record the CMAP from the hypothenar muscle by simultaneously visualizing the CMAP morphology and applying a 1 ms square-wave pulse to the ulnar nerve with the cathode needle electrode.
		<ol>
			<li>To achieve optimal recordings, carefully adjust the angle and/or position of the cathode until an optimal biphasic response curve with constant amplitude is achieved (Figure 2A.). Once the optimal position has been determined, stabilize the cathode with a repositionable electrode holder. 
			NOTE: The software used automatically delivers the test stimuli, described below, and increases or decreases the current required to achieve the threshold.</li>
		</ol>
		</li>
		<li>Record a stimulus-response curve by incrementally increasing the stimulus intensity of a 1 ms impulse by 1 mA until a maximum response is achieved. 
		NOTE: The purple and green line in figure 2B and 2C represents the incremental increase of the stimulus intensity and the automated system respectively. The target amplitude for threshold tracking is automatically set to 40% of the maximal amplitude corresponding to the area of the steepest slope on the stimulus-response curve. The change in &#39;threshold&#39; (i.e. stimulus required to elicit 40% CMAP) induced by various test stimuli is the variable of obtained throughout the remainder of the protocol.</li>
		<li>Record multiple axonal excitability parameters, including threshold electrotonus (TE), current-threshold (I/V) relationship and recovery cycle (RC) as described in reference12. 
		NOTE: The computer controlled axonal excitability program assesses the threshold (TE) with a 100 ms subthreshold depolarizing and hyperpolarizing currents that is set at &#177;20% and &#177;40% of the control threshold current. The change in threshold after both depolarizing and hyperpolarizing currents is automatically recorded at 14 time-points during the 100 ms polarizing current and 12 points after the polarizing current. Threshold electrotonus indirectly assesses the internodal conductances and is a marker of membrane potential. The I/V relationship is assessed with 200 ms subthreshold conditioning currents that alternate in intensity from +50% to -100% of the control threshold in 10% increments. The I/V relationship differs to TE in that the currents are longer in duration, the intensity of polarization is progressively altered from +50% to -100% and the threshold change is assessed at each intensity 1 ms after the polarizing current has ceased. The I/V relationship provides insight into the rectifying properties of the axon1. The RC is assessed with a paired pulse paradigm, where an initial supramaximal conditioning stimulus is applied followed by a series of test stimuli at specific intervals from 2.5 ms to 200 ms. This parameter tests a well-defined series of events that occur in the nodal region of the axon following supramaximal stimulation. These events include, the inactivation of voltage-gated Na+ channels which makes it more difficult to elicit a subsequent response and is quantified by refractoriness and the relative refractory period 1. This is followed by a charging of the internode and a period of heightened excitability mediated by fast potassium channels, quantified by superexcitability. Finally, slowly activating potassium channels mediate a late period of reduced excitability, quantified as subexcitability.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56102/56102fig2.jpg" /> 
Figure 2: Raw data from an axonal excitability test. Panel A demonstrates abiphasic CMAP response curve after incremental increase of stimulation to the ulnar nerve. Panels B and C represent the stimulus intensity (mA) and amplitude of CMAP (mV), respectively. The green component of B and C depicts the automated incremental decrease in stimulus intensity and the associated sigmoid shaped decrease in CMAP required for threshold tracking.
4. Post Electrophysiology Procedures
<ol>
	<li>Transfer the rat to a separate cage until it has regained sufficient consciousness to maintain sternal recumbency. Do not leave an animal unattended and in company of other animals until it has fully recovered from the anesthesia. Once the rat has fully recovered from the anesthesia, transfer it back into its original cage.</li>
	<li>At completion of the study, euthanize the rat without pain for the animal, e.g. lethal intraperitoneal injection of a barbiturate (e.g. lethabarb).</li>
</ol>

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Y., Krishnan, A. V., Friedlander, M. L., Kiernan, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+abnormalities+of+sensory+nerve+function+associated+with+oxaliplatin-induced+neurotoxicity.">Acute abnormalities of sensory nerve function associated with oxaliplatin-induced neurotoxicity.</a> J. Clin. Oncol. 27 (8), 1243-1249 (2009).</li><li>Arnold, R., Pussell, B. A., Pianta, T. J., Lin, C. S. -. Y., Kiernan, M. C., Krishnan, A. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Association+between+calcineurin+inhibitor+treatment+and+peripheral+nerve+dysfunction+in+renal+transplant+recipients.">Association between calcineurin inhibitor treatment and peripheral nerve dysfunction in renal transplant recipients.</a> Am. J. 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The authors have nothing to disclose.

The described procedure demonstrates a simple and reliable, minimally-invasive technique that allows assessment of the biophysical properties and membrane potential of the axon in a short period of time. Compared with other more invasive techniques, which require the exposure of the nerve, the present method of axonal excitability testing induces minimal tissue damage thus enabling in vivo assessment that preserves the physiological conditions of the nerve of interest and allows for repeated measurements.
<p...

The use of electrophysiological techniques is an essential tool for the in vivo investigation of peripheral nerve function in neurological disorders. Conventional nerve conduction methods utilize supramaximal stimuli to record motor action potential amplitude and latency. These techniques therefore provide useful information on the number of conducting fibers and on the conduction velocity of the fastest fibers. A valuable complementary tool is that of axonal excitability testing. This technique uses sophisticated electrophysiological stimulation patterns to indirectly assess the biophysical properties of peripheral nerves, such as the activity of ion channels, energy-dependent pumps, ion exchange processes and membrane potential1.
Axonal excitability testing is commonly utilized in the clinical setting to investigate pathophysiological processes and effects of therapeutic interventions on various neurological disorders. Importantly, axonal excitability measures are sensitive to therapeutic interventions that affect peripheral nerve function such as intravenous immunoglobulin (IVIg) therapy2, chemotherapy3 and calcineurin inhibitor (CNI) treatment4. Although these studies have provided important insights, clinical studies often preclude investigation of early disease traits and novel therapeutic options5. Therefore, the use of these methods in animal models of neurological disorders has recently gained traction6,7,8,9. Indeed, these methods provide an opportunity to understand the specific nerve excitability changes associated with these disorders, thus advancing translational research.
The procedure described here is a simple and reliable method to record axonal excitability measures on the ulnar nerves of the intact rat.

<table><tbody><tr><td>QTracS Program</td><td>Digitimer Ltd.</td><td></td><td>Axonal excitability program</td></tr><tr><td>AM-Systems 2200, Analog Stimulus Isolator, 2200V/50Hz</td><td>SDR Scientific</td><td>850005</td><td>Stimulator</td></tr><tr><td>High Performance AC Amplifier Model LP511</td><td>Grass Technologies</td><td></td><td>Amplifier</td></tr><tr><td>Humbug 50/60Hz Noise eliminator</td><td>Quest Scientific Instruments</td><td>726310</td><td>Noise eliminator</td></tr><tr><td>Low Impedance Platinum Monopolar Subdermal Needle Electrodes</td><td>Grass Technologies</td><td>F-E2-24</td><td>Recording electrodes, 10 mm length, 30 gauge</td></tr><tr><td>Low Impedance Platinum Electroencephalography Needle Electrodes</td><td>Cephalon</td><td>9013L0702</td><td>Stimulating electrodes, 10 mm length, 30 gauge</td></tr><tr><td>Multifunction I/O Device Model USB-6341</td><td>National Instruments</td><td></td><td>Multifunction input/output device</td></tr><tr><td>Iron Base Plate IP</td><td>Narishige Scientific Instrument Laboratory</td><td></td><td>Used for holding stimulating needle electrode in place</td></tr><tr><td>Rotating X-block X-4</td><td>Narishige Scientific Instrument Laboratory</td><td></td><td>Used for holding stimulating needle electrode in place</td></tr><tr><td>Magnetic Stand GJ-8</td><td>Narishige Scientific Instrument Laboratory</td><td></td><td>Used for holding stimulating needle electrode in place</td></tr><tr><td>Micromanipulator M-3333</td><td>Narishige Scientific Instrument Laboratory</td><td></td><td>Used for holding stimulating needle electrode in place</td></tr></tbody></table>

in vivo electrophysiological measurement of the rat ulnar nerve with axonal excitability testing

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and latency detect chronic axon loss and demyelination, respectively. Axonal excitability techniques &#34;by threshold tracking&#34; expand upon these measures by providing information regarding the activity of ion channels, pumps and exchangers that relate to acute function and may precede degenerative events. As such, the use of axonal excitability in animal models of neurological disorders may provide a useful in vivo measure to assess novel therapeutic interventions. Here we describe an experimental setup for multiple measures of motor axonal excitability techniques in the rat ulnar nerve.
The animals are anesthetized with isoflurane and carefully monitored to ensure constant and adequate depth of anesthesia. Body temperature, respiration rate, heart rate and saturation of oxygen in the blood are continuously monitored. Axonal excitability studies are performed using percutaneous stimulation of the ulnar nerve and recording from the hypothenar muscles of the forelimb paw. With correct electrode placement, a clear compound muscle action potential that increases in amplitude with increasing stimulus intensity is recorded. An automated program is then utilized to deliver a series of electrical pulses which generate 5 specific excitability measures in the following sequence: stimulus response behavior, strength duration time constant, threshold electrotonus, current-threshold relationship and the recovery cycle.
Data presented here indicate that these measures are repeatable and show similarity between left and right ulnar nerves when assessed on the same day. A limitation of these techniques in this setting is the effect of dose and time under anesthesia. Careful monitoring and recording of these variables should be undertaken for consideration at the time of analysis.

Electrophysiological measures of the rat ulnar nerve were obtained with the present protocol. Figure 3 demonstrates a representative recording from the left ulnar nerve of a 12 week-old female Long Evans rat. Compound muscle action potential relates to the number of conducting fibers that are simultaneously activated. The supramaximal peak response (mV) (Figure 3A) demonstrates the peak response achieved when incrementally increa...

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In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Axonal excitability techniques provide a powerful tool to examine pathophysiology and biophysical changes that precede irreversible degenerative events. This manuscript demonstrates the use of these techniques on the ulnar nerve of anesthetized rats.

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and ...

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9.1K Views.  University of New South Wales. Axonal excitability techniques provide a powerful tool to examine pathophysiology and biophysical changes that precede irreversible degenerative events. This manuscript demonstrates the use of these techniques on the ulnar nerve of anesthetized rats.

Video: In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Measuring Neuromuscular Junction Functionality

Neuromuscular junction (NMJ) functionality plays a pivotal role when studying diseases in which the communication between motor neuron and muscle is impaired, such as aging and amyotrophic lateral sclerosis (ALS). Here we describe an experimental protocol that can be used to measure NMJ functionality by combining two types of electrical stimulation: direct muscle membrane stimulation and the stimulation through the nerve. The comparison of the muscle response to these two different stimulations can help to define, at the functional level, potential alterations in the NMJ that lead to functional decline in muscle.
Ex vivo preparations are suited to well-controlled studies. Here we describe an intensive protocol to measure several parameters of muscle and NMJ functionality for the soleus-sciatic nerve preparation and for the diaphragm-phrenic nerve preparation. The protocol lasts approximately 60 min and is conducted uninterruptedly by means of a custom-made software that measures the twitch kinetics properties, the force-frequency relationship for both muscle and nerve stimulations, and two parameters specific to NMJ functionality, i.e. neurotransmission failure and intratetanic fatigue. This methodology was used to detect damages in soleus and diaphragm muscle-nerve preparations by using SOD1G93A transgenic mouse, an experimental model of ALS that ubiquitously overexpresses the mutant antioxidant enzyme superoxide dismutase 1 (SOD1).

A functional assessment of the neuromuscular junction (NMJ) can provide essential information on the communication between muscle and nerve. Here we describe a protocol to comprehensively evaluate both the NMJ and muscle functionality using two different muscle-nerve preparations, i.e. soleus-sciatic and diaphragm-phrenic.

Neuromuscular junction (NMJ) functionality plays a pivotal role when studying diseases in which the communication between motor neuron and muscle is ...

Establishing a Mouse Model of a Pure Small Fiber Neuropathy with the Ultrapotent Agonist of Transient Receptor Potential Vanilloid Type 1

Patients with diabetes mellitus (DM) or those experiencing the neurotoxic effects of chemotherapeutic agents may develop sensation disorders due to degeneration and injury of small-diameter sensory neurons, referred to as small fiber neuropathy. Present animal models of small fiber neuropathy affect both large- and small-diameter sensory fibers and thus create a neuropathology too complex to properly assess the effects of injured small-diameter sensory fibers. Therefore, it is necessary to develop an experimental model of pure small fiber neuropathy to adequately examine these issues. This protocol describes an experimental model of small fiber neuropathy specifically affecting small-diameter sensory nerves with resiniferatoxin (RTX), an ultrapotent agonist of transient receptor potential vanilloid type 1 (TRPV1), through a single dose of intraperitoneal injection, referred to as RTX neuropathy. This RTX neuropathy showed pathological manifestations and behavioral abnormalities that mimic the clinical characteristics of patients with small fiber neuropathy, including intraepidermal nerve fiber (IENF) degeneration, specifically injury in small-diameter neurons, and induction of thermal hypoalgesia and mechanical allodynia. This protocol tested three doses of RTX (200, 50, and 10 &#181;g/kg, respectively) and concluded that a critical dose of RTX (50 &#181;g/kg) is required for the development of typical small fiber neuropathy manifestations, and prepared a modified immunostaining procedure to investigate IENF degeneration and neuronal soma injury. The modified procedure is fast, systematic, and economic. Behavioral evaluation of neuropathic pain is critical to reveal the function of small-diameter sensory nerves. The evaluation of mechanical thresholds in experimental rodents is particularly challenging and this protocol describes a customized metal mesh that is suitable for this type of assessment in rodents. In summary, RTX neuropathy is a new and easily established experimental model to evaluate the molecular significance and intervention underlying neuropathic pain for the development of therapeutic agents.

This study establishes an experimental model of pure small fiber neuropathy with resiniferatoxin (RTX). A unique dose of RTX (50 &#181;g/kg) is optimal for developing a small fiber neuropathy model that mimics patient characteristics and could help investigate the nociceptive molecular significance underlying neuropathic pain.

Patients with diabetes mellitus (DM) or those experiencing the neurotoxic effects of chemotherapeutic agents may develop sensation disorders due to ...

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings provide a sensitive approach to measure nerve conduction in humans and rodent models. To broaden the technical possibilities for electromyography in mice, the measurement of compound muscle action potentials (CMAPs) from the brachial plexus nerve in the forelimb using needle electrodes is described here. CMAP recordings after stimulating the sciatic nerve in hindlimbs have been previously described. The newly introduced method here allows for the evaluation of the nerve conductivity at an additional site, and thus provides a more profound overview of the neuromuscular functionality. The technique provides information on both the relative number of functional axons and the myelination level. Thereby, this method can be applied to assess both axonal diseases as well as demyelinating conditions. This minimally invasive method does not require extraction of the nerve and therefore it is suitable for repeated measurements for longitudinal follow-up in the same animal. Similar recordings are performed in clinical setups to emphasize the translational relevance of the method.

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

The measurement of nerve conduction is a useful tool to assess mouse models of neurodegeneration but it is frequently only applied to stimulate the sciatic nerve in hindlimbs. Here, we describe a technique to measure compound muscle action potential (CMAP) in vivo in the mouse forelimb muscles innervated by the brachial plexus.

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings ...

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for nerve fibers in the central and peripheral nervous systems. This method is particularly applicable to dorsal root ganglia (DRG), which are primary sensory neurons that exhibit a pseudo-unipolar structure of nervous processes. The patterns and features of the action potentials passed along axons are recordable in these neurons. The present study uses in vivo single-fiber recordings to observe the conduction failure of sciatic nerves in complete Freund&#8217;s adjuvant (CFA)-treated rats. As the underlying mechanism cannot be studied using in vivo single-fiber recordings, patch-clamp-recordings of DRG neurons are performed on preparations of intact DRG with the attached sciatic nerve. These recordings reveal a positive correlation between conduction failure and the rising slope of the after-hyperpolarization potential (AHP) of DRG neurons in CFA-treated animals. The protocol for in vivo single fiber-recordings allows the classification of nerve fibers via the measurement of conduction velocity and monitoring of abnormal conditions in nerve fibers in certain diseases. Intact DRG with attached peripheral nerve allows observation of the activity of DRG neurons in most physiological conditions. Conclusively, single-fiber recording combined with electrophysiological recording of intact DRGs is an effective method to examine the role of conduction failure during the analgesic process.

Single-fiber recording is an effective electrophysiological technique that is applicable to the central and peripheral nervous systems. Along with the preparation of intact DRG with the attached sciatic nerve, the mechanism of conduction failure is examined. Both protocols improve the understanding of the peripheral nervous system&#39;s relationship with pain.

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for ...

Functional and Physiological Methods of Evaluating Median Nerve Regeneration in the Rat

The main goal of this investigation is to show how to create and repair different types of median nerve (MN) lesions in the rat. Moreover, different methods of simulating postoperative physiotherapy are presented. Multiple standardized strategies are used to assess motor and sensory recovery using an MN model of peripheral nerve lesion and repair, thus permitting easy comparison of the results. Several options are included for providing a postoperative physiotherapy-like environment to rats that have undergone MN injuries. Finally, the paper provides a method to evaluate the recovery of the MN using several noninvasive tests (i.e., grasping test, pin prick test, ladder rung walking test, rope climbing test, and walking track analysis), and physiological measurements (infrared thermography, electroneuromyography, flexion strength evaluation, and flexor carpi radialis muscle weight determination). Hence, this model seems particularly appropriate to replicate a clinical scenario, facilitating extrapolation of results to the human species.
Although the sciatic nerve is the most studied nerve in peripheral nerve research, analysis of the rat MN presents various advantages. For example, there is a reduced incidence of joint contractures and automutilation of the affected limb in MN lesion studies. Furthermore, the MN is not covered by muscle masses, making its dissection easier than that of the sciatic nerve. In addition, MN recovery is observed sooner, because the MN is shorter than the sciatic nerve. Also, the MN has a parallel path to the ulnar nerve in the arm. Hence, the ulnar nerve can be easily used as the nerve graft for repairing MN injuries. Finally, the MN in rats is located in the forelimb, akin to the human upper limb; in humans, the upper limb is the site of most peripheral nerve lesions.

Presented is a protocol to produce different types of median nerve (MN) lesions and repair in the rat. Additionally, the protocol shows how to evaluate the functional recovery of the nerve using several noninvasive behavioral tests and physiological measurements.

The main goal of this investigation is to show how to create and repair different types of median nerve (MN) lesions in the rat. Moreover, different ...

Medicine

Maximum Isometric Tetanic Force Measurement of the Tibialis Anterior Muscle in the Rat

Traumatic nerve injuries result in substantial functional loss and segmental nerve defects often necessitate the use of autologous interposition nerve grafts. Due to their limited availability and associated donor side morbidity, many studies in the field of nerve regeneration focus on alternative techniques to bridge a segmental nerve gap. In order to investigate the outcomes of surgical or pharmacological experimental treatment options, the rat sciatic nerve model is often used as a bioassay. There are a variety of outcome measurements used in rat models to determine the extent of nerve regeneration. The maximum output force of the target muscle remains the most relevant outcome for clinical translation of experimental therapies. Isometric force measurement of tetanic muscle contraction has previously been described as a reproducible and valid technique for evaluating motor recovery after nerve injury or repair in both rat and rabbit models. In this video, we will provide a step-by-step instruction of this invaluable procedure for assessment of functional recovery of the tibialis anterior muscle in a rat sciatic nerve defect model using optimized parameters. We will describe the necessary pre-surgical preparations in addition to the surgical approach and dissection of the common peroneal nerve and tibialis anterior muscle tendon. The isometric tetanic force measurement technique will be detailed. Determining the optimal muscle length and stimulus pulse frequency is explained and measuring the maximum tetanic muscle contraction is demonstrated.

Evaluation of motor recovery remains the benchmark outcome measure in experimental peripheral nerve studies. The isometric tetanic force measurement of the tibialis anterior muscle in the rat is an invaluable tool to assess functional outcomes after reconstruction of sciatic nerve defects. The methods and nuances are detailed in this article.

Traumatic nerve injuries result in substantial functional loss and segmental nerve defects often necessitate the use of autologous interposition nerve ...

Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

Ex vivo preparations enable the study of many neurophysiological processes in isolation from the rest of the body while preserving local tissue structure. This work describes the preparation of rat sciatic nerves for ex vivo neurophysiology, including buffer preparation, animal procedures, equipment setup and neurophysiological recording. This work provides an overview of the different types of experiments possible with this method. The outlined method aims to provide 6 h of stimulation and recording on extracted peripheral nerve tissue in tightly controlled conditions for optimal consistency in results. Results obtained using this method are A-fibre compound action potentials (CAP) with peak-to-peak amplitudes in the millivolt range over the entire duration of the experiment. CAP amplitudes and shapes are consistent and reliable, making them useful to test and compare new electrodes to existing models, or the effects of interventions on the tissue, such as the use of chemicals, surgical alterations, or neuromodulatory stimulation techniques. Both conventional commercially available cuff electrodes with platinum-iridium contacts and custom-made conductive elastomer electrodes were tested and gave similar results in terms of nerve stimulus strength-duration response.

Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

This protocol describes the preparation of rat whole sciatic nerve tissue for ex vivo electrophysiological stimulation and recording in an environmentally-regulated, two-compartment, perfused saline bath.

Ex vivo preparations enable the study of many neurophysiological processes in isolation from the rest of the body while preserving local tissue structure. ...

Terminal H-reflex Measurements in Mice

The Hoffmann reflex (H-reflex), as an electrical analog to the stretch reflex, allows electrophysiological validation of the integrity of neural circuits after injuries such as spinal cord damage or stroke. An increase of the H-reflex response, together with symptoms like non-voluntary muscle contractions, pathologically augmented stretch reflex, and hypertonia in the corresponding muscle, is an indicator of post-stroke spasticity (PSS).
In contrast to rather nerve-unspecific transcutaneous measurements, here, we present a protocol to quantify the H-reflex directly at the ulnar and median nerves of the forepaw, which is applicable, with minor modifications, to the tibial and sciatic nerve of the hindpaw. Based on the direct stimulation and the adaptation to different nerves, the method represents a reliable and versatile tool to validate electrophysiological changes in spasticity-related disease models.

The clinical evaluation of spasticity based on the Hoffmann reflex (H-reflex) and using electrical stimulation of peripheral nerves is an established method. Here, we provide a protocol for a terminal and direct nerve stimulation for H-reflex quantification in the mouse forepaw.

The Hoffmann reflex (H-reflex), as an electrical analog to the stretch reflex, allows electrophysiological validation of the integrity of neural circuits ...

Biology

In Vivo Measurement of the Compound Muscle Action Potential in a Rat

Source: Wild, B. M., et al. <a href="https://app.jove.com/v/56102">In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing.</a> J. Vis. Exp. (2018)
This video demonstrates the procedure for recording compound muscle action potential (CMAP) in rats by applying electrical stimuli to the ulnar nerve, adjusting electrodes for optimal signal, and increasing stimulation to assess nerve functionality. This method enables the in vivo investigation of the peripheral nerve functions in neurological disorders.

Pomiar in vivo złożonego potencjału czynnościowego mięśni u szczura

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

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In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

Functional and Physiological Methods of Evaluating Median Nerve Regeneration in the Rat

Maximum Isometric Tetanic Force Measurement of the Tibialis Anterior Muscle in the Rat

Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

Terminal H-reflex Measurements in Mice

Pomiar in vivo złożonego potencjału czynnościowego mięśni u szczura