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.remotexs.ntu.edu.sg/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>

<ol>
	<li>Krishnan, A. V., Lin, C. S. -. Y., Park, S. B., 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=Axonal+ion+channels+from+bench+to+bedside:+a+translational+neuroscience+perspective.">Axonal ion channels from bench to bedside: a translational neuroscience perspective.</a> Prog neurobiol. 89 (3), 288-313 (2009).</li><li>Lin, C. S. -. Y., Krishnan, A. V., Park, S. B., 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=Modulatory+effects+on+axonal+function+after+intravenous+immunoglobulin+therapy+in+chronic+inflammatory+demyelinating+polyneuropathy.">Modulatory effects on axonal function after intravenous immunoglobulin therapy in chronic inflammatory demyelinating polyneuropathy.</a> Arch neurol. 68 (7), 862-869 (2011).</li><li>Park, S. B., Goldstein, D., Lin, C. S. -. 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. Transplant. 13 (9), 2426-2432 (2013).</li><li>Bo&#235;rio, D., Greensmith, L., Bostock, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Excitability+properties+of+motor+axons+in+the+maturing+mouse.">Excitability properties of motor axons in the maturing mouse.</a> J. Peripher. Nerv. Syst. 14 (1), 45-53 (2009).</li><li>Bo&#235;rio, D., Kalmar, B., Greensmith, L., Bostock, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Excitability+properties+of+mouse+motor+axons+in+the+mutant+SOD1(G93A)+model+of+amyotrophic+lateral+sclerosis.">Excitability properties of mouse motor axons in the mutant SOD1(G93A) model of amyotrophic lateral sclerosis.</a> Muscle &amp; Nerve. 41 (6), 774-784 (2010).</li><li>Alvarez, S., Calin, A., Graffmo, K. S., Moldovan, M., Krarup, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peripheral+motor+axons+of+SOD1(G127X)+mutant+mice+are+susceptible+to+activity-dependent+degeneration.">Peripheral motor axons of SOD1(G127X) mutant mice are susceptible to activity-dependent degeneration.</a> Neurosci. 241, 239-249 (2013).</li><li>Fledrich, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Soluble+neuregulin-1+modulates+disease+pathogenesis+in+rodent+models+of+Charcot-Marie-Tooth+disease+1A.">Soluble neuregulin-1 modulates disease pathogenesis in rodent models of Charcot-Marie-Tooth disease 1A.</a> Nat. Med. 20 (9), 1055-1061 (2014).</li><li>Vianello, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Low+doses+of+arginine+butyrate+derivatives+improve+dystrophic+phenotype+and+restore+membrane+integrity+in+DMD+models.">Low doses of arginine butyrate derivatives improve dystrophic phenotype and restore membrane integrity in DMD models.</a> FASEB J. 28 (6), 2603-2619 (2014).</li><li>Osaki, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+anesthetic+agents+on+in+vivo+axonal+HCN+current+in+normal+mice.">Effects of anesthetic agents on in vivo axonal HCN current in normal mice.</a> Clin Neurophysiol. 126 (10), 2033-2039 (2015).</li><li>Biessels, G. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenotyping+animal+models+of+diabetic+neuropathy:+a+consensus+statement+of+the+diabetic+neuropathy+study+group+of+the+EASD+(Neurodiab).">Phenotyping animal models of diabetic neuropathy: a consensus statement of the diabetic neuropathy study group of the EASD (Neurodiab).</a> J. Peripher. Nerv. Syst. 19 (2), 77-87 (2014).</li><li>Bo&#235;rio, D., Greensmith, L., Bostock, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+model+of+mouse+motor+nerve+excitability+and+the+effects+of+polarizing+currents.">A model of mouse motor nerve excitability and the effects of polarizing currents.</a> J. Peripher. Nerv. Syst. 16 (4), 322-333 (2011).</li><li>Arnold, R., Moldovan, M., Rosberg, M. R., Krishnan, A. V., Morris, R., Krarup, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nerve+excitability+in+the+rat+forelimb:+a+technique+to+improve+translational+utility.">Nerve excitability in the rat forelimb: a technique to improve translational utility.</a> J. Neurosci. Methods. 275, 19-24 (2017).</li><li>Moldovan, M., Alvarez, S., Krarup, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Motor+axon+excitability+during+Wallerian+degeneration.">Motor axon excitability during Wallerian degeneration.</a> Brain. 132 (Pt 2), 511-523 (2009).</li><li>Madison, R. D., Robinson, G. A., Krarup, C., Moldovan, M., Li, Q., Wilson, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+electrophoresis+and+in+vivo+electrophysiology+of+peripheral+nerve+using+DC+field+stimulation.">In vitro electrophoresis and in vivo electrophysiology of peripheral nerve using DC field stimulation.</a> J. Neurosci. Methods. 225, 90-96 (2014).</li><li>Moldovan, M., Krarup, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+Na+/K++pump+function+following+repetitive+activity+in+mouse+peripheral+nerve.">Evaluation of Na+/K+ pump function following repetitive activity in mouse peripheral nerve.</a> J. Neurosci. Methods. 155 (2), 161-171 (2006).</li></ol>

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.
To ensure consistent results there are a few methodological considerations that need to be addressed. One such factor is the concentration and type of anesthetic. It has been previously demonstrated that isoflurane has an effect on hyperpolarization-activated cyclic nucleotide-gated channels10. Alternatively, an injectable anesthetic mixture of medetomidine, midazolam and butorphanol was investigated and also demonstrated effects on nerve excitability over time10. Medetomidine/midazolam/fentanyl (MMF) appears to have little effect on nerve excitability and has been successfully used in many nerve excitability studies7,13,14, though its effects over time on nerve excitability have not been systematically investigated. Of note, this anesthetic is difficult to obtain in the US and Australia as fentanyl is a controlled substance with strict import regulations. Regardless of chosen anesthesia, dosage and time under anesthesia should be carefully monitored for consideration at analysis.
Another factor to consider is electrode quality. Due to the strong and prolonged hyperpolarizing and depolarizing currents applied in axonal excitability studies high quality electrodes are required. In this study, low impedance platinum electroencephalogram (EEG) needle electrodes were used. Low impedance electrodes are typically within the kilo-ohms range and the platinum EEG electrodes conventionally operate within the 0.5 - 5 kilo-ohm range which qualifies as low impedance electrodes. The axonal excitability program can record the current output and calculate impedance versus the target out voltage and it has been previously established that the current was stable for 30 min, which is the typical length of this protocol15. Additionally, these electrodes have been recently introduced in mice16 and were found to not be subject to polarizing effects14,15. Therefore, polarization would not be a problem during the experimental procedure.
In contrast to protocols that expose the nerve of interest, these studies utilize an intact model which makes positioning of stimulating electrodes approximate. Thus, exact replication of electrode placement in longitudinal studies may be difficult. Despite this, a previous study with different anesthesia demonstrated good repeatability of 30/34 ulnar nerve excitability parameters across 3 separate studies similar to that seen in tibial and caudal nerves13. Furthermore, in this study, comparison of right and left CMAPs showed consistency (Figure 4), suggesting precise and proper electrode placement is sufficient to offset this possible limitation.
There are several critical steps in the process of acquiring CMAP responses. Proper and consistent placement of the stimulating needle electrode is crucial for the reproducible measurements of the amplitude. Additionally, it is essential to ensure proper placement of recording needle electrodes to minimize background noise. Therefore, simultaneously visualizing the response curve while placing the stimulating needle electrode is crucial to confirm consistent placement.
The use of this minimally-invasive technique on animal models of neurological disorders may be a valuable tool to investigate pathophysiological changes and early disease traits. These may act as biomarkers and facilitate the investigation of novel therapeutic interventions in conjunction with behavioral measures of hand function. Moreover, validation of these techniques in rodents may enable the investigation of pharmacokinetic and pharmacodynamic relationships of new compounds. This may enable better translation of therapeutic compounds prior to Phase 1 and 2 clinical trials

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 increasing the stimulus until the there is no change in the response (Figure 2B).
Figure 3B represents the current-voltage (I/V) relationship produced by currents longer in duration (200 ms) and their intensities changed in 10% increments from +50% to -100% of the threshold. The I/V relationship assesses inward and outward rectification by examining differences in threshold current in response to depolarizing and hyperpolarizing currents. The bottom left quadrant reflects inward rectification accommodating to hyperpolarization and activation of inwardly rectifying conductances1. The upper right quadrant reflects fast and slow K+ channel activation and the outward rectification accommodating to the depolarizing current.
Internodal conductances can be examined using threshold electrotonus waveform (Figure 3C) in response to long subthreshold depolarizing and hyperpolarizing currents. Both hyperpolarizing and depolarizing TE have variables that can be calculated by averaging the threshold change between specific time points 10-20 ms, 20-40 ms and 90-100 ms.
Changes over time due to injury or the application of therapeutic interventions may cause specific changes to nerve excitability parameters. This may provide useful in vivo information regarding pathophysiological changes, early disease traits and therapeutic efficacy in animal models of neurological disorders.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56102/56102fig3.jpg" /> 
Figure 3: A representative axonal excitability plot. (A) Stimulus-response curve depicting the supermaximal peak response (mV). (B) The current-voltage (I/V) relationship produced by a 200ms polarizing stimulus ranging from +50% to -100% of the threshold current. (C) Threshold electrotonus illustrating waveforms in response to prolonged subthreshold polarizing current elicited by a 20% and 40% depolarizing (above 0 on y axis) and hyperpolarizing (below 0 on the y axis) current. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56102/56102fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Sequential axonal excitability testing was undertaken on the left then right forelimb of n=4 rats (12 week-old). Both left and right recordings were completed within 35 min after the loss of the pedal withdrawal reflex. The analysis was performed using the paired non-parametric Wilcoxon signed rank test. These analyses revealed no significant differences between left and right ulnar nerves for any of the axonal excitability variables. Consistency was demonstrated in standard nerve conduction parameters, CMAP amplitude and latency (Figure 4A and B) as well as nerve excitability variables including superexcitability and threshold electrotonus hyperpolarizing (TEh) 90-100 (Figure 4C and D). However, previous studies10 have indicated significant change over time in parameters does occur under isoflurane anesthesia (see discussion).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56102/56102fig4.jpg" /> 
Figure 4: Mean recordings (n=4) obtained for (A) peak response (B) latency (C) superexcitability and (D) hyperpolarizing threshold electrotonus (90-100ms) in both left (red) and right (blue) ulnar nerves. The error bars indicate the standard error of the mean (SEM). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56102/56102fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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

Nerve Excitability Assessment in Chemotherapy-induced Neurotoxicity

Chemotherapy-induced neurotoxicity is a serious consequence of cancer treatment, which occurs with some of the most commonly used chemotherapies1,2. Chemotherapy-induced peripheral neuropathy produces symptoms of numbness and paraesthesia in the limbs and may progress to difficulties with fine motor skills and walking, leading to functional impairment. In addition to producing troubling symptoms, chemotherapy-induced neuropathy may limit treatment success leading to dose reduction or early cessation of treatment. Neuropathic symptoms may persist long-term, leaving permanent nerve damage in patients with an otherwise good prognosis3. As chemotherapy is utilised more often as a preventative measure, and survival rates increase, the importance of long-lasting and significant neurotoxicity will increase.
There are no established neuroprotective or treatment options and a lack of sensitive assessment methods. Appropriate assessment of neurotoxicity will be critical as a prognostic factor and as suitable endpoints for future trials of neuroprotective agents. Current methods to assess the severity of chemotherapy-induced neuropathy utilise clinician-based grading scales which have been demonstrated to lack sensitivity to change and inter-observer objectivity4. Conventional nerve conduction studies provide information about compound action potential amplitude and conduction velocity, which are relatively non-specific measures and do not provide insight into ion channel function or resting membrane potential. Accordingly, prior studies have demonstrated that conventional nerve conduction studies are not sensitive to early change in chemotherapy-induced neurotoxicity4-6. In comparison, nerve excitability studies utilize threshold tracking techniques which have been developed to enable assessment of ion channels, pumps and exchangers in vivo in large myelinated human axons7-9.
Nerve excitability techniques have been established as a tool to examine the development and severity of chemotherapy-induced neurotoxicity10-13. Comprising a number of excitability parameters, nerve excitability studies can be used to assess acute neurotoxicity arising immediately following infusion and the development of chronic, cumulative neurotoxicity. Nerve excitability techniques are feasible in the clinical setting, with each test requiring only 5 -10 minutes to complete. Nerve excitability equipment is readily commercially available, and a portable system has been devised so that patients can be tested in situ in the infusion centre setting. In addition, these techniques can be adapted for use in multiple chemotherapies.
In patients treated with the chemotherapy oxaliplatin, primarily utilised for colorectal cancer, nerve excitability techniques provide a method to identify patients at-risk for neurotoxicity prior to the onset of chronic neuropathy. Nerve excitability studies have revealed the development of an acute Na+ channelopathy in motor and sensory axons10-13. Importantly, patients who demonstrated changes in excitability in early treatment were subsequently more likely to develop moderate to severe neurotoxicity11. However, across treatment, striking longitudinal changes were identified only in sensory axons which were able to predict clinical neurological outcome in 80% of patients10. These changes demonstrated a different pattern to those seen acutely following oxaliplatin infusion, and most likely reflect the development of significant axonal damage and membrane potential change in sensory nerves which develops longitudinally during oxaliplatin treatment10. Significant abnormalities developed during early treatment, prior to any reduction in conventional measures of nerve function, suggesting that excitability parameters may provide a sensitive biomarker.

This abstract describes a novel method to assess the development of neurotoxicity in patients receiving chemotherapy treatment. While conventional assessment methods are limited in their ability to detect early changes in nerve function, nerve excitability techniques provide early identification of patients at risk of severe neurotoxicity and insight into pathophysiology.

Chemotherapy-induced neurotoxicity is a serious consequence of cancer treatment, which occurs with some of the most commonly used chemotherapies1,2. ...

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 ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Visual Evoked Potential Recording in a Rat Model of Experimental Optic Nerve Demyelination

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to monitor the disease course in the follow-up period. Changes in the VEP parameters closely correlate with pathological damage in the optic nerve. This protocol provides a detailed description about the rodent model of optic nerve microinjection, in which a partial demyelination lesion is produced in the optic nerve. VEP recording techniques are also discussed. Using skull implanted electrodes, we are able to acquire reproducible intra-session and between-session VEP traces. VEPs can be recorded on individual animals over a period of time to assess the functional changes in the optic nerve longitudinally. The optic nerve demyelination model, in conjunction with the VEP recording protocol, provides a tool to investigate the disease processes associated with demyelination and remyelination, and can potentially be employed to evaluate the effects of new remyelinating drugs or neuroprotective therapies.

Focal demyelination is induced in the optic nerve using lysolecithin microinjection. Visual evoked potentials are recorded via skull electrodes implanted over the visual cortex to examine the signal conduction along the visual pathway in vivo. This protocol details the surgical procedures underlying electrode implantation and optic nerve microinjection.

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to ...

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

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 ...

Screening of Axonal Degeneration in Carpal Tunnel Syndrome Using Ultrasonography and Nerve Conduction Studies

Axonal degeneration, indicative of surgical decompression, may coexist in carpal tunnel syndrome (CTS) as the disease progresses. However, the current diagnostic and severity gradation system cannot clearly indicate its coexistence, resulting in&#160;confusion of appropriate treatment prescription. There are also constraints in conventional methods for&#160;differentiation as well. This study aims at introducing an innovative, efficient, and quick screening protocol to differentiate axonal degeneration associated with CTS, using ultrasound and nerve conduction studies (NCS). It starts by using NCS to perform orthodromic stimulation at the wrist, to obtain the sensory conduction of the median and the ulnar nerves respectively. Meanwhile, the motor conduction&#160;of the median nerve is collected by stimulating the palm, wrist, and elbow, followed by the stimulation of the ulnar nerve at the wrist,&#160;below and above the elbow. Then, an ultrasound assessment is performed, using a linear array transducer, with cross-sectional area (CSA) and perimeter (P) at the wrist and at the one-third distal forearm calipered. Ratios (R-CSA, R-P) and changes from wrist to one-third distal forearm (&#916;CSA and &#916;P) are calculated according to a standard format. Potential axonal degeneration coexisting in CTS will be screened according to the criteria of NCS and cut-off values of ultrasound measurements established in a previous study. In terms of its noninvasiveness, low cost, convenience, and efficiency, it is easy to apply ultrasound complimentarily in clinical practice to prescreen patients with potential coexisting axonal degeneration. Nevertheless, the ultrasonographic imaging cannot directly reflect axonal degeneration. It still relies on conventional but invasive methods&#160;such as electromyography (EMG) and biopsy&#160;for confirmation if needed.

Here we present a protocol using nerve conduction studies and ultrasound to screen potential axonal degeneration associated with carpal tunnel syndrome. The criteria for differentiation are established. Compared to conventional approaches, this method is noninvasive, convenient, and efficient, with an overall satisfactory accuracy, sensitivity, and specificity.

Axonal degeneration, indicative of surgical decompression, may coexist in carpal tunnel syndrome (CTS) as the disease progresses. However, the current ...

Facial Nerve Surgery in the Rat Model to Study Axonal Inhibition and Regeneration

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial nerve can be manipulated along its entire length, from its intracranial segment to its extratemporal course. There are three primary types of nerve injury used for the experimental study of regenerative properties: nerve crush, transection, and nerve gap. The range of possible interventions is vast, including surgical manipulation of the nerve, delivery of neuroactive reagents or cells, and either central or end-organ manipulations. Advantages of this model for studying nerve regeneration include simplicity, reproducibility, interspecies consistency, reliable survival rates of the rat, and an increased anatomic size relative to murine models. Its limitations involve a more limited genetic manipulation versus the mouse model and the superlative regenerative capability of the rat, such that the facial nerve scientist must carefully assess time points for recovery and whether to translate results to higher animals and human studies. The rat model for facial nerve injury allows for functional, electrophysiological, and histomorphometric parameters for the interpretation and comparison of nerve regeneration. It thereby boasts tremendous potential toward furthering the understanding and treatment of the devastating consequences of facial nerve injury in human patients.

This protocol describes a reproducible approach to facial nerve surgery in the rat model, including descriptions of various inducible patterns of injury.

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial ...