The authors have no acknowledgements.

Animals were housed in the University of Southern Denmark Animal Care Facility as per institutional guidelines. All animal experiments were carried out in accordance with the National Institutes of Health guide for the care and use of laboratory animals. This is an acute, non-survival surgery procedure.&#160;
1. Preparation of Tubing, Electrode, and Instruments for the Surgical Procedure
<ol>
	<li>Use the following microsurgical instruments: surgical scissors, angled micro scissors, a tissue forceps, a pair of Dumont #7 and #5 curved micro forceps, a micro needle holder, and retractors. 
	Note: As this is an acute procedure, the instruments do not need to be sterilized. After use, clean and wipe the tips with 70% ethanol.</li>
	<li>Soak the tubing in 70% ethanol and then flush it with sterile 0.9% NaCl with 100 U/mL heparin before use. Leave the tubing filled to avoid introducing air bubbles into the system.</li>
	<li>Cut a 20-30 cm length piece of polyethylene (PE)-50 tubing to make a catheter for the ICP measurement. Ensure that the tubing is as short as possible to reduce dampening the pressure.</li>
	<li>Bend a sterile 24G needle side-to-side until the needle breaks in the middle. Connect the piece with the bevel to the distal end of the PE tubing for insertion into the penile crus. Insert the other half to the hub for connection to the pressure transducer. Fill the system with heparinized saline (100 U/mL).</li>
	<li>To make the bipolar Teflon coated electrode, cut two 125 &#181;m silver wires to equal length. Use a piece of tape to attach the wires to the edge of the table and thread them together. Subsequently, attach the electrode to a black plate.</li>
	<li>Make a small incision into the Teflon and use #5 micro forceps to strip a 4-5 mm length of Teflon coating off the tips of the electrodes. Cut the tips off with a scalpel to achieve even length and create hooks by bending the ends around the blunt edge of a scalpel blade.</li>
	<li>Tape the electrode with the end extending slightly over the edge of the black plate with the hooks pointing upwards. Mix the silicon glue on a plastic plate for 10 s and wrap a glue bubble around the electrode 1-2 mm from the hook.</li>
	<li>Allow it to dry for approximately 5 min before use (Figure 1). Strip the Teflon from a longer section on the other end, to allow for connection to the stimulator. 
	Note: With re-doing the hooks on the distal end, the electrode can be reused many times.</li>
</ol>
2. Preparation of the Animal
<ol>
	<li>After anesthetizing the animal, shave the lower half of the abdomen, neck, and perineum. Scrub the animal with 70% alcohol followed by povidone-iodine three times. Place the rat on a heated surgical pad in a supine position. Apply vet eye ointment and switch the anesthesia over to a nose cone with 2.5% isoflurane and 0.8 L/min oxygen as the carrier. 
	Note: Adjust the level of isoflurane and oxygen to achieve an acceptable level of anesthesia.</li>
</ol>
3. Presurgical Preparation
<ol>
	<li>Perform the entire surgical procedure under an operating microscope: a magnification ranging from 3.15X to 20X is sufficient. Use gloves and maintain a clean environment throughout the surgery. Place the rat on a drape.</li>
</ol>
4. Ischiocavernosus Muscle Dissection for the ICP Measurement
<ol>
	<li>Use a scalpel, straight scissors, and Dumont #7 curved micro forceps to make a 1 cm vertical skin incision 5 mm lateral to the midline starting at the level of the base of the penis and extending downward (Figure 2A). Use a Q-tip and carefully separate the fascia lateral to the scrotum (Figure 2B). After dissecting the fascia, attach retractors and palpate with a cotton-tipped-swap to find the ischial tuberosity (Figure 2C).</li>
	<li>Dissect through the adipose tissue medial of this point until the ischiocavernosus muscle is visualized (Figure 3A). Use a pair of Dumont #7 curved micro forceps and longitudinally separate the muscle. The tunica albuginea will appear as a bright white structure (Figure 3B). Using micro forceps and a micro scissor, expose the tunica albuginea adequately to see its course (Figure 3C).</li>
	<li>After calibration of the systems settings, insert tubing through the skin on the perineum, making sure that it runs parallel to the penile crus (Figure 4). Leave the line in place and keep the incision moist with saline.</li>
</ol>
5. CN Dissection for Stimulation
<ol>
	<li>Make a 2 cm lower, midline abdominal incision through the skin using, first, a scalpel, and then a pair of straight scissors and micro forceps. Create a matching incision through the fascia along the linea alba and the underlying muscular tissue to expose the bladder and the prostate.</li>
	<li>Use retractors to achieve good exposure. Use cotton-tip swabs to separate the prostate from the adipose tissue to obtain clear visualization of the major pelvic ganglion (MPG) and CN, running on the dorsolateral aspect of the prostate (Figure 5).</li>
	<li>After visualization of the MPG and the CN, carefully incise the fascia overlying the nerve 2-5 mm distal to the MPG with angled micro scissors (Figure 6a). With the use of #5 micro forceps, spread the tissue on each side of the nerve and underneath it to free a 4-mm long portion (Figure 6b), and slide a 9-0 suture under the nerve (Figure 6c).</li>
	<li>Elevate the nerve slightly with the help of the suture (Figure 7a) to facilitate placement of the hooks of the bipolar electrode around the nerve (Figure 7b). Let an assistant mix the two-component silicon glue with the tip of an insulin needle for 5 s. Dry the nerve and apply the glue to the area around the hooks and the nerve (Figure 7c, d). Keep the nerve elevated by pulling slightly on the electrode for approximately 1 min to allow the glue to dry.</li>
	<li>Remove the retractors, except for the retractors on the right side to avoid any pulling or twisting of the electrode. Wet the exposed organs with saline and lay gauze soaked in saline over the incision.</li>
</ol>
6. Cavernous Body Cannulation for the ICP Measurement
<ol>
	<li>Restore visualization of the tunica albuginea using retractors. Make sure not to attach the retractors to the ischiocavernosus muscle as it will distort the crus.</li>
	<li>Attach the needle and flush the tubing with heparinized saline before introducing it into the tunica albuginea. Keep the tunica albuginea stretched, using Dumont #7 curved micro forceps in one hand (non-dominant), holding the tunica albuginea and the rest of the overlying muscle distal to the point of insertion. Hold the needle with straight micro forceps in the other dominant hand and make sure to introduce it parallel with the course of the cavernous body (Figure 8).</li>
	<li>Push the needle 5-8 mm into the cavernous body. Flush the tubing and press on the crus to test the line (Figure 9). Ensure that there are no leaks. Fasten the tubing to the table with tape to prevent accidental pulling on the line. Remove the retractors.</li>
</ol>
7. Stimulation of the CN
<ol>
	<li>With the recording program (e.g., Spike 2) running, continuously record both the intracavernous and mean arterial pressure.</li>
	<li>Set the following parameters on a stimulator (e.g., SD9 Grass Instruments, see Table of Materials) for CN stimulation: current at 1.5 mA, frequency at 16 Hz, voltage at 3 V, and pulse width at 5 ms. Apply 50 s of stimulation with a minimum of 1 min of rest between stimulations. 
	NOTE: The first stimulations usually result in diminished response (Figure 10).</li>
</ol>

<ol>
	<li>Eckhard, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Untersuchungen &#252;ber die Erektion des Hundes In: Beitr&#228;ge zur Anatomie und Physiologie. , 123-166 (1863).</li><li>Quinlan, D. M., Nelson, R. J., Partin, A. W., Mostwin, J. L., Walsh, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+rat+as+a+model+for+the+study+of+penile+erection.">The rat as a model for the study of penile erection.</a> J Urol. 141 (3), 656-661 (1989).</li><li>Heaton, J. P., Varrin, S. J., Morales, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+characterization+of+a+bio-assay+of+erectile+function+in+a+rat+model.">The characterization of a bio-assay of erectile function in a rat model.</a> J Urol. 145 (5), 1099-1102 (1991).</li><li>Martinez-Pineiro, L., 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=Rat+model+for+the+study+of+penile+erection:+pharmacologic+and+electrical-stimulation+parameters.">Rat model for the study of penile erection: pharmacologic and electrical-stimulation parameters.</a> Eur Urol. 25 (1), 62-70 (1994).</li><li>Hayashi, N., 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=The+effect+of+FK1706+on+erectile+function+following+bilateral+cavernous+nerve+crush+injury+in+a+rat+model.">The effect of FK1706 on erectile function following bilateral cavernous nerve crush injury in a rat model.</a> J Urol. 176 (2), 824-829 (2006).</li><li>Burnett, A. L., Becker, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunophilin+ligands+promote+penile+neurogenesis+and+erection+recovery+after+cavernous+nerve+injury.">Immunophilin ligands promote penile neurogenesis and erection recovery after cavernous nerve injury.</a> J Urol. 171 (1), 495-500 (2004).</li><li>Yamashita, 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=Nerve+injury-related+erectile+dysfunction+following+nerve-sparing+radical+prostatectomy:+a+novel+experimental+dissection+model.">Nerve injury-related erectile dysfunction following nerve-sparing radical prostatectomy: a novel experimental dissection model.</a> Int J Urol. 16 (11), 905-911 (2009).</li><li>Burnett, A. L., 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=GGF2+is+neuroprotective+in+a+rat+model+of+cavernous+nerve+injury-induced+erectile+dysfunction.">GGF2 is neuroprotective in a rat model of cavernous nerve injury-induced erectile dysfunction.</a> J Sex Med. 12 (4), 897-905 (2015).</li><li>Lin, H., 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=Nanoparticle+Improved+Stem+Cell+Therapy+for+Erectile+Dysfunction+in+a+Rat+Model+of+Cavernous+Nerve+Injury.">Nanoparticle Improved Stem Cell Therapy for Erectile Dysfunction in a Rat Model of Cavernous Nerve Injury.</a> J Urol. 195 (3), 788-795 (2016).</li><li>Kapoor, M. S., Khan, S. A., Gupta, S. K., Choudhary, R., Bodakhe, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+erectile+dysfunction.">Animal models of erectile dysfunction.</a> J Pharmacol Toxicol Methods. 76, 43-54 (2015).</li><li>Mehta, N., Sikka, S., Rajasekaran, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+as+an+animal+model+for+male+erectile+function+evaluation+in+sexual+medicine+research.">Rat as an animal model for male erectile function evaluation in sexual medicine research.</a> J Sex Med. 5 (6), 1278-1283 (2008).</li><li>Cellek, S., Bivalacqua, T. J., Burnett, A. L., Chitaley, K., Lin, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Common+pitfalls+in+some+of+the+experimental+studies+in+erectile+function+and+dysfunction:+a+consensus+article.">Common pitfalls in some of the experimental studies in erectile function and dysfunction: a consensus article.</a> J Sex Med. 9 (11), 2770-2784 (2012).</li><li>Chung, E., De Young, L., Brock, G. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Investigative+models+in+erectile+dysfunction:+a+state-of-the-art+review+of+current+animal+models.">Investigative models in erectile dysfunction: a state-of-the-art review of current animal models.</a> J Sex Med. 8 (12), 3291-3305 (2011).</li><li>Mullerad, M., Donohue, J. F., Li, P. S., Scardino, P. T., Mulhall, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+sequelae+of+cavernous+nerve+injury+in+the+rat:+is+there+model+dependency.">Functional sequelae of cavernous nerve injury in the rat: is there model dependency.</a> J Sex Med. 3 (1), 77-83 (2006).</li><li>Albersen, M., 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=Injections+of+adipose+tissue-derived+stem+cells+and+stem+cell+lysate+improve+recovery+of+erectile+function+in+a+rat+model+of+cavernous+nerve+injury.">Injections of adipose tissue-derived stem cells and stem cell lysate improve recovery of erectile function in a rat model of cavernous nerve injury.</a> J Sex Med. 7 (10), 3331-3340 (2010).</li></ol>

The authors have nothing to disclose.

The principal goal of this study was to describe a simplified surgical technique of penile crus cannulation for ICP recording and isolation of the CN for electrostimulation. We introduced modifications to the dissection of the cavernous body to simplify the surgery and provide reproducible recordings of the increase of the ICP with CN stimulation. With a 1 cm vertical skin incision, lateral to the base of the penis, using the palpable ischial tuberosity as guidance, we achieved good exposure of the ischiocavernosus muscle and tunica albuginea. This procedure is quicker (under 15 min) than those described in the literature and causes minimal tissue disruption2,3,4.
The currently used technique of CN stimulation includes hooking, lifting, and drying of the CN before each electrostimulation is applied10. This technique does not guarantee that the conditions remain the same for each stimulation. Also, lifting of the nerve requires the use of a micromanipulator and repeated stretching and releasing of the nerve, which could lead to neuropraxia. Compared to the current CN stimulation technique, the use of a biocompatible silicon glue to isolate the nerve-electrode complex reduces the frequency of the nerve stretching to two instances; once to place the bipolar electrode around the nerve, and once to isolate the nerve and the electrode using glue. Subsequently, multiple neurostimulations could be performed without the need for nerve manipulations.
Insertion of the needle into the cavernous body for pressure recording constitutes a critical step. New users of this technique should practice this part before starting experiments. When deploying the needle into the crus, it is critical that the crus is stretched and the needle is deployed parallel to its course. The other critical step is the nerve handling during its dissection from the surrounding tissue and placement of the electrode underneath. While dissection itself is not a difficult task, any stretch or crush of the nerve could result in damage. Our modification of this step makes the need for a micromanipulator unnecessary and simplifies nerve handling. The use of the biocompatible silicon glue to isolate the nerve and provide stable and reliable contact between the electrode and the nerve reduces the necessary manipulation. Silicone glue could also be used in other animal models where in vivo neurostimulation is applied.
The stimulation parameters listed in numerous studies vary from 14-20 Hz and 1.5-12 V11,13,14. This technique showed that stimulation using a 1.5 mA, 16 Hz, 3 V, and 5 ms pulse produced a full response. With increasing stimulation parameters, the ICP does not increase (Figure 18). This suggests that stimulation using any parameters, which are above the threshold for triggering arterial (cavernous artery), arteriolar, and sinusoidal smooth muscle relaxation, are sufficient to trigger the reflex and would result in a full physiological response. The subsequent increase in the frequency, amplitude, or pulse width does not lead to a stronger physiological response and it could deplete neurotransmitters or even cause nerve injury. With the above-described parameters, we were able to achieve a reproducible response with a rest period as short as 30 s more than 40 times in a row.
The depth of anesthesia clearly affects the physiological response. With inhalation anesthesia, it could be well controlled, however the peak response is approximately 25% lower, when compared to fentanyl/midazolam anesthesia administered by injection. We observed that the dose of isoflurane could be maintained at the level determined by the minimum isoflurane concentration necessary to eliminate toe pinch response. Most investigators, use injection anesthesia with sodium pentobarbital, fentanyl/midazolam, ketamine/xylazine, or ketamine/midazolam15.
None of the published articles mentioned the use of controlled oxygenation. Results from this study show that this had a major impact on ICP. Therefore, it is important that, regardless of type of anesthesia used, the animal receives oxygen through the nose cone.
The use of rats is advantageous due to their size and resilience. Both in previously published studies and confirmed in this research, Sprague Dawley rats have been shown to have an intact response to CN stimulation in the range of 6-12 weeks old and weighing 200-550 g. Using mice is more challenging, but beneficial due to the availability of transgenic technology12.

In vivo study of erectile function in experimental animals started in 1863 with the pioneering experimental work of Eckhard1. Electrostimulation of the pelvic nerves was used to induce the increased ICP in dogs. Throughout the 20th century, similar experimental protocols were used in larger animals such as dogs, monkeys, cats and rabbits. Evaluating erectile function in a rat was first developed by Quinlan et al. in 19892. The method has since been modified and updated by several other groups34. Today, the rat is the most widely used animal model for studying the pathology of erectile dysfunction and evaluating emerging treatment options. The main steps of the procedure include, recording systemic blood pressure using a line in the carotid artery, cannulation of the penile crus to measure ICP, and stimulation of the CN to induce an increase in the ICP. Although several researchers have refined the model, its reproducibility remains a problem, and variable results have been reported by different laboratories. Several pitfalls still persist.
Previous articles5,6,7,8,9,10 describe the use of full penile exposure with degloving of the penis for cavernous body cannulation. This is not an optimal approach as manipulation and disruptive dissection causes injury to structures, which are essential to erectile function. The dissection of the CN has been well described10,11, but stimulation of the nerve is not optimal due to multiple factors that could affect experimental results. The technique of CN stimulation includes lifting the nerve from the surrounding tissue by pulling on the bipolar hook electrode, which is positioned around the nerve, and drying the nerve before each stimulation. This can lead to various degrees of nerve damage and electrical current leakage, resulting in a diminished response or false increase in the ICP through stimulation of alternative pathways e.g., pelvic floor muscles, bladder and gastrointestinal tract12. All of these factors limit reproducibility.
During our study, we observed that both the depth and type of anesthesia have a profound effect on the ICP. The anesthetics used are sodium pentobarbital, ketamine/xylazine or ketamine/midazolam injection, or isoflurane/oxygen inhalation.
Here we describe a simplified surgical method and provide data in support of standardization of the experimental protocol.

<table><tbody><tr><td>Adson forceps</td><td>Fine Science Tool</td><td>11006-12</td><td></td></tr><tr><td>Dumont #7 forceps</td><td>Fine Science Tool</td><td>11271-30</td><td></td></tr><tr><td>Dumont #5 forceps</td><td>Fine Science Tool</td><td>11273-20</td><td></td></tr><tr><td>ToughCut Mayo scissor</td><td>Fine Science Tool</td><td>14110-15</td><td></td></tr><tr><td>Miniature Vannas Spring scissor</td><td>Fine Science Tool</td><td>15006-09</td><td></td></tr><tr><td>Ultra Fine Hemostat</td><td>Fine Science Tool</td><td>13020-12</td><td></td></tr><tr><td>Crile Hemostat</td><td>Fine Science Tool</td><td>13004-14</td><td></td></tr><tr><td>Kwik-Sil Adhesive</td><td>World Precision Instruments</td><td>KWIK-SIL</td><td></td></tr><tr><td>Teflon coated silver wire 0.125 mm</td><td>World Precision Instruments</td><td>AGT0510</td><td></td></tr><tr><td>Elastic wire retractors</td><td>Custom made</td><td></td><td></td></tr><tr><td>Scalpel blade</td><td>Fine Science Tool</td><td>10023-00</td><td></td></tr><tr><td>PE-50 tubing</td><td>Warner Instruments</td><td>64-0753</td><td></td></tr><tr><td>23 G Needle</td><td>Kruuse</td><td>121272</td><td></td></tr><tr><td>SD-9 Square Pulse Stimulator</td><td>Somatco</td><td>1077/183</td><td></td></tr><tr><td>Blood pressure transducer and cable</td><td>World Precision Instruments</td><td>BLPR2</td><td></td></tr><tr><td>Raucotupf Cotton-tipped Applicators</td><td>Lohmann-Raucher</td><td>11966</td><td></td></tr><tr><td>Pro-ophta Ocular Sticks</td><td>Lohmann-Raucher</td><td>16515</td><td></td></tr><tr><td>NaCl 0,9 % 100 mL</td><td>Local pharmacy</td><td></td><td></td></tr><tr><td>Heparin</td><td>Local pharmacy</td><td></td><td></td></tr><tr><td>25 mL Syringe</td><td>Odense University Hospital</td><td></td><td></td></tr><tr><td>Vet eye ointment - Viscotears</td><td>Local pharmacy</td><td></td><td></td></tr><tr><td>silver wires&amp;nbsp;</td><td>Science Products GmbH, Heidelberg, Germany</td><td></td><td></td></tr><tr><td>Silicon Glue&amp;nbsp;</td><td>Kwik-Sil, World Precision Instruments, Sarasota, FL, USA</td><td></td><td></td></tr></tbody></table>

cavernous nerve stimulation and recording of intracavernous pressure in a rat

The stimulation of the cavernous nerve (CN) and measurement of intracavernous pressure (ICP) have been used extensively to test and evaluate therapies for erectile dysfunction. However, the methods used vary between laboratories, and pitfalls still exist. The goal of this study was to describe a surgical technique that would provide a reliable and reproducible model. By exposing the ischiocavernosus muscle at its point of insertion on the ischial tuberosity, the penile crus could be cannulated with minimal dissection and injury to the structures involved in erectile function. Repeated stimulation of the CN, without the need for lifting and drying, was achieved by using a 125 &#181;m bipolar silver electrode and biocompatible silicon glue to isolate the electrode-nerve complex. This method prevents neuropraxia by reducing stretching and drying the nerve and provides complete isolation of the nerve, negating electrical leakage and preventing stimulation of alternative pathways.

The use of this protocol with the recommended stimulation settings, under inhalation anesthesia with isoflurane 2.0% oxygen 0.8 L/min, should produce results as shown in Figure 11 and Figure12, where there is multiple back to back stimulations between 75-80 mm Hg. Figure 13 shows the same stable response over a 20-min stimulation with the response stable at 73-77 mm Hg. Test the line for ICP measurement by flushing the tube and tapping on the crus (Figure 9). The rapid response back to baseline is the hallmark of a well-placed line. If the integrity of the tunica albuginea is damaged, the test would result in lower peak pressure and slow response back to baseline after flushing and tapping, and a diminished response when stimulating (Figure 14). There would also be leakage of the heparinized NaCl when flushing and bleeding during stimulations.
The types and levels of anesthesia as well as the use of oxygen had a major impact on ICP. Figure 15 demonstrates the effect of the different levels of isoflurane on ICP, with both a decreased response and a less stable plateau. With isoflurane at 2%, there was a stable response in the ICP measurement with multiple stimulations on 78 mm Hg. Increasing the concentration of isoflurane to 3.5%, however, resulted in a rapid 50% decrease to 34 mm Hg in multiple, subsequent stimulations. The same effect was observed when switching the isoflurane from 2.0% to 3.0%, where a 19% decrease in response was observed, and from 2.5% to 5%, where an even more rapid decrease in response of 70% was seen. Blood pressure remained stable throughout all stimulations. In rats anesthetized using the isoflurane/oxygen anesthesia during surgery and initial stimulations, who then received 25% of the recommended dose of fentanyl/midazolam (while the isoflurane was discontinued), there was a similarly stable response but it increased by 25% during the fentanyl/midazolam anesthesia compared to isoflurane (Figure 16).
Administration of oxygen through a nose cone increased the oxygen saturation in the blood from 61-75% to 99-100% in about 20 s. When the oxygenation was stopped, the same decrease was seen over approximately 1 min. Blood pressure was stable throughout the stimulations, but oxygen administration through a nose cone (0.8 L/min) had a large effect on the maximum ICP measurement, reducing it by 35-45% in back-to-back stimulations (Figure 17).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56807/56807fig1.jpg" /> 
Figure 1. The bipolar Teflon coated silver electrode. (A) Glue bubble in the transition zone between the coated and uncoated electrode. (B) Distal 2 cm of the electrode tightly braded. (C) Parallel uncoated hooks 1 mm apart. <a href="/files/ftp_upload/56807/56807fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56807/56807fig2.jpg" /> 
Figure 2. Dissection of the cavernous body - landmarks. (A) 1 cm vertical skin incision, downward starting 2 mm lateral from the base of the penis. (B) Fascia lateral to the scrotum separated using cotton-tipped swabs. (C) View of the operating field after placement of retractors (pm: pyramidalis muscle, it: insertion point of crus to the ischial tuberosity). <a href="/files/ftp_upload/56807/56807fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56807/56807fig3.jpg" /> 
Figure 3. Exposing the tunica albuginea. (A) The ischiocavernosus muscle (arrow). (B) Tunica albuginea (arrow). (C) Low power magnification showing the course of the cavernous body. <a href="/files/ftp_upload/56807/56807fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56807/56807fig4.jpg" /> 
Figure 4. Line for cavernous pressure recording. Needle introduced through the skin on the perineum running parallel with cavernous body (arrow). <a href="/files/ftp_upload/56807/56807fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56807/56807fig5.jpg" /> 
Figure 5. Exposure of the MPG marked by arrow and CN running vertical on the dorsolateral aspect of the prostate. Major pelvic ganglion (MPG) marked by arrow, cavernous nerve (CN). <a href="/files/ftp_upload/56807/56807fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56807/56807fig6.jpg" /> 
Figure 6. Dissection of the cavernous nerve. (A) Cutting the fascia overlying the cavernous nerve with micro scissors. (B) Separating the nerve from the underlying tissue using micro forceps. (C) Placing a 9-0 ligature underneath the nerve. Major pelvic ganglion (MPG) marked by arrow, cavernous nerve (CN). <a href="/files/ftp_upload/56807/56807fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56807/56807fig7.jpg" /> 
Figure 7. Hooking of the nerve. (A) Elevating the nerve by gently pulling on the suture. (B) Nerve resting in the hooks of the electrode. (C) Nerve and electrode complex isolated with the biocompatible silicon glue. (D) Additional glue bubble added to completely isolate the nerve-electrode complex. <a href="/files/ftp_upload/56807/56807fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/56807/56807fig8.jpg" /> 
Figure 8. Cannulation of the tunica albuginea. A 23 G needle connected to PE-50 tubing inserted into the tunica albuginea. Point of insertion marked by arrow. <a href="/files/ftp_upload/56807/56807fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/56807/56807fig9.jpg" /> 
Figure 9. Testing the intracavernous line. The responses seen with a correct line placement. Note the flushing of the line and response to tapping on the crus. Also, note the quick pressure drop back to baseline. <a href="/files/ftp_upload/56807/56807fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/56807/56807fig10.jpg" /> 
Figure 10. Initial responses to cavernous nerve stimulation. Diminished first response. First stimulation of 50 mm Hg and a fluctuating plateau. Second and third stimulation of 66 mm Hg. The following measurements were recorded at the normal level at 73 mm Hg. <a href="/files/ftp_upload/56807/56807fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/56807/56807fig11.jpg" /> 
Figure 11. Repeated cavernous nerve stimulation and intracavernous pressure recording. Showing the stability of the results using this protocol. Ten back-to-back stimulations between 75-78 mm Hg. <a href="/files/ftp_upload/56807/56807fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" class="xfigimg" src="/files/ftp_upload/56807/56807fig12.jpg" /> 
Figure 12. Cavernous nerve stimulation and pressure recording. Approximately 30 back-to-back stimulations with &#60; 6 mm Hg variability in the pressure. <a href="/files/ftp_upload/56807/56807fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 13" class="xfigimg" src="/files/ftp_upload/56807/56807fig13.jpg" /> 
Figure 13. Continuous stimulation lasting 20 min. Increased fluctuation at the end, but the subsequent stimulations, after a 1 min rest, produced a stable response. <a href="/files/ftp_upload/56807/56807fig13large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 14" class="xfigimg" src="/files/ftp_upload/56807/56807fig14.jpg" /> 
Figure 14. Leaking tunica albuginea. Prolonged response back to baseline after flushing and tapping. Decreased response after stimulation. <a href="/files/ftp_upload/56807/56807fig14large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 15" class="xfigimg" src="/files/ftp_upload/56807/56807fig15.jpg" /> 
Figure 15. The effect of anesthesia dose on intracavernous pressure. Decreased and more fluctuating response on increasing isoflurane compared to the stable response on 2% in the first two, and the last three stimulations. The blue trace on top showing constant mean arterial pressure. <a href="/files/ftp_upload/56807/56807fig15large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 16" class="xfigimg" src="/files/ftp_upload/56807/56807fig16.jpg" /> 
Figure 16. The effect of anesthesia type on intracavernous pressure. The initial stimulations performed under isoflurane anesthesia show a pressure increase of 80 mm Hg, once fentanyl/midazolam was administered there was an increase in response to 110 mm Hg. <a href="/files/ftp_upload/56807/56807fig16large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 17" class="xfigimg" src="/files/ftp_upload/56807/56807fig17.jpg" /> 
Figure 17. The effect of oxygen administration through the nose cone. Discontinuing oxygen administration through the nose cone resulted in a significant reduction in cavernous pressure increase with cavernous nerve stimulation. No effect on the mean arterial pressure was noted (blue trace above). <a href="/files/ftp_upload/56807/56807fig17large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 18" class="xfigimg" src="/files/ftp_upload/56807/56807fig18.jpg" /> 
Figure 18. The effect of voltage on pressure response to stimulation. The voltage between 1.5-6 V produced an identical pressure response. The response diminished below 1.5 V. <a href="/files/ftp_upload/56807/56807fig18large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat at JoVE.com

Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat

This study describes a simplified surgical procedure and technique for performing cavernous nerve stimulation with the isolation of the nerve-electrode complex using silicone glue and intracavernous pressure measurement.

The stimulation of the cavernous nerve (CN) and measurement of intracavernous pressure (ICP) have been used extensively to test and evaluate therapies for ...

cavernous-nerve-stimulation-recording-intracavernous-pressure

Nanyang Technological University

Research

JoVE Journal

Medicine

12.5K Views.  University of Southern Denmark. Deze studie beschrijft een vereenvoudigde chirurgische ingreep en techniek voor het uitvoeren van cavernous zenuwstimulatie met het isolement van de zenuw-elektrode complex met siliconen lijm en meting van de intracavernous druk.

Video: Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat

Epidural Intracranial Pressure Measurement in Rats Using a Fiber-optic Pressure Transducer

Elevated intracranial pressure (ICP) is a significant problem in several forms of ischemic brain injury including stroke, traumatic brain injury and cardiac arrest. This elevation may result in further neurological injury, in the form of transtentorial herniation1,2,3,4, midbrain compression, neurological deficit or increased cerebral infarct2,4. Current therapies are often inadequate to control elevated ICP in the clinical setting5,6,7 . Thus there is a need for accurate methods of ICP measurement in animal models to further our understanding of the basic mechanisms and to develop new treatments for elevated ICP.
In both the clinical and experimental setting ICP cannot be estimated without direct measurement. Several methods of ICP catheter insertion currently exist. Of these the intraventricular catheter has become the clinical 'gold standard' of ICP measurement in humans8. This method involves the partial removal of skull and the instrumentation of the catheter through brain tissue. Consequently, intraventricular catheters have an infection rate of 6-11%9. For this reason, subdural and epidural cannulations have become the preferred methods in animal models of ischemic injury. 
Various ICP measurement techniques have been adapted for animal models, and of these, fluid-filled telemetry catheters10 and solid state catheters are the most frequently used11,12,13,14,15. The fluid-filled systems are prone to developing air bubbles in the line, resulting in false ICP readings. Solid state probes avoid this problem (Figure 1). An additional problem is fitting catheters under the skull or into the ventricles without causing any brain injury that might alter the experimental outcomes. Therefore, we have developed a method that places an ICP catheter contiguous with the epidural space, but avoids the need to insert it between skull and brain. 
An optic fibre pressure catheter (420LP, SAMBA Sensors, Sweden) was used to measure ICP at the epidural location because the location of the pressure sensor (at the very tip of the catheter) was found to produce a high fidelity ICP signal in this model. There are other manufacturers of similar optic fibre technologies13 that may be used with our methodology. Alternative solid state catheters, which have the pressure sensor located at the side of the catheter tip, would not be appropriate for this model as the signal would be dampened by the presence of the monitoring screw. 
Here, we present a relatively simple and accurate method to measure ICP. This method can be used across a wide range of ICP related animal models.

A novel technique to record the pressures within the skull is described. The minimally invasive method uses a fibre-optic pressure sensing system to accurately measure intracranial pressure (ICP) in anaesthetized rats without causing significant brain trauma. The technique may be used in a wide range of experimental models.

Elevated intracranial pressure (ICP) is a significant problem in several forms of ischemic brain injury including stroke, traumatic brain injury and ...

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer scale. However, current methods mostly rely on tethered cable recordings or only use unidirectional systems, allowing either recording or stimulation of neural activity, but not at the same time or same target. Here, a new wireless, bidirectional device for simultaneous multichannel recording and stimulation of neural activity in freely behaving rats is described. The system operates through a single portable head stage that both transmits recorded activity and can be targeted in real-time for brain stimulation using a telemetry-based multichannel software. The head stage is equipped with a preamplifier and a rechargeable battery, allowing stable long-term recordings or stimulation for up to 1 h. Importantly, the head stage is compact, weighs 12 g (including battery) and thus has minimal impact on the animal&#180;s behavioral repertoire, making the method applicable to a broad set of behavioral tasks. Moreover, the method has the major advantage that the effect of brain stimulation on neural activity and behavior can be measured simultaneously, providing a tool to assess the causal relationships between specific brain activation patterns and behavior. This feature makes the method particularly valuable for the field of deep brain stimulation, allowing precise assessment, monitoring, and adjustment of stimulation parameters during long-term behavioral experiments. The applicability of the system has been validated using the inferior colliculus as a model structure.

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

A wireless, bidirectional system for multi-channel neural recordings and stimulation in freely behaving rats is introduced. The system is light and compact, thus having minimal impact on the animal&#180;s behavioral repertoire. Moreover, this bidirectional system provides a sophisticated tool to assess causal relationships between brain activation patterns and behavior.

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer ...

Behavior

Novel Approach for Simultaneous Recording of Renal Sympathetic Nerve Activity and Blood Pressure with Intravenous Infusion in Conscious, Unrestrained Mice.

Renal sympathetic nerves contribute significantly to both physiological and pathophysiological phenomena. Evaluating renal sympathetic nerve activity (RSNA) is of great interest in many areas of research such as chronic kidney disease, hypertension, heart failure, diabetes and obesity. Unequivocal assessment of the role of the sympathetic nervous system is thus imperative for proper interpretation of experimental results and understanding of disease processes. RSNA has been traditionally measured in anesthetized rodents, including mice. However, mice usually exhibit very low systemic blood pressure and hemodynamic instability for several hours during anesthesia and surgery. Meaningful interpretation of RSNA is confounded by this non-physiological state, given the intimate relationship between sympathetic nervous tone and cardiovascular status. To address this limitation of traditional approaches, we developed a new method for measuring RSNA in conscious, freely-moving mice. Mice were chronically instrumented with radio-telemeters for continuous monitoring of blood pressure as well as a jugular venous infusion catheter and custom-designed bipolar electrode for direct recording of RSNA. Following a 48-72 hour recovery period, survival rate was 100% and all mice behaved normally. At this time-point, RSNA was successfully recorded in 80% of mice, with viable signals acquired up to 4 and 5 days post-surgery in 70% and 50% of mice, respectively. Physiological blood pressures were recorded in all mice (116&plusmn;2 mmHg; n=10). Recorded RSNA increased with eating and grooming, as well-established in the literature. Furthermore, RSNA was validated by ganglionic blockade and modulation of blood pressure with pharmacological agents. Herein, an effective and manageable method for clear recording of RSNA in conscious, freely-moving mice is described.

Anesthetized mice exhibit non-physiological systemic blood pressure, which precludes meaningful assessment of autonomic tone given the intimate relationship between blood pressure and the autonomic nervous system. Thus, a novel method to simultaneously record renal sympathetic nerve activity and blood pressure with intravenous infusion in conscious mice is outlined.

Renal sympathetic nerves contribute significantly to both physiological and pathophysiological phenomena. Evaluating renal sympathetic nerve activity ...

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. Several recent studies have demonstrated the effectiveness of chronic VNS in enhancing central nervous system plasticity to improve motor rehabilitation, extinction learning, and sensory discrimination. Construction of chronically implantable devices for use in such studies is challenging due to rats&#8217; small size, and typical protocols require extensive training of personnel and time-consuming microfabrication methods. Alternatively, commercially available implantable cuff electrodes can be purchased at a significantly higher cost. In this protocol, we present a simple, low-cost method for construction of small, chronically implantable peripheral nerve cuff electrodes for use in rats. We validate the short and long-term reliability of our cuff electrodes by demonstrating that VNS in ketamine/xylazine anesthetized rats produces decreases in breathing rate consistent with activation of the Hering-Breuer reflex, both at the time of implantation and up to 10 weeks after device implantation. We further demonstrate the suitability of the cuff electrodes for use in chronic stimulation studies by pairing VNS with skilled lever press performance to induce motor cortical map plasticity.

Existing approaches for constructing chronically implantable peripheral nerve cuff electrodes for use in small rodents often require specialized equipment and/or highly trained personnel. In this protocol we demonstrate a simple, low-cost approach for fabricating chronically implantable cuff electrodes, and demonstrate their effectiveness for vagus nerve stimulation (VNS) in rats.

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. ...

Neuroscience

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 ...

Abdominal VNS for Treating Inflammatory Conditions in Rat Models

Implantation Surgery for Abdominal Vagus Nerve Stimulation and Recording Studies in Awake Rats

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does not have any respiratory and cardiac off-target effects commonly associated with cervical VNS. The lack of respiratory and cardiac off-target effects means that the intensity of stimulation does not need to be lowered to reduce side effects commonly experienced during cervical VNS. Few recent studies demonstrate the anti-inflammatory effects of abdominal VNS in rat models of inflammatory bowel disease, rheumatoid arthritis, and glycemia reduction in a rat model of type 2 diabetes. Rat is a great model to explore the potential of this technology because of the well-established anatomy of the vagus nerve, the large size of the nerve that allows easy handling, and the availability of many disease models. Here, we describe the methods for cleaning and sterilizing the abdominal VNS electrode array and surgical protocol in rats. We also describe the technology required for confirmation of suprathreshold stimulation by recording evoked compound action potentials. Abdominal VNS has the potential to offer selective, effective treatment for a variety of conditions, including inflammatory diseases, and the application is expected to expand similarly to cervical VNS.

Author Spotlight: Exploring Abdominal VNS for Inflammatory Conditions

The present protocol describes the surgical technique for implanting an electrode array onto the abdominal vagus nerve in rats, along with methods for chronic electrophysiology testing and stimulation using the implanted device.

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does ...

Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents

Erectile dysfunction (ED) is defined as the inability to attain or keep an erection of the penis, and this has become a prevalent male sexual disorder. Rodents are employed by many studies to research the physiology/pathology of erectile function. Erectile function in rodents can be evaluated by measuring the intracavernosal pressure (ICP). In practice, ICP can be monitored following electrical stimulation of the cavernous nerves (CNs). The arterial pressure of the carotid artery (the mean arterial pressure) is used as the reference for ICP. Using ICP recording protocols, many key parameters of erectile function can be measured from the ICP response curve. The ICP measurement provides more information than the apomorphine-induced penile erection test, and is cheaper than telemetric monitoring of the corpus spongiosum penis, making this method the most popular one to evaluate erectile function. However, compared to the easily-performed APO-induced erectile function test, successful ICP recordings require attention to detail, practice, and adherence to the operation method. In this work, an introduction to ICP recording in rats is provided to complement the procedure efficiently.

Intracavernosal pressure recording (ICP) is an important method to evaluate the erectile function of experimental animals. Here, a detailed protocol is demonstrated for the recording procedure of ICP by catheterizing the crura penis and then electrically stimulating the cavernous nerves in rats.

Erectile dysfunction (ED) is defined as the inability to attain or keep an erection of the penis, and this has become a prevalent male sexual disorder. ...

Biology

Intracavernous Pressure Recording in a Cavernous Nerve Injury Rat Model

The bilateral cavernous nerve (CN) injury rat model has been extensively used to simulate clinical cavernous nerve injury associated with erectile dysfunction (ED) for evaluating the effect of clinical therapeutic methods. However, the methods of CN injury model construction are flawed and varied in the ED research field. It is CN crush injury that is the most commonly used method in recent years. This study aims to provide a detailed description of the procedure of bilateral CN injury rat model construction and measurement of intracavernous pressure (ICP) recording, providing a reliable and reproducible CN injury rat model. This work successfully developed the CN injury method of hemostat crush injury using a syringe needle as hard support and a hemostat with a rubber sleeve. Also, this method concludes that a voltage of 1.0 V, frequency of 20 Hz, and pulse-width of 5 ms are the optimized stimulation parameters for ICP recording in a bilateral CN injury rat model.

This protocol describes developing a stable bilateral cavernous nerve injury rat model of radical prostatectomy associated with erectile dysfunction and intracavernous pressure measurement.

The bilateral cavernous nerve (CN) injury rat model has been extensively used to simulate clinical cavernous nerve injury associated with erectile ...

Performing Cavernous Nerve Stimulation in Rats to Measure Intracavernous Pressure

Source: Hox, M., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/56807">Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat.</a> J. Vis. Exp. (2018).
This video demonstrates the isolation and stimulation of the cavernous nerve in an anesthetized rat to study blood pressure changes in the cavernous body.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

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

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

Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat

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