The authors acknowledge Dr. Gerald Hunsberger of GlaxoSmithKline Pharmaceuticals, King of Prussia, PA, USA, and Galvani Bioelectronics (Stevenage, UK) for sharing their original nerve preparation technique with us. The authors acknowledge Robert Toth for the Dual-Chamber nerve bath design. The authors acknowledge funding from the Healthcare Technologies Challenge Awards (HTCA) grant of the Engineering and Physical Sciences Research Council (EPSRC). The authors acknowledge the High Performance Embedded and Distributed Systems Centre for Doctoral Training (HiPEDS CDT) of Imperial College London for funding Adrien Rapeaux (EP/L016796/1 ). Adrien Rapeaux is currently funded by the UK Dementia Research Institute, Care Research and Technology Centre.&#160;The authors gratefully acknowledge Zack Bailey of Imperial College, in the Department of Bioengineering, for help with experiments and access to animal tissues during the production of the JoVE video article.

All animal care and procedures were performed under appropriate licenses issued by the UK Home office under the Animals (Scientific Procedures) Act (1986) and were approved by the Animal Welfare and Ethical Review Board of Imperial College London.
1. Preparation of buffers
NOTE: This part of the protocol can be carried out well in advance of the rest of the protocol, except for the final steps involving the preparation of modified Krebs-Henseleit Buffer (mKHB) at 1x concentration.
<ol>
	<li>Prepare 1 M CaCl2 stock solution
	<ol>
		<li>Add 14.701 g of CaCl2 dihydrate to a clean 100 mL beaker. Add ~75 mL of deionized water and stir until complete dissolution of the salt.</li>
		<li>Transfer the solution to a 100 mL graduated flask and add deionized water until 100 mL volume has been reached. Transfer the solution to a bottle and store it in a refrigerator at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare 10x concentrated mKHB stock
	<ol>
		<li>Add 66.03 g of sodium chloride (NaCl), 3.57 g of potassium chloride (KCl), 1.63 g of potassium dihydrogen phosphate (KH2PO4), and 1.44 g of magnesium sulfate to a 2 L beaker.</li>
		<li>Add ~750 mL of deionized water to the beaker and stir until the salts have dissolved (there may be small salt crystals left at the bottom). Add 25 mL of 1 M CaCl2 stock solution (step 1.1) and stir; ensure this is the last salt added.</li>
		<li>Transfer the solution to a 1 L graduated flask and add deionized water to reach a total volume of 1 L. Transfer the solution to a 1 L bottle. Store concentrated 10x mKHB at 4 &#176;C in a refrigerator. 
		NOTE: Concentrated mKHB stock can be stored for approximately 1 month before replacement.</li>
	</ol>
	</li>
	<li>Prepare dissection Petri dish (coating)
	<ol>
		<li>Prepare a clean glass Petri dish (120 mm diameter) for coating by washing and drying the dish carefully.</li>
		<li>Following the manufacturer&#39;s instructions for usage and curing of the conformal alkoxy coating, coat the bottom of the Petri Dish with ~3-5 mm of coating by carefully pouring the conformal coating mixture into the dish until the desired thickness has been reached.</li>
		<li>Cure the coating in an oven at 60 &#176;C until it is firm to the touch. The dissection Petri dish is now ready. 
		&#8203;NOTE: Gentle cleaning of the dish after each use will ensure that the coating lasts for years before replacement is needed. When replacing the coating, ensure to remove all the coating material before applying a new coating. Note that the conformal alkoxy coating (Table of Materials) used in this protocol has a shelf life shorter than a year.</li>
	</ol>
	</li>
</ol>
2. Pre-dissection preparations
NOTE: This step starts the experiment. The below steps must be carried out on the same day, in this order.
<ol>
	<li>Prepare 1x mKHB
	<ol>
		<li>Prepare a clean 2 L beaker for the buffer. Transfer 200 mL of 10x mKHB stock to the 2 L beaker. Add 2.1 g of sodium carbonate (NaHCO3) and 0.99 g of anhydrous Dextrose (D-glucose) to the 2 L beaker.</li>
		<li>Add approximately 1 L of deionized water to the beaker. Stir until salts have been completely dissolved. Transfer the solution to a 2 L graduated flask and add deionized water to reach a total volume of 2 L.</li>
		<li>Transfer the solution to a 2 L glass bottle placed on a heating stirrer with the temperature set to 37 &#176;C. 
		NOTE: Smaller heating stirrer will often be unable to reach the target 37 &#176;C temperature due to the thermal inertia of large containers full of water. Adjust the temperature upward so the contents of the bottle reach 37 &#176;C with stirring. Monitor the temperature during the experiment and lower the set temperature in the event of overshoot.</li>
		<li>Place a thermometer in the 2 L bottle to monitor the temperature, using grippers to prevent the thermometer from being impacted by the stirring flea. Aerate the buffer with carbogen for a minimum of 30 min to oxygenate the solution. This should set the pH to 7.4. Measure the pH with a pH meter to ensure it is within 0.1 pH units of 7.4 (at 37 &#176;C). 
		NOTE: To adjust the pH to 7.4, use hydrochloric acid or sodium hydroxide when needed.</li>
		<li>Fill two 15 mL centrifuge tubes and one 100 mL bottle with mKHB and place them on ice to cool. 
		NOTE: The centrifuge tubes and the bottle can be cleaned and reused after each experiment without autoclaving.</li>
		<li>For later parts of the experiment, continue to aerate the buffer in the 2 L bottle with carbogen at 37 &#176;C with continuous stirring. 
		NOTE: Unsupervised use of carbogen may be dangerous if no automatic systems are in place to shut off gas flow in the event of a leak. An automated sensor and electronic shutoff valve can mitigate this; otherwise, a person must be left to supervise the equipment if dissection takes place in a different room. In all cases, an oxygen sensor should be used near the setup to warn operators when ambient oxygen concentration rises above 25%. Use a room with active ventilation if possible.</li>
	</ol>
	</li>
	<li>Turn on the signal acquisition device, low-noise preamplifier, line noise filter, and oscilloscope for recording and stimulation, to allow enough time for temperature stabilization.</li>
	<li>Ensure all electrical recording equipment is correctly configured
	<ol>
		<li>Set the low-noise preamplifier to AC-coupled input with the input band-pass filter set to 6 dB roll-off per decade, and cutoff frequencies set to 30 Hz and 3 kHz for the high-pass and low-pass filters, respectively.</li>
		<li>Set the gain of the low-noise preamplifier to 100.</li>
	</ol>
	</li>
</ol>
3. Animal anesthesia and euthanization
NOTE: Female rats between 250 and 330 g (Table of Materials) were used for the studies.
<ol>
	<li>Prepare surgical tools and consumables: 12 cm straight scissors (blunt); 2 mm cutting edge angled spring scissors; 4 cm fine scissors, sharp or semi-sharp; #7 Dumont forceps; 45&#176; angled fine forceps and 6-0 suturing silk or thread.</li>
	<li>Place the rat in an anaesthetization container or chamber. Connect oxygen and the anesthetic diffuser to the container. Set anesthetic (isoflurane) concentration to 3.5% and wait approximately 10 min or until the animal shows signs of anesthesia such as loss of righting reflex.</li>
	<li>After the rat shows signs of anesthesia, confirm the loss of consciousness with a toe pinch withdrawal reflex test. Proceed only when there is no toe withdrawal; otherwise, check all the connections and anesthetic levels in the diffuser and repeat step 3.2.</li>
	<li>Take the animal out of the container and proceed with cervical dislocation, followed by incision of a femoral artery for confirmation of death.</li>
</ol>
4. Dissection protocol
NOTE: Place the animal with its belly down on the dissection table. Repeat the following steps for both legs. Typically, the right leg is dissected first.
<ol>
	<li>Holding the ankle firmly between the thumb, index finger, and middle finger, sever the calcaneal tendon using 12 cm straight blunt scissors.</li>
	<li>With fine sharp scissors, make a skin incision from the calcaneal tendon along the back of the leg all the way up to the base of the spine, taking care not to dissect the muscle tissue below.</li>
	<li>Using fine forceps and fine scissors, make careful incisions through the muscle layers near the middle of the back of the leg until the sciatic nerve is exposed. As soon as the sciatic nerve is visible, moisturize the cavity using ice-cold mKHB to prevent the nerve from drying out.</li>
	<li>Using hemostats, pull the flaps of skin on each side apart, and maintain the incision open for finer dissection work. Starting from the location of the calcaneal tendon incision, with fine scissors, interrupt the muscle on the medial side of the leg to free the nerve. Continue to maintain moisture levels in the area with ice-cold mKHB.</li>
	<li>As the nerve is exposed while moving up the leg, dissect the overlying muscle tissue. Free the nerve nearer to the spine from connective tissues until reaching the spinal cleft at which point there is a kink in the nerve. Do not attempt to clean the nerve yet as speed is essential at this stage.</li>
	<li>Sever the nerve as close to the spine as possible with fine scissors. To make the dissection easier, very gently pull the end of the nerve near the ankle using forceps. Never pinch the nerve in the middle, but only the ends. Never pull a nerve taut. 
	OPTIONAL: At this point, if time allows, suturing of both ends of the nerve can be carried out to help maintain viability. Keep handling to a minimum to prevent damage. If there is insufficient time (see step 4.5), skip this step and follow step 5.2 later on.</li>
	<li>Place the dissected nerve in the 15 mL centrifuge tube filled with mKHB (step 2.1.5), close the tube, and place the tube back on ice until the start of the cleaning procedure.</li>
	<li>Repeat steps 4.1-4.7 for the other leg. Ideally, each nerve should take 5-10 min to extract, in order to maximize tissue viability.</li>
</ol>
5. Nerve cleaning procedure
<ol>
	<li>Fill the coated Petri dish approximately halfway with chilled oxygenated mKHB. Place one of the dissected sciatic nerves in the dish and pin both ends of the nerve to the dish such that the nerve is straight without kinks, torsion, or twists. Pin the nerve as close to the ends as possible.</li>
	<li>Using 6-0 silk sutures or fine thread, tie a double knot around each end of the nerve to prevent cytosol leakage into the buffer. Place the knots just next to the insect pins on the side closer to the center of the nerve, as this will prevent leakage from nerve tissue into the buffer. Do not carry out this step if the nerve has been previously ligated.</li>
	<li>Using the microscope for precision and the 2 mm angled spring scissors, remove fat, blood vessels, and muscle tissue from the nerve. Prune any nerve branches that will not be used in the stimulation and recording protocol.
	<ol>
		<li>Every 5 min of cleaning, replace the buffer with fresh chilled oxygenated mKHB. Use fine forceps to pull on connective tissues, fat, and blood vessels to ease dissection.</li>
	</ol>
	</li>
	<li>Place the nerves back in the transport tubes filled with fresh oxygenated chilled mKHB and place the tubes on ice.</li>
</ol>
6. Equipment setup
NOTE: The equipment setup used to carry out experiments is illustrated in Figure 1. Briefly, it consists of a dual-compartment nerve bath, a 2 L bottle placed on a heating stirrer, a source of carbogen for buffer aeration, and tubing to allow the buffer to flow from the bottle to the bath, and back to the bottle using a peristaltic pump. The bath can be machined out of plexiglass or 3D-printed from watertight materials. It has a depth of approximately 2 cm, and the partition separating the two chambers of the bath features a 1.5 mm diameter hole to allow the threading of a peripheral nerve across both chambers. One chamber is large, must be at least 4 or 5 cm long, and will be filled with buffer. The other chamber should be at least 3 cm long and will be filled with silicone or mineral oil. The bath must not be made too large as this will degrade control of perfusion, temperature, and pH. Different bath sizes may be required depending on the size of the nerve tissue being studied.
<ol>
	<li>Prepare a clean dual-chamber nerve bath (see Supplementary File for design specifications). Place the nerve bath below the level of the 2 L bottle placed on the heating stirrer, using standard laboratory boss heads and grippers. Connect the drain of the bath to the peristaltic pump inlet.</li>
	<li>Connect the outlet of the peristaltic pump to a tube leading back to the 2 L buffer bottle. Connect the bath inlet to a tube with an adjustable flow valve and put the tube inside the 2 L bottle. Use a three-way valve with a syringe connected to the middle outlet to help with priming the tube as a siphon for gravity-assisted buffer inflow.</li>
	<li>Prime the siphon by drawing the syringe until buffer flows into it. Configure the valve such that the flow rate of buffer into the bath is ~5-6 mL&#183;min-1. The flow can be increased initially to fill the bath. Once the bath buffer level has reached the drain, place the nerves in the bath.</li>
	<li>Using an insect pin, secure the end of the nerve at the corner of the buffer-filled bath chamber. Using 45&#176; angled fine forceps and pinching the nerve only at the ends, carefully thread the nerve to be stimulated through the hole in the partition between the two bath chambers.</li>
	<li>Secure the other end of the nerve in the oil chamber of the bath with an insect pin, ensuring that the nerve is straight without being stretched and is free of kinks and twists. Using silicone grease, make a seal to prevent buffer leakage from the buffer chamber into the oil chamber. Fill the oil chamber with silicone or mineral oil.</li>
	<li>Place the Ag/AgCl recording electrode hooks in the oil bath chamber and secure them using boss heads and grippers. Drape the portion of the nerve in the oil bath over the hooks without pulling the nerve taut. Do not pinch the nerve; use angled forceps to lift the nerve without pinching.</li>
	<li>Adjust or repair the silicone grease seal if any leakage is observed after moving the nerve.</li>
	<li>Connect the reference Ag/AgCl electrode to the amplifier ground and place the electrode in the buffer-filled bath chamber by securing it using a laboratory gripper.</li>
</ol>
7. Electrode implantation on the nerve in the bath
<ol>
	<li>Prepare a clean nerve cuff electrode for stimulation according to reference21. Place the electrode in the buffer-filled bath chamber. Using forceps or fine tweezers with blunt or angled tips, open the electrode in the bath to wet the inside of the cuff.</li>
	<li>If bubbles remain, use a fine syringe to draw buffer from the bath and force the bubbles out of the cuff. With a tweezer under the nerve, gently open the cuff and slide it under the nerve. Close the cuff around the nerve, taking care to avoid any kinking or twisting of the nerve.</li>
	<li>Connect the stimulation electrode to the stimulator and secure the stimulation electrode lead with tape. Connect the current return electrode to the stimulator and secure the lead with tape. If using a square platinum sheet as the current return electrode, position the sheet away from the nerve in the bath.</li>
</ol>
8. Stimulation and recording
<ol>
	<li>Connect the stimulator TTL signal output to channel 4 of the oscilloscope, which will be used to trigger the oscilloscope. On the oscilloscope screen, press the Trigger channel tab, and specify Channel 4 as the triggering channel. Set the trigger level to 1 V using the Level knob.</li>
	<li>On the oscilloscope, set time resolution to 1 ms/division and voltage resolution to 10 mV/division. Center the trigger reference in time and set the trigger level to 1 V. 
	NOTE: Follow steps 8.3 to 8.6 if using the custom stimulator (see Table of Materials). It is assumed that the MATLAB software and device drivers have been installed on the laboratory computer using instructions provided freely online for the custom neural stimulator21. Otherwise, follow the manufacturer&#39;s instructions for the use of a commercially available stimulator as an alternative.</li>
	<li>Connect the stimulator to the laboratory computer. Turn on the stimulator by connecting the battery power supply to the power input. Start the MATLAB software on the laboratory computer.</li>
	<li>Execute the custom MATLAB script: HFAC_4ch_Stimulator_Initialization.m (Table of Materials). 
	NOTE: The USB communication cable between the computer and stimulator should flash green. If it does not, then there is a configuration error, and the power supply and connections must be verified.</li>
	<li>Open the MATLAB script: HFAC_4ch_Monophasic_Stimulation.m (Table of Materials). By directly editing the MATLAB script, set the parameters as follows: stimulator pulse amplitude = -300 &#181;A, stimulator pulse width = 300 &#181;s, stimulator number of pulses = 10 and stimulator time between pulses = 1 s.</li>
	<li>Start the stimulation protocol by clicking on Run in the MATLAB software.</li>
</ol>

<ol><li>McIntyre, C. C., Richardson, A. G., Grill, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Modeling+the+excitability+of+mammalian+nerve+fibers%3A+Influence+of+afterpotentials+on+the+recovery+cycle%5BTitle%5D)+AND+%22Journal+of+Neurophysiology%22%5BJournal%5D)">Modeling the excitability of mammalian nerve fibers: Influence of afterpotentials on the recovery cycle.</a> Journal of Neurophysiology. 87 (2), 995-1006 (2002).</li>
<li>Pelot, N. A., Grill, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((In+vivo+quantification+of+excitation+and+kilohertz+frequency+block+of+the+rat+vagus+nerve%5BTitle%5D)+AND+%22Journal+of+Neural+Engineering%22%5BJournal%5D)">In vivo quantification of excitation and kilohertz frequency block of the rat vagus nerve.</a> Journal of Neural Engineering. 17 (2), 026005(2020).</li>
<li>Patel, Y. A., Kim, B. S., Rountree, W. S., Butera, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Kilohertz+electrical+stimulation+nerve+conduction+block%3A+Effects+of+electrode+surface+area%5BTitle%5D)+AND+%22IEEE+Transactions+on+Neural+Systems+and+Rehabilitation+Engineering%22%5BJournal%5D)">Kilohertz electrical stimulation nerve conduction block: Effects of electrode surface area.</a> IEEE Transactions on Neural Systems and Rehabilitation Engineering. 25 (10), 1906-1916 (2017).</li>
<li>Kortelainen, J., Al-Nashash, H., Vipin, A., Thow, X. Y., All, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+effect+of+anaesthesia+on+somatosensory+evoked+potential+measurement+in+a+rat+model%5BTitle%5D)+AND+%22Laboratory+Animals%22%5BJournal%5D)">The effect of anaesthesia on somatosensory evoked potential measurement in a rat model.</a> Laboratory Animals. 50 (1), 63-66 (2016).</li>
<li>Oh, S. S., Hayes, J. M., Sims-Robinson, C., Sullivan, K. A., Feldman, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+effects+of+anesthesia+on+measures+of+nerve+conduction+velocity+in+male+C57Bl6%2FJ+mice%5BTitle%5D)+AND+%22Neuroscience+Letters%22%5BJournal%5D)">The effects of anesthesia on measures of nerve conduction velocity in male C57Bl6/J mice.</a> Neuroscience Letters. 483 (2), 127-131 (2010).</li>
<li>Kuffler, S. W., Williams, E. M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Small-nerve+junctional+potentials.+The+distribution+of+small+motor+nerves+to+frog+skeletal+muscle%2C+and+the+membrane+characteristics+of+the+fibres+they+innervate%5BTitle%5D)+AND+%22The+Journal+of+Physiology%22%5BJournal%5D)">Small-nerve junctional potentials. The distribution of small motor nerves to frog skeletal muscle, and the membrane characteristics of the fibres they innervate.</a> The Journal of Physiology. 121 (2), 289-317 (1953).</li>
<li>Brunton, E., Blau, C. W., Nazarpour, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Separability+of+neural+responses+to+standardised+mechanical+stimulation+of+limbs%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Separability of neural responses to standardised mechanical stimulation of limbs.</a> Scientific Reports. 7 (1), 11138(2017).</li>
<li>Schmalbruch, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fiber+composition+of+the+rat+sciatic+nerve%5BTitle%5D)+AND+%22The+Anatomical+Record%22%5BJournal%5D)">Fiber composition of the rat sciatic nerve.</a> The Anatomical Record. 215 (1), 71-81 (1986).</li>
<li>Kagiava, A., Theophilidis, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Assessing+the+permeability+of+the+rat+sciatic+nerve+epineural+sheath+against+compounds+with+local+anesthetic+activity%3A+an+ex+vivo+electrophysiological+study%5BTitle%5D)+AND+%22Toxicology+Mechanisms+and+Methods%22%5BJournal%5D)">Assessing the permeability of the rat sciatic nerve epineural sheath against compounds with local anesthetic activity: an ex vivo electrophysiological study.</a> Toxicology Mechanisms and Methods. 23 (8), 634-640 (2013).</li>
<li>Motori, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Neuronal+metabolic+rewiring+promotes+resilience+to+neurodegeneration+caused+by+mitochondrial+dysfunction%5BTitle%5D)+AND+%22Science+Advances%22%5BJournal%5D)">Neuronal metabolic rewiring promotes resilience to neurodegeneration caused by mitochondrial dysfunction.</a> Science Advances. 6 (35), 8271(2020).</li>
<li>Schwarz, J. R., Eikhof, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Na+currents+and+action+potentials+in+rat+myelinated+nerve+fibres+at+20+and+37%C2%B0+C%5BTitle%5D)+AND+%22Pfl%C3%BCgers+Archiv%22%5BJournal%5D)">Na currents and action potentials in rat myelinated nerve fibres at 20 and 37° C.</a> Pflügers Archiv. 409 (6), 569-577 (1987).</li>
<li>Cranefield, P. F., Brink, F., Bronk, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+oxygen+uptake+of+the+peripheral+nerve+of+the+rat%5BTitle%5D)+AND+%22Journal+of+Neurochemistry%22%5BJournal%5D)">The oxygen uptake of the peripheral nerve of the rat.</a> Journal of Neurochemistry. 1 (3), 245-249 (1957).</li>
<li>Lehmann, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+effect+of+changes+in+pH+on+the+action+of+mammalian+A+nerve+fibers%5BTitle%5D)+AND+%22American+Journal+of+Physiology-Legacy+Content%22%5BJournal%5D)">The effect of changes in pH on the action of mammalian A nerve fibers.</a> American Journal of Physiology-Legacy Content. 118 (3), 600-612 (1937).</li>
<li>Hamm, L. L., Nakhoul, N., Hering-Smith, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Acid-base+homeostasis%5BTitle%5D)+AND+%22Clinical+Journal+of+the+American+Society+of+Nephrology%3A+CJASN%22%5BJournal%5D)">Acid-base homeostasis.</a> Clinical Journal of the American Society of Nephrology: CJASN. 10 (12), 2232-2242 (2015).</li>
<li>Minasian, S. M., Galagudza, M. M., Dmitriev, Y. V., Kurapeev, D. I., Vlasov, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Myocardial+protection+against+global+ischemia+with+Krebs-Henseleit+buffer-based+cardioplegic+solution%5BTitle%5D)+AND+%22Journal+of+Cardiothoracic+Surgery%22%5BJournal%5D)">Myocardial protection against global ischemia with Krebs-Henseleit buffer-based cardioplegic solution.</a> Journal of Cardiothoracic Surgery. 8, 60(2013).</li>
<li>Bailey, L. E., Ong, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Krebs-Henseleit+solution+as+a+physiological+buffer+in+perfused+and+superfused+preparations%5BTitle%5D)+AND+%22Journal+of+Pharmacological+Methods%22%5BJournal%5D)">Krebs-Henseleit solution as a physiological buffer in perfused and superfused preparations.</a> Journal of Pharmacological Methods. 1 (2), 171-175 (1978).</li>
<li>Miller, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Sydney+Ringer%3A+physiological+saline%2C+calcium+and+the+contraction+of+the+heart%5BTitle%5D)+AND+%22The+Journal+of+Physiology%22%5BJournal%5D)">Sydney Ringer: physiological saline, calcium and the contraction of the heart.</a> The Journal of Physiology. 555, Pt 3 585-587 (2004).</li>
<li>Yamamoto, D., Suzuki, N., Miledi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Blockage+of+chloride+channels+by+HEPES+buffer%29%5BTitle%5D)+AND+%22Proceedings+of+the+Royal+Society+of+London.+Series+B.+Biological+Sciences%22%5BJournal%5D)">Blockage of chloride channels by HEPES buffer).</a> Proceedings of the Royal Society of London. Series B. Biological Sciences. 230 (1258), 93-100 (1987).</li>
<li>Janz, G. J., Taniguchi, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+silver-silver+halide+electrodes.+Preparation%2C+stability%2C+and+standard+potentials+in+aqueous+and+non-aqueous+media%5BTitle%5D)+AND+%22Chemical+Reviews%22%5BJournal%5D)">The silver-silver halide electrodes. Preparation, stability, and standard potentials in aqueous and non-aqueous media.</a> Chemical Reviews. 53 (3), 397-437 (1953).</li>
<li>Rapeaux, A., Constandinou, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+HFAC+block-capable+and+module-extendable+4-channel+stimulator+for+acute+neurophysiology%5BTitle%5D)+AND+%22Journal+of+Neural+Engineering%22%5BJournal%5D)">An HFAC block-capable and module-extendable 4-channel stimulator for acute neurophysiology.</a> Journal of Neural Engineering. 17 (4), 046013(2020).</li>
<li>Next-Generation-Neural-Interfaces/HFAC_Stimulator_4ch. Next Generation Neural Interfaces. , Available from: 
 <a target="_blank" href="https://github.com/Next-Generation-Neural-Interfaces/HFAC_Stimulator_4ch>">https://github.com/Next-Generation-Neural-Interfaces/HFAC_Stimulator_4ch></a> (2021).</li>
<li>Cuttaz, E. A., Chapman, C. A. R., Syed, O., Goding, J. A., Stretchable Green, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((fully+polymeric+electrode+arrays+for+peripheral+nerve+stimulation%5BTitle%5D)+AND+%22Advanced+Science%22%5BJournal%5D)">fully polymeric electrode arrays for peripheral nerve stimulation.</a> Advanced Science. 8 (8), 2004033(2021).</li>
<li>Lossi, L., Merighi, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+use+of+ex+vivo+rodent+platforms+in+neuroscience+translational+research+with+attention+to+the+3Rs+philosophy%5BTitle%5D)+AND+%22Frontiers+in+Veterinary+Science%22%5BJournal%5D)">The use of ex vivo rodent platforms in neuroscience translational research with attention to the 3Rs philosophy.</a> Frontiers in Veterinary Science. 5, 164(2018).</li>
</ol>

The authors have no conflicts of interest.
OPEN ACCESS STATEMENT: 
For the purpose of open-access, the author has applied a Creative Common Attribution (CC BY) license (where permitted by UKRI, &#8216;Open Government Licence&#8217; or &#8216;Creative Commons Attribution No-derivatives (CC BY-ND) licence may be stated instead) to any Author Accepted Manuscript version arising.

DATA SHARING: 
The raw data used in the figures of this article will be made available by the authors, without undue reservation.

In this work, we described a protocol to prepare rat sciatic nerves for ex vivo neurophysiology. Tissue extraction takes approximately 30 min, including animal handling, anesthesia, culling, and dissection, while nerve cleaning, placement in the bath, and electrode implantation should require an additional 30 min before recording can be started. Buffer preparation can be carried out in 30 min, though this can be done ahead of the rest of the experiment. This type of preparation and experiment has been used and d...

The current understanding of fundamental nerve function as modeled in silico is lacking in several aspects, notably with respect to the effects of nerve tissue compartmentalization outside of the soma, axon, and dendrites. Axon-myelin interactions are still poorly understood as evidenced by the fact that even detailed computational nerve models such as MRG1 (for mammalian nerves) that adequately capture conventional electrical stimulation response, do not capture other experimentally observed behaviors such as high-frequency block carryover2 or secondary onset response3.
This protocol provides a method to efficiently investigate neurophysiological processes at the nerve level in an acute small laboratory animal model, using a standardized preparation protocol to isolate the nerve, control its environment, and remove it from an in vivo context to an ex vivo context. This will prevent other body processes or anesthetics used by in vivo nerve stimulation protocols to alter nerve behavior and confound measured results or their interpretation4,5. This enables the development of more realistic models focusing solely on effects specific to nerve tissues that are poorly understood. This protocol is also useful as a testbed for new nerve stimulation and recording electrode materials and geometries, as well as new stimulation paradigms such as high-frequency block2,3. Variations of this technique have been used previously to study nerve physiology in tightly controlled conditions6, for example, to measure ion channel dynamics and properties or the effects of local anesthetics7.
This technique provides several advantages compared to alternatives such as acute in vivo small animal experimentation8. The technique obviates the need to maintain anesthesia depth as the tissue has been extracted from the body, reducing the amount of required equipment such as an anesthetic diffuser, oxygen concentrator, and heating pad. This simplifies the experimental protocol, reducing the risk of mistakes. As anesthetics can potentially alter nerve function4, this technique ensures that measures will not be confounded by side effects from these anesthetic compounds. Finally, this technique is more appropriate than acute in vivo experiments when studying the effects of neurotoxic compounds such as tetrodotoxin, which would kill an anesthetized animal by paralysis.
Peripheral nerve sections are a unique ex vivo system since there is a high chance that the fibers responsible for recorded neural signals do not contain any soma. As these would normally be located, for motor neurons, in the spine, and for sensory neurons in the dorsal root ganglia next to the spine, the preparation of a section of the mammalian nerve can be roughly modeled as a collection of tubular membranes with ion channels, open at both ends9. Metabolism is maintained by the mitochondria located in the axon at the time of tissue dissection10. Suturing of the open ends of the axolemma is encouraged after extraction to close them and thereby help maintain existing ionic gradients across the membrane, which are essential for normal nerve function.
To maintain tissue homeostasis outside the body, several environmental variables must be tightly controlled. These are temperature11, oxygenation12, osmolarity, pH13,14, and access to glucose to maintain metabolism. For this protocol, the approach is to use a modified Krebs-Henseleit buffer15,16 (mKHB) continuously aerated with a mixture of oxygen and carbon dioxide. The mKHB is in the family of cardioplegic buffers6,17 used to preserve dissected tissues outside of the body, for example, in ex vivo experiments. These buffers do not contain any hemoglobin, antibiotics, or antifungals and are, therefore, only suitable for preparations involving small amounts of tissue for a limited time. pH control was achieved with the carbonate and carbon dioxide redox pair, requiring constant aeration of the buffer with carbon dioxide to maintain pH equilibrium. This is to avoid using other common buffering agents such as HEPES, which can modify nerve cell function18. To oxygenate the buffer and provide pH control, a mixture of 5% carbon dioxide in oxygen called carbogen (95% O2, 5% CO2) was used. A heating stirrer was used for temperature control of a buffer container, and the buffer was perfused through a nerve bath, and then recirculated to the starting container. A typical experiment would last 6-8 h before the nerve loses its viability and no longer responds sufficiently to stimulation for measures to be representative of healthy tissue.
To optimize the signal-to-noise ratio, silver-chloride electrodes were used for recording, which were prepared according to previously described methods19. For stimulation, a combination of commercial off-the-shelf platinum cuff electrodes and custom-made conductive polymer cuff electrodes can be used. Conductive polymer cuff electrodes have notably higher charge capacities, which are useful when stimulating the nerve using high amplitude waveforms20.
The stimulator used in this protocol has been previously described20. Documentation, design files, and software scripts to use it are publicly available21. Other stimulators can be used to execute this protocol; however, the custom stimulator is also capable of high-frequency alternative current (HFAC) block2,20, which enables a wider range of neurophysiology experiments. To use HFAC block, conductive elastomer cuffs are recommended to avoid damage to the nerve. Conductive elastomer nerve cuffs are soft and fully polymeric electrode arrays produced from conductive elastomers as the conductive component and polydimethylsiloxane as the insulation22. Devices were manufactured in a bipolar configuration using conventional laser microfabrication techniques.

<table><tbody><tr><td>1 L Glass bottle</td><td>VWR International Ltd</td><td>215-1595</td><td>Borosilicate glass</td></tr><tr><td>1 L Glass graduated flask</td><td>VWR International Ltd</td><td>612-3626</td><td>Borosilicate glass</td></tr><tr><td>2 L Glass bottle</td><td>VWR International Ltd</td><td>215-1596</td><td>Borosilicate glass</td></tr><tr><td>2 L Glass graduated flask</td><td>VWR International Ltd</td><td>BRND937254</td><td>Borosilicate glass</td></tr><tr><td>Adaptor, pneumatic, 8 mm to 1/4 NPT</td><td>RS UK</td><td>536-2599</td><td>push-to-fit straight adaptor between oxygen hose and gas dispersion tube</td></tr><tr><td>Alkoxy conformal coating</td><td>Farnell</td><td>1971829</td><td>ACC15 Alkoxy conformal coating for dissection petri dish preparation</td></tr><tr><td>Anesthetic</td><td>Chanelle</td><td>N/A</td><td>Isoflurane inhalation anesthetic, 250 mL bottle</td></tr><tr><td>Beaker, 2 L</td><td>VWR International Ltd</td><td>213-0469</td><td>Borosilicate glass</td></tr><tr><td>Bipolar nerve cuff</td><td>Cortec GMBH</td><td>N/A</td><td>800 micron inner diameter, perpendicular lead out, no connector termination</td></tr><tr><td>Bossheads</td><td>N/A</td><td>N/A</td><td>Standard wet laboratory bossheads for attaching grippers to rods</td></tr><tr><td>Calcium Chloride dihydrate</td><td>Sigma Aldrich</td><td>C7902-500g</td><td>500 g in plastic bottle</td></tr><tr><td>Carbogen canister</td><td>BOC</td><td>N/A</td><td>F-size canister</td></tr><tr><td>Centrifuge Tubes, 15 mL volume</td><td>VWR International Ltd</td><td>734-0451</td><td>Falcon tubes</td></tr><tr><td>Conductive elastomer nerve cuff</td><td>N/A</td><td>N/A</td><td>high charge capacity nerve cuff for stimulation, see protocol for fabrication reference</td></tr><tr><td>Connector, Termimate</td><td>Mouser UK</td><td>538-505073-1100-LP</td><td>These should be soldered to wire terminated with crocodile clips (see entry 11)</td></tr><tr><td>Crocodile clip connectors</td><td>RS UK</td><td>212-1203</td><td>These should be soldered to wire terminated with TermiMate connectors (see entry 10)</td></tr><tr><td>Deionized Water</td><td>N/A</td><td>N/A</td><td>Obtained from deionized water dispenser</td></tr><tr><td>Forceps angled 45 degrees</td><td>InterFocus Ltd</td><td>91110-10</td><td>Fine forceps, student range</td></tr><tr><td>Forceps standard Dumont #7</td><td>InterFocus Ltd</td><td>91197-00</td><td>Student range forceps</td></tr><tr><td>Gas Disperson Tube, Porosity 3</td><td>Merck</td><td>12547866</td><td>N/A</td></tr><tr><td>Glucose anhydrous, powder</td><td>VWR International Ltd</td><td>101174Y</td><td>500 g in plastic bottle</td></tr><tr><td>Grippers</td><td>N/A</td><td>N/A</td><td>Standard wet laboratory rod-mounted grippers</td></tr><tr><td>Heating Stirrer</td><td>RS UK</td><td>768-9672</td><td>Stuart US152</td></tr><tr><td>Hemostats</td><td>N/A</td><td>N/A</td><td>Any hemostat &gt;12 cm in length is suitable</td></tr><tr><td>Insect Pins, stainless steel, size 2</td><td>InterFocus Ltd</td><td>26001-45</td><td>N/A</td></tr><tr><td>Laptop computer</td><td>N/A</td><td>N/A</td><td>Any laboratory-safe portable computer with at least 2 unused USB ports is suitable</td></tr><tr><td>Line Noise Filter</td><td>Digitimer</td><td>N/A</td><td>Humbug noise eliminator (50 Hz line noise filter)</td></tr><tr><td>Low-Noise Preamplifier, SR560</td><td>Stanford Research Systems</td><td>SR560</td><td>Low-noise voltage preamplifier</td></tr><tr><td>Magnesium Sulphate salt</td><td>VWR International Ltd</td><td>291184P</td><td>500g in plastic bottle</td></tr><tr><td>MATLAB scripts</td><td>Github</td><td>https://github.com/Next-Generation-Neural-Interfaces/HFAC_Stimulator_4ch</td><td>Initialization, calibration and stimulation scripts for the custom stimulator</td></tr><tr><td>MATLAB software</td><td>Mathworks</td><td>N/A</td><td>Standard package</td></tr><tr><td>Microscope Light, PL-2000</td><td>Photonic</td><td>N/A</td><td>Light source with swan necks. Product may be obtained from third party supplier</td></tr><tr><td>Microscope, SMZ 745</td><td>Nikon</td><td>SM745</td><td>Stereoscopic Microscope</td></tr><tr><td>Mineral oil, non-toxic</td><td>VWR International Ltd</td><td>31911.A1</td><td>Oil for nerve bath</td></tr><tr><td>Nerve Bath</td><td>N/A</td><td>N/A</td><td>Plexiglas machined nerve bath, see protocol for details.</td></tr><tr><td>Oscilloscope</td><td>LeCroy</td><td>N/A</td><td>434 Wavesurfer. Product may be obtained from 3rd party suppliers</td></tr><tr><td>Oxygen Hose, 1 meter</td><td>BOC</td><td>N/A</td><td>1/4&quot; NPT terminations</td></tr><tr><td>Oxygen Regulator</td><td>BOC</td><td>C106X/2B:3.5BAR-BS3-1/4&quot;NPTF 230Bar</td><td>N/A</td></tr><tr><td>Peristaltic Pump P-1</td><td>Pharmacia Biotech</td><td>N/A</td><td>Product may be obtained from third party supplier</td></tr><tr><td>Petri Dish, Glass</td><td>VWR International Ltd</td><td>391-0580&amp;nbsp;</td><td>N/A</td></tr><tr><td>Potassium Chloride salt</td><td>Sigma Aldrich</td><td>P5405-250g</td><td>250 g in plastic bottle</td></tr><tr><td>Potassium Dihydrogen Sulphate salt</td><td>Merck</td><td>1.04873.0250</td><td>250 g in plastic bottle</td></tr><tr><td>Rat</td><td>Charles River Laboratories</td><td>N/A</td><td>Sprague Dawley, 250-330 grams, female</td></tr><tr><td>Reference electrode, ET072</td><td>eDaQ (Australia)</td><td>ET072-1</td><td>Silver silver-chloride reference electrode</td></tr><tr><td>Rod</td><td>N/A</td><td>N/A</td><td>Standard wet laboratory rods with fittings for stands</td></tr><tr><td>Scale</td><td>Sartorius</td><td>N/A</td><td>M-Power scale, for weighing powders. Product may be obtained from third-party suppliers</td></tr><tr><td>Scissors straight 12 cm edge</td><td>InterFocus Ltd</td><td>91400-12</td><td>blunt-blunt termination, student range</td></tr><tr><td>Signal Acquisition Device</td><td>Cambridge Electronic Design</td><td>Micro3-1401</td><td>Micro3-1401 Multichannel ADC</td></tr><tr><td>Silicone grease, non-toxic</td><td>Farnell</td><td>3821559</td><td>for sealing of bath partition</td></tr><tr><td>Silicone tubing, 2 mm inner diameter</td><td>N/A</td><td>N/A</td><td>N/A</td></tr><tr><td>Silicone tubing, 5 mm inner diameter</td><td>N/A</td><td>N/A</td><td>N/A</td></tr><tr><td>Silver wire</td><td>Alfa Aesar</td><td>41390</td><td>0.5 mm, annealed</td></tr><tr><td>Sodium Bicarbonate salt</td><td>Sigma Aldrich</td><td>S5761-500g</td><td>500 g in plastic bottle</td></tr><tr><td>Sodium Chloride salt</td><td>VWR International Ltd</td><td>27810.295</td><td>1 kg in plastic bottle</td></tr><tr><td>Spring scissors angled 2 mm edge</td><td>InterFocus Ltd</td><td>15010-09</td><td>N/A</td></tr><tr><td>Stand</td><td>N/A</td><td>N/A</td><td>Standard wet laboratory stands with sockets for rods</td></tr><tr><td>Stimulator</td><td>Digitimer</td><td>DS3</td><td>DS3 or Custom Stimulator (see references)</td></tr><tr><td>Stirring flea</td><td>VWR International Ltd</td><td>442-0270</td><td>For use with the heating stirrer</td></tr><tr><td>Syringe tip, blunt, 1 mm diameter</td><td>N/A</td><td>N/A</td><td>N/A</td></tr><tr><td>Syringe tip, blunt, 2 mm diameter</td><td>N/A</td><td>N/A</td><td>N/A</td></tr><tr><td>Syringe, plastic, 10 mL volume</td><td>N/A</td><td>N/A</td><td>syringe should have luer lock fitting</td></tr><tr><td>Tape, water-resistant</td><td>N/A</td><td>N/A</td><td>For securing tubing and wiring to workbench</td></tr><tr><td>Thermometer</td><td>VWR International Ltd</td><td>620-0806</td><td>glass thermometer</td></tr><tr><td>USB Power Bank</td><td>RS UK</td><td>135-1000</td><td>Custom Stimulator power supply, fully charge before experiment. Not needed if using DS3</td></tr><tr><td>Valve, Leuer Lock, 3-Way</td><td>VWR International Ltd</td><td>229-7440</td><td>For attaching syringe to bath feed tube and priming siphon</td></tr></tbody></table>

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.

Representative results that can be obtained with this protocol are the consistent compound action potentials from A-type nerve fibers within the sciatic nerve. These action potentials typically have a peak-to-peak amplitude of approximately 1 mV at the electrode and therefore 100 mV once amplified (Figure 2). Similar stimulation amplitudes and pulse widths should yield similar CAP amplitudes. Conductive elastomer cuff electrodes will generally require slightly higher stimulation amplitudes i...

Watch this Scientific Journal Video about Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology at JoVE.com

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.

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. ...

preparation-of-rat-sciatic-nerve-for-ex-vivo-neurophysiology

Research

JoVE Journal

Neuroscience

4.2K Views.  Imperial College London. 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.

Video: Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

Axoplasm Isolation from Rat Sciatic Nerve

Isolation of pure axonal cytoplasm (axoplasm) from peripheral nerve is crucial for biochemical studies of many biological processes. In this article, we demonstrate and describe a protocol for axoplasm isolation from adult rat sciatic nerve based on the following steps: (1) dissection of nerve fascicles and separation of connective tissue; (2) incubation of short segments of nerve fascicles in hypotonic medium to release myelin and lyse non-axonal structures; and (3) extraction of the remaining axon-enriched material. Proteomic and biochemical characterization of this preparation has confirmed a high degree of enrichment for axonal components.

We demonstrate a protocol for axoplasm isolation from adult rat sciatic nerve based on dissection of nerve fascicles and incubation in hypotonic medium to release myelin and lyse non-axonal structures, followed by extraction of the remaining axon-enriched material.

Isolation of pure axonal cytoplasm (axoplasm) from peripheral nerve is crucial for biochemical studies of many biological processes. In this article, we ...

Combining Peripheral Nerve Grafting and Matrix Modulation to Repair the Injured Rat Spinal Cord

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of communication with the brain. One of the goals of our research is to promote axon regeneration to restore connectivity across the lesion site. To accomplish this we developed a peripheral nerve (PN) grafting technique where segments of sciatic nerve are either placed directly between the damaged ends of the spinal cord or are used to form a bridge across the lesion. There are several advantages to this approach compared to transplantation of other neural tissues; regenerating axons can be directed towards a specific target area, the number and source of regenerating axons is easily determined by tracing techniques, the graft can be used for electrophysiological experiments to measure functional recovery associated with axons in the graft, and it is possible to use an autologous nerve to reduce the possibility of graft rejection. In our lab we have performed both autologous (donor and recipient are the same animal) and heterologous (donor and recipient are different animals) grafts with comparable results. This approach has been used successfully in both acute and chronic injury situations. Regenerated axons that reach the distal end of the PN graft often fail to extend back into the spinal cord, so we use microinjections of chondroitinase to degrade inhibitory molecules associated with the scar tissue surrounding the area of SCI. At the same time we have found that providing exogenous growth and trophic molecules encourages longer distance axonal regrowth into the spinal cord. Several months after transplantation we perform a variety of anatomical, behavioral and electrophysiological tests to evaluate the recovery of function in our spinal cord injured animals. This experimental approach has been used successfully in several spinal cord injury models, at different levels of injury and in different species (mouse, rat and cat). Importantly, the peripheral nerve grafting approach is effective in promoting regeneration by acute and chronically injured neurons.

Traumatic injury to the spinal cord disrupts communication with the brain. To restore lost connectivity we utilize a peripheral nerve graft to provide a substratum for regenerating fibers in combination with neurotrophic factors and matrix-modulating enzymes to remove inhibitory molecules to promote long distance growth.

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of ...

Biology

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 ...

Toluidine Blue Staining of Resin-Embedded Sections for Evaluation of Peripheral Nerve Morphology

Peripheral nerves extend throughout the body, innervating target tissues with motor or sensory axons. Due to widespread distribution, peripheral nerves are frequently damaged because of trauma or disease. As methods and strategies have been developed to assess peripheral nerve injury in animal models, function and regeneration, analyzing the morphometry of the peripheral nerve has become an essential terminal outcome measurement. Toluidine blue staining of nerve cross sections obtained from resin embedded nerve sections is a reproducible method for qualitative and quantitative assessments of peripheral nerves, enabling visualization of morphology number of axons and degree of myelination. This technique, as with many other histological methods, can be difficult to learn and master using standard written protocols. The intent of this publication is therefore to accentuate written protocols for toluidine blue staining of peripheral nerves with videography of the method, using sciatic nerves harvested from rats. In this protocol, we describe in vivo peripheral nerve fixation and collection of the tissue, and post-fixation with 2% osmium tetroxide, embedding of nerves in epoxy resin, and ultramicrotome sectioning of nerves to 1-2&#956;m thickness. Nerve sections then transferred to a glass slide and stained with toluidine blue, after which they are quantitatively and qualitatively assessed. Examples of the most common problems are shown, as well as steps for mitigating these issues.

Here we present a protocol to visualize fine structures of peripheral nerves by obtaining and staining 1-2 &#181;m sections with toluidine blue

Peripheral nerves extend throughout the body, innervating target tissues with motor or sensory axons. Due to widespread distribution, peripheral nerves ...

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 ...

In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological characteristics in the intact spinal network and holds significant advantages relative to classical in vitro or extracellular recording techniques. An advantage of in vivo intracellular recordings is that this method can be performed on adult animals with a fully mature nervous system, and therefore many observed physiological mechanisms can be translated to practical applications. In this methodological paper, we describe this procedure combined with externally applied constant current stimulation, which mimics polarization processes occurring within spinal neuronal networks. Trans-spinal direct current stimulation (tsDCS) is an innovative method increasingly used as a neuromodulatory intervention in rehabilitation after various neurological injuries as well as in sports. The influence of tsDCS on the nervous system remains poorly understood and the physiological mechanisms behind its actions are largely unknown. The application of the tsDCS simultaneously with intracellular recordings enables us to directly observe changes of motoneuron membrane properties and characteristics of rhythmic firing in response to the polarization of the spinal neuronal network, which is crucial for the understanding of tsDCS actions. Moreover, when the presented protocol includes the identification of the motoneuron with respect to an innervated muscle and its function (flexor versus extensor) as well as the physiological type (fast versus slow) it provides an opportunity to selectively investigate the influence of tsDCS on identified components of spinal circuitry, which seem to be differently affected by polarization. The presented procedure focuses on surgical preparation for intracellular recordings and stimulation with an emphasis on the steps which are necessary to achieve preparation stability and reproducibility of results. The details of the methodology of the anodal or cathodal tsDCS application are discussed while paying attention to practical and safety issues.

This protocol describes in vivo intracellular recording of rat lumbar motoneurons with simultaneous trans-spinal direct current stimulation. The method enables us to measure membrane properties and to record rhythmic firing of motoneurons before, during and after anodal or cathodal polarization of the spinal cord.

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological ...

Automated Gait Analysis to Assess Functional Recovery in Rodents with Peripheral Nerve or Spinal Cord Contusion Injury

Peripheral and central nerve injuries are mostly studied in rodents, especially rats, given the fact that these animal models are both cost-effective and a lot of comparative data has been published in the literature. This includes a multitude of assessment methods to study functional recovery following nerve injury and repair. Besides evaluation of nerve regeneration by means of histology, electrophysiology, and other in vivo and in vitro assessment techniques, functional recovery is the most important criterion to determine the degree of neural regeneration. Automated gait analysis allows recording of a vast quantity of gait-related parameters such as Paw Print Area and Paw Swing Speed as well as measures of inter-limb coordination. Additionally, the method provides digital data of the rats&#8217; paws after neuronal damage and during nerve regeneration, adding to our understanding of how peripheral and central nervous injuries affect their locomotor behavior. Besides the predominantly used sciatic nerve injury model, other models of peripheral nerve injury such as the femoral nerve can be studied by means of this method. In addition to injuries of the peripheral nervous systems, lesions of the central nervous system, e.g., spinal cord contusion can be evaluated. Valid and reproducible data assessment is strongly dependent on meticulous adjustment of the hard- and software settings prior to data acquisition. Additionally, proper training of the experimental animals is of crucial importance. This work aims to illustrate the use of computerized automated gait analysis to assess functional recovery in different animal models of peripheral nerve injury as well as spinal cord contusion injury. It also emphasizes the method&#8217;s limitations, e.g., evaluation of nerve regeneration in rats with sciatic nerve neurotmesis due to limited functional recovery. Therefore, this protocol is thought to help researchers interested in peripheral and central nervous injuries to assess functional recovery in rodent models.

Automated gait analysis is a feasible tool to evaluate functional recovery in rodent models of peripheral nerve injury and spinal cord contusion injury. While it requires only one setup to assess locomotor function in various experimental models, meticulous hard- and soft-ware adjustment and training of the animals is highly important.

Peripheral and central nerve injuries are mostly studied in rodents, especially rats, given the fact that these animal models are both cost-effective and ...

In Vitro Myelination of Peripheral Axons in a Coculture of Rat Dorsal Root Ganglion Explants and Schwann Cells

The process of myelination is essential to enable rapid and sufficient signal transduction in the nervous system. In the peripheral nervous system, neurons and Schwann cells engage in a complex interaction to control the myelination of axons. Disturbances of this interaction and breakdown of the myelin sheath are hallmarks of inflammatory neuropathies and occur secondarily in neurodegenerative disorders. Here, we present a coculture model of dorsal root ganglion explants and Schwann cells, which develops a robust myelination of peripheral axons to investigate the process of myelination in the peripheral nervous system, study axon-Schwann cell interactions, and evaluate the potential effects of therapeutic agents on each cell type separately. Methodologically, dorsal root ganglions of embryonic rats (E13.5) were harvested, dissociated from their surrounding tissue, and cultured as whole explants for 3 days. Schwann cells were isolated from 3-week-old adult rats, and sciatic nerves were enzymatically digested. The resulting Schwann cells were purified by magnetic-activated cell sorting and cultured under neuregulin and forskolin-enriched conditions. After 3 days of dorsal root ganglion explant culture, 30,000 Schwann cells were added to one dorsal root ganglion explant in a medium containing ascorbic acid. The first signs of myelination were detected on day 10 of coculture, through scattered signals for myelin basic protein in immunocytochemical staining. From day 14 onward, myelin sheaths were formed and propagated along the axons. Myelination can be quantified by myelin basic protein staining as a ratio of the myelination area and axon area, to account for the differences in axonal density. This model provides experimental opportunities to study various aspects of peripheral myelination in vitro, which is crucial for understanding the pathology of and possible treatment opportunities for demyelination and neurodegeneration in inflammatory and neurodegenerative diseases of the peripheral nervous system.

In the coculture system of dorsal root ganglia and Schwann cells, myelination of the peripheral nervous system can be studied. This model provides experimental opportunities to observe and quantify peripheral myelination and to study the effects of compounds of interest on the myelin sheath.

The process of myelination is essential to enable rapid and sufficient signal transduction in the nervous system. In the peripheral nervous system, ...

Understanding Motor Neuron Responses to Spinal Cord Stimulation

The Ex vivo Preparation of Spinal Cord Slice for the Whole-Cell Patch-Clamp Recording in Motor Neurons During Spinal Cord Stimulation

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to execute sensorimotor behaviors, directly studying the electrical responses of motor neurons with SCS can help us understand the underlying logic of spinal motor modulation. To simultaneously record diverse stimulus characteristics and cellular responses, a patch-clamp is a good method to study the electrophysiological characteristics at a single-cell scale. However, there are still some complex difficulties in achieving this goal, including maintaining cell viability, quickly separating the spinal cord from the bony structure, and using the SCS to successfully induce action potentials. Here, we present a detailed protocol using patch-clamp to study the electrical responses of motor neurons to SCS with high spatiotemporal resolution, which can help researcher improve their skills in separating the spinal cord and maintaining the cell viability at the same time to smoothly study the electrical mechanism of SCS on motor neuron and avoid unnecessary trial and mistake.

Author Spotlight: Advancing Spinal Cord Stimulation - Exploring the Cellular Responses of Motor Neurons Through Patch-Clamp Electrophysiology 

This protocol describes a method using a patch-clamp to study the electrical responses of motor neurons to spinal cord stimulation (SCS) with high spatiotemporal resolution, which can help researchers improve their skills in separating the spinal cord and maintaining cell viability simultaneously.

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to ...

Trans-Spinal Direct Current Stimulation of a Rat&#39;s Spinal Motoneuron

Source: B&#261;czyk, M., et al. <a href="https://app.jove.com/v/61439">In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation.</a> J. Vis. Exp. (2020).
This video describes the in vivo intracellular recording of rat motoneurons under trans-spinal direct current stimulation. The protocol describes how to measure membrane properties and record the rhythmic firing of motoneurons before, during, and after anodal or cathodal polarization of the spinal cord.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
NOTE: ...

Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

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The Ex vivo Preparation of Spinal Cord Slice for the Whole-Cell Patch-Clamp Recording in Motor Neurons During Spinal Cord Stimulation

Trans-Spinal Direct Current Stimulation of a Rat's Spinal Motoneuron