Name: Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat (Video) | JoVE Journal
Uploaded: 2018-04-23T12:30:00.000+00:00
Duration: 7 min 43 s
Description: Learn cavernous nerve stimulation and intracavernous pressure recording in rats using a simplified surgical technique with silicone glue isolation. This reproducible method enables erectile dysfunction research and therapeutic testing.

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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://scholar.google.com/scholar?hl=en&safe=off&q=author%3aEckhard%2C+C+%22Untersuchungen+%C3%BCber+die+Erektion+des+Hundes+In%3A+Beitr%C3%A4ge+zur+Anatomie+und+Physiologie%22">Untersuchungen über die Erektion des Hundes In: Beiträge zur Anatomie und Physiologie</a>. , Vol. Band III Giessen: Ferber 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/pubmed?term=((The+rat+as+a+model+for+the+study+of+penile+erection%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">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/pubmed?term=((The+characterization+of+a+bio-assay+of+erectile+function+in+a+rat+model%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">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/pubmed?term=((Rat+model+for+the+study+of+penile+erection%3A+pharmacologic+and+electrical-stimulation+parameters%5BTitle%5D)+AND+%22Eur+Urol%22%5BJournal%5D)">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/pubmed?term=((The+effect+of+FK1706+on+erectile+function+following+bilateral+cavernous+nerve+crush+injury+in+a+rat+model%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">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/pubmed?term=((Immunophilin+ligands+promote+penile+neurogenesis+and+erection+recovery+after+cavernous+nerve+injury%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">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/pubmed?term=((Nerve+injury-related+erectile+dysfunction+following+nerve-sparing+radical+prostatectomy%3A+a+novel+experimental+dissection+model%5BTitle%5D)+AND+%22Int+J+Urol%22%5BJournal%5D)">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/pubmed?term=((GGF2+is+neuroprotective+in+a+rat+model+of+cavernous+nerve+injury-induced+erectile+dysfunction%5BTitle%5D)+AND+%22J+Sex+Med%22%5BJournal%5D)">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/pubmed?term=((Nanoparticle+Improved+Stem+Cell+Therapy+for+Erectile+Dysfunction+in+a+Rat+Model+of+Cavernous+Nerve+Injury%5BTitle%5D)+AND+%22J+Urol%22%5BJournal%5D)">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/pubmed?term=((Animal+models+of+erectile+dysfunction%5BTitle%5D)+AND+%22J+Pharmacol+Toxicol+Methods%22%5BJournal%5D)">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/pubmed?term=((Rat+as+an+animal+model+for+male+erectile+function+evaluation+in+sexual+medicine+research%5BTitle%5D)+AND+%22J+Sex+Med%22%5BJournal%5D)">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/pubmed?term=((Common+pitfalls+in+some+of+the+experimental+studies+in+erectile+function+and+dysfunction%3A+a+consensus+article%5BTitle%5D)+AND+%22J+Sex+Med%22%5BJournal%5D)">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/pubmed?term=((Investigative+models+in+erectile+dysfunction%3A+a+state-of-the-art+review+of+current+animal+models%5BTitle%5D)+AND+%22J+Sex+Med%22%5BJournal%5D)">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/pubmed?term=((Functional+sequelae+of+cavernous+nerve+injury+in+the+rat%3A+is+there+model+dependency%5BTitle%5D)+AND+%22J+Sex+Med%22%5BJournal%5D)">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/pubmed?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%5BTitle%5D)+AND+%22J+Sex+Med%22%5BJournal%5D)">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 muscl...

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

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 meas...

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 ...

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Vagus Nerve Stimulation via Chronically Implanted Peripheral Nerve Cuff Electrodes in Rats

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

This video demonstrates the isolation and stimulation of the cavernous nerve in an anesthetized rat to study blood pressure changes in the cavernous body.

Performing Cavernous Nerve Stimulation in Rats to Measure Intracavernous Pressure

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

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

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.

Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents

We developed a technique that simultaneously records both electrocorticography and local field potentials in response to nociceptive laser stimuli from freely moving rats. This technique helps establish a direct relationship of electrocortical signals at the mesoscopic and macroscopic levels, which facilitates the investigation of nociceptive information processing in the brain.

Simultaneous Recordings of Cortical Local Field Potentials and Electrocorticograms in Response to Nociceptive Laser Stimuli from Freely Moving Rats

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.

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

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.

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

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.

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

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.

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

Learn cavernous nerve stimulation and intracavernous pressure recording in rats using a simplified surgical technique with silicone glue isolation. This reproducible method enables erectile dysfunction research and therapeutic testing.

Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat

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