Name: intracavernosal pressure recording to evaluate erectile function in rodents
Uploaded: 2018-06-06T12:30:00.000+00:00
Duration: 8 min 3 s
Description: Watch this Scientific Journal Video about Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents at JoVE.com

This work was supported by the Fundamental Research Funds for the Central Universities (020814380018, 020814380077), The China Scholarship Council (CSC, No. 201606195024), Natural Science Foundation of Jiangsu Province (BK20160138), and Key Project supported by Science and Technology Development Foundation, Nanjing Medical University (2014NJMUZD053).

Three-month-old and 18-month-old Sprague-Dawley rats were used in the present study. All animals were handled in accordance with the NIH Guidelines for the Care and Use of Laboratory Animals. Procedures involving animal subjects were approved by the local Institutional Animal Care and Ethics Committee, with an effort to minimize animal suffering. The protocols were approved by the Institutional Animal Care and Use Committee (IACUC) at Nanjing Tech University (Nanjing, China).
The rats were divided into two groups according to their age and preliminary performance in the APO-induced erectile function test prior to ICP recording: the young normal group (YN group) and aged erectile dysfunction group (AE group)10.
1. Preparation Before Surgery
<ol>
	<li>Manually make a pair of bipolar electrodes for the ICP recording (Figure 1). Slightly bend the ends of electrodes, and adjust the distance between two electrodes to 1-2 mm wide, as shown in Figure 1A.</li>
	<li>Connect the electrodes to the stimulator using two crocodile clamps (Figure 1A,B).</li>
	<li>Assemble the catheter system: First, connect a hypodermic 23G needle to a 3-way stopcock with tubing, then connect the stopcock to the pressure transducer. Next, attach a 10-mL syringe to the third end of stopcock to provide heparin saline.</li>
	<li>Carefully check for leakage after filling the whole system with heparin saline (200 U/mL). Then turn the 3-way stopcock to close the syringe channel or the pressure transducer channel (Figure 1C).</li>
	<li>Lift the needle 20 cm up the level of the wooden pad. Then calibrate the pressure recording system to 20 cm H2O. After that, move the height of the needle to verify the accuracy of the recording system. Repeat the calibration until the accuracy was confirmed.</li>
	<li>Transfer the rats from the animal facility to the surgery room, and allow them to become accustomed to the surgery room for at least 30 min.</li>
	<li>The autoclaved instruments are sprayed with 70% ethanol just before surgery</li>
</ol>
2. Surgery Procedure
<ol>
	<li>Anesthetize the rat with an intraperitoneal injection of sodium pentobarbital at a dose of 45 mg/kg body weight, and wait for 5-10 min. Pinch the toes to confirm a proper anesthetization.</li>
	<li>Shave the fur of the abdomen and neck with an electric shaver, and place the rat on its back on a heating pad.</li>
	<li>Wipe the surgery area with 10% povidone-iodine solution soaked cotton balls followed by 70% ethanol soaked cotton balls. Also, apply ophthalmic ointment to prevent the eyes from drying out.</li>
	<li>Catheterize the left carotid artery.
	<ol>
		<li>Grasp the skin of the neck with forceps and make a horizontal incision in the middle of the neck. Incise the muscles, carefully expose the left carotid artery, and isolate a 5-mm section of the vessel.</li>
		<li>Carefully separate the carotid artery from the vagus nerve using forceps, draw a silk suture under the carotid artery, and put a loose tie on the caudal end of the vessel, then make another tight knot on the cranial end of the vessel.</li>
		<li>Caudally clamp the vessel with a bulldog clamp above the suture to stop the blood flow.</li>
		<li>Carefully make an incision on the vessel with microsurgical scissors, and insert the arterial catheter towards the heart with the assistance of the micro-dissecting hook and forceps.</li>
		<li>Fasten the loose caudal ligature around the catheter to secure it. Remove the bulldog clamp to recover the blood flow.</li>
	</ol>
	</li>
	<li>Isolate the CN and Place the Electrode
	<ol>
		<li>Lift the skin and muscle of the abdomen with a pair of forceps. With the dissecting scissors, cut through the lower abdomen to the penis to make a midline incision.</li>
		<li>Gently push the intestine with swab into the upper part of the abdominal cavity.</li>
		<li>Grasp the bladder with a pair of forceps and pull out the bladder from the abdominal cavity. Expose the ventral lobes of the prostate, which is located on the ventral portion of the urethra.</li>
		<li>Pull out the ventral lobes of the prostate, seminal vesicle, and vas deferens to expose the dorsal lobe of the prostate. Find the point of adhesion of the vas deferens and prostate.</li>
		<li>Separate the space between the prostate and vas deferens. Carefully expose the fibrous capsule, which is located posterior to the junction point of the prostate and vas deferens. Then find the major pelvic ganglion (Figure 2). 
		NOTE: The major pelvic ganglion and cavernous nerves can be seen at the surface of the prostate.</li>
		<li>Carefully dry CN area with a sterile swab.&#160;Carefully isolate and hook the right cavernous nerve with the bipolar electrodes.</li>
	</ol>
	</li>
	<li>Catheterize the Left Crura
	<ol>
		<li>Cut a small incision in the skin of the penis with dissecting scissors, and then carefully denude the skin of the penis shaft.</li>
		<li>Dissect the striated penile musculature. Find the upper branch of pubis bone.</li>
		<li>Expose the bulbospongiosus muscle, which covers the spongious bulb.</li>
		<li>Divide the bulbospongiosus muscle from the ischiocavernosus muscle by using curved forceps.</li>
		<li>Carefully isolate the ischiocavernosus muscle with curved forceps, and then cut the ischiocavernosus muscle to expose the white tunica albuginea of the crus corpus cavernosum.</li>
		<li>Following the anatomical direction of the crus corpus cavernosum, carefully insert the needle into the crus corpus cavernosum through the white tunica albuginea. 
		NOTE: This is a crucial step for successful catheterization. A small amount of heparinized saline can be injected, and a slight penile tumescence should be observed, if the needle has been inserted correctly.</li>
		<li>Carefully release the needle and avoid any sliding of the needle or disruption of the connecting tube. Inspect for any leakage.</li>
	</ol>
	</li>
</ol>
3. Stimulate the CN
<ol>
	<li>Open the software program for pressure signal recording and start the pressure signal recording.</li>
	<li>Set the parameters of the stimulation: 15 Hertz, pulse width of 5 milliseconds, 5 volts, and a 60 s duration. Stimulate CN at a frequency of 15&#160;Hz with a pulse width of 5 ms. 
	NOTE: A sharp rise of the ICP can be observed while applying the electrical stimulation.</li>
	<li>Allow a 30-minute rest interval between stimulations. The maximum of consecutive stimulation in each animal is three times.</li>
</ol>
4. End the Procedure
<ol>
	<li>After recording, administer euthanasia by injecting an overdose of pentobarbital sodium at a dose of 150 mg/kg body weight. Confirm the death of rats by checking their arterial pressure. Remove the rats, and clean the surgery tools.</li>
</ol>
5. Data Analysis
<ol>
	<li>Save and export the data from the software. The response is commonly expressed as the ratio of ICP to systemic mean arterial pressure (MAP). The ratio of peak ICP/MAP was calculated to evaluate the erectile function.</li>
	<li>Pool data from at least five rats, and analyze with statistical software. Differences are considered statistically significant when p &#60; 0.05, using Student&#39;s t-test.</li>
</ol>

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W. <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>Albersen, M., Orabi, H., Lue, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+and+treatment+of+erectile+dysfunction+in+the+aging+male:+A+mini-review.">Evaluation and treatment of erectile dysfunction in the aging male: A mini-review.</a> Gerontology. 58 (1), 3-14 (2012).</li><li>Hedlund, P., Matsumoto, K., Andersson, K. -. 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The authors have nothing to disclose

As a direct measure of erectile function, ICP is a reliable method14. It allows for the acquisition of data on basal ICP, peak ICP, plateau ICP, time to erection and detumescence time, duration of response, etc. Besides these direct measured parameters, there are some other index parameters: (1) &#34;T80&#34;, the time to reach 80% of peak ICP; (2) &#34;D20&#34;, the time to decrease to 20% of peak ICP; (3) &#34;&#916;T80&#34;, the rate of increase in pressure (per second) at T80; and (4)...

ED is defined as the inability to attain or keep a penile erection, and has become a common male sexual disorder1. Experimental animals are used and provide reproducible models to investigate erectile function2. For a long time, several larger animal models have been employed for investigating erectile function3,4,5. Although rodents are relatively small compared to other animals, they are also used for the study of male erectile dysfunction due to exhibiting several advantages6. First, the morphological and functional sexual characteristics of humans are recapitulated in rodents. Second, compared to larger animals used in ED studies, rodents are more economical to purchase, house, and maintain. Third, genetically modified rodent models provide advantages in reproducible and subsequent behavioral as well as neurophysiological studies. Therefore, rodents have quickly become the primary animals used in the study of male erectile dysfunction.
Benefiting from a pure genetic background and consistent culture conditions, rodent models have provided consistently reproducible data5,6,7,8. Among the numerous available studies related to many aspects of erectile functions, the apomorphine (APO)-induced erectile response test and the electric-stimulation-induced ICP response test are the most widely used methods that reliably reflect erectile function9,10,11,12. The APO-induced erectile function test, developed by Heaton et al.13, is a bio-assay that utilizes the phenomenon that administration of apomorphine to rats elicits erections and yawns. As an easy, noninvasive, and stable bio-assay to evaluate erectile function, the APO-induced erectile function test is widely used in many studies. However, this assay does not adequately reflect the quality of erections or the dynamic changes in blood flow associated with an erectile response14. ICP measurements were initially developed by Quinlan et al.15. In this method, a catheter is placed into the carotid artery to measure systemic blood pressure, and another catheter is inserted into the crus corpus cavernosum to record the ICP. Before or during the ICP recording, a vasoactive agent and/or electrical field stimulation of the major pelvic ganglion (MPG) or CN were often given to the rats14. This assay has been a reliable tool for evaluating the therapies and medicines for ED, and will likely be used as a vital evaluation method in the future6.
Compared to the easily-performed APO-induced erectile function test, successful ICP recordings require attention to detail, practice, and adherence to the operation method. Therefore, here, we provide a detailed description of how to perform ICP recording.

<table><tbody><tr><td>&lt;strong&gt;Animal&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Rats</td><td></td><td></td><td>Strain: Sprague-Dawley Age: 2-3 month</td></tr><tr><td>Rats</td><td></td><td></td><td>Strain: Sprague-Dawley Age: 15-18 month</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Saline</td><td>Sigma-Aldrich, S7653</td><td></td><td>dissolve 8.5 gram sodium chloride in distilled water</td></tr><tr><td>Pentobarbital sodium solution</td><td>Sigma-Aldrich, P3761</td><td></td><td>dissolve 1 gram in 100 ml saline</td></tr><tr><td>Povidone-iodine</td><td>BTP Pharmaceutical Co. Limited</td><td></td><td>10% (V/V)</td></tr><tr><td>Ethanol</td><td>China National Pharmaceutical Group Corporation (SINOPHARM)</td><td></td><td>70% (V/V)</td></tr><tr><td>Heparin</td><td>Sigma-Aldrich, H3149</td><td></td><td>dissolve 20000U heparin in 100 ml saline</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Hypodermic needle</td><td>Shandong Weigao Group Medical Polymer Co., Ltd.&amp;nbsp;</td><td>23G</td><td></td></tr><tr><td>Syringe</td><td>Shandong Weigao Group Medical Polymer Co., Ltd</td><td>10 ml</td><td></td></tr><tr><td>Three-way stopcock</td><td>Chengdu Instrument factory</td><td>TSK 01</td><td></td></tr><tr><td>Electrode</td><td>Chengdu Instrument factory</td><td>JST-1</td><td></td></tr><tr><td>Catheter tube</td><td>Chengdu Instrument factory</td><td>PE-10, PE-50</td><td></td></tr><tr><td>Operating scissors</td><td>Shanghai operation equipment factory</td><td>J22010, J22020</td><td></td></tr><tr><td>Ophthalmic operating scissors</td><td>Shanghai operation equipment factory</td><td>Y00010, Y00020</td><td></td></tr><tr><td>Ophthalmic forceps</td><td>Shanghai operation equipment factory</td><td>JD1010, JD1020</td><td></td></tr><tr><td>MicroScissors</td><td>World Precision Instruments</td><td>WAA260</td><td></td></tr><tr><td>silk suture</td><td>Shandong Weigao Group Medical Polymer Co., Ltd.&amp;nbsp;</td><td>5-0</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Stimulator</td><td>Nanjing medease science and technology co. ltd (model 4C501H)</td><td></td><td>15 Hz, 5 ms pulse, 5 V, 60 s duration and 5 minutes interval</td></tr><tr><td>Multichannel signal collection processing system</td><td>Nanjing medease science and technology co. ltd (model 4C501H)</td><td></td><td>Blood pressure model</td></tr><tr><td>Pressure transducer</td><td>Beijing Xin Hang Xing Ye Technology Trading Company Limited (model YP100)</td><td></td><td>40KPa</td></tr></tbody></table>

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.

Numerous studies have shown that erectile dysfunction in aged males is becoming a common problem. However, medical treatment is limited in the management of aging-related ED16. In rodent models of aging-related ED, many therapies are tested on the erectile function of aged rats. As we have introduced above, the ICP recording test could be used to distinguish the ED animals from the total population of experimental animals, which is also valuable to quantify the eff...

Watch this Scientific Journal Video about Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents at JoVE.com

Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents

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. ...

intracavernosal-pressure-recording-to-evaluate-erectile-function

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14.1K Views.  Nanjing University. Intracavernosal druk opname (ICP) is een belangrijke methode om te evalueren van de erectiele functie van proefdieren. Hier, een gedetailleerd protocol blijkt procedure voor de opname van ICP door catheterizing de crura penis en dan elektrisch stimuleren de cavernous zenuwen bij ratten.

Video: Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents

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 ...

Medicine

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. Despite the discovery of key regulators of uterine contractility, this process is still not fully understood. Transgenic mice provide an ideal model in which to study parturition. Previously, the only method to study uterine contractility in the mouse was ex vivo isometric tension recordings, which are suboptimal for several reasons. The uterus must be removed from its physiological environment, a limited time course of investigation is possible, and the mice must be sacrificed. The recent development of radiometric telemetry has allowed for longitudinal, real-time measurements of in vivo intrauterine pressure in mice. Here, the implantation of an intrauterine telemeter to measure pressure changes in the mouse uterus from mid-pregnancy until delivery is described. By comparing differences in pressures between wild type and transgenic mice, the physiological impact of a gene of interest can be elucidated. This technique should expedite the development of therapeutics used to treat myometrial disorders during pregnancy, including preterm labor.

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

This manuscript provides a protocol for implanting telemeters in the mouse for the purpose of measuring intrauterine pressures during pregnancy.

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. ...

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 ...

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.

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 ...

Simultaneous Electrocardiography Recording and Invasive Blood Pressure Measurement in Rats

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. Research focusing on cardiovascular disease models, potential cardiovascular therapeutic targets or pharmaceutical agents requires assessment of systemic arterial pressure and heart rhythm changes. In situations where radio telemetry systems are not available or affordable, the technique of femoral artery cannulation is an alternative way to obtain intra-arterial pressure waveform recordings and systemic BP measurements. This technique is economical and can be performed with standard equipment in animal facilities. However, invasive arterial pressure recording requires cannulation of small arteries, which can be a challenging surgical skill. Here, we present step-by-step protocols for femoral artery cannulation procedures. Key procedures include the calibration of the data acquisition system, tissue dissection and femoral artery cannulation, and setup of the arterial cannulation system for pressure recording. Surface electrocardiography recording procedures are also included. We also present examples of BP recordings from normotensive and hypertensive rats. This protocol allows reliable direct recordings of systemic BP with simultaneous electrocardiography.

Here, we describe a setup for simultaneous recording of electrocardiography and intra-arterial blood pressure (BP) in experimental rats, which can be done with standard equipment in animal facilities and can be applied to physiological or pharmacological studies to investigate pathogenic or therapeutic mechanisms in cardiovascular medicine.

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. ...

In Vivo Luminal Measurement of Distension-Evoked Urothelial ATP Release in Rodents

ATP, released from the urothelium in response to bladder distension, is thought to play a significant sensory role in the control of micturition. Therefore, accurate measurement of urothelial ATP release in a physiological setting is an important first step in studying the mechanisms that control purinergic signaling in the urinary bladder. Existing techniques to study mechanically evoked urothelial ATP release utilize cultured cells plated on flexible supports or bladder tissue pinned into Ussing chambers; however, each of these techniques does not fully emulate conditions in the intact bladder. Therefore, an experimental setup was developed to directly measure ATP concentrations in the lumen of the rodent urinary bladder.
In this setup, the bladders of anesthetized rodents are perfused through catheters in both the dome of the bladder and via the external urethral orifice. Pressure in the bladder is increased by capping the urethral catheter while perfusing sterile fluid into the bladder through the dome. Measurement of intravesical pressure is achieved using a pressure transducer attached to the bladder dome catheter, akin to the setup used for cystometry. Once the desired pressure is reached, the urethral catheter's cap is removed, and fluid collected for ATP quantification by luciferin-luciferase assay. Through this experimental setup, the mechanisms controlling both mechanical and chemical stimulation of urothelial ATP release can be interrogated by including various agonists or antagonists into the perfusate or by comparing results between wildtype and genetically modified animals.

In Vivo Luminal Measurement of Distension-Evoked Urothelial ATP Release in Rodents

This protocol describes the procedure for measuring ATP concentrations in the lumen of the bladder in an anesthetized rodent.

ATP, released from the urothelium in response to bladder distension, is thought to play a significant sensory role in the control of micturition. ...

Cystometric and External Urethral Sphincter Measurements in Awake Rats with Implanted Catheter and Electrodes Allowing for Repeated Measurements

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually performed as terminal analysis under urethane anesthesia. It is well known that anesthetic drugs can influence bladder function. Hence, the aim of this technique is to perform cystometric measurements of the urinary bladder and external urethral sphincter in lightly restrained awake rats. For this purpose, a bladder catheter is implanted into the bladder dome. Subsequently, two electrodes are implanted bilateral to the external urethral sphincter and a ground electrode is sutured to a non-responsive skeletal muscle. The bladder catheter and the three electrodes are finally tunneled subcutaneously to the neck region and affixed to a harness. With this technique, the lower urinary tract can be measured at multiple time points in the same animal to assess lower urinary tract function. The main application of this technique is the follow-up of simultaneous urinary bladder and external urethral sphincter function in awake healthy rats and after induction of a disease or injury. Moreover, subsequent lower urinary tract monitoring can be performed during evaluation of the disease/injury and to monitor treatment efficacy.

This protocol first describes the surgical procedure of the permanent implantation of a urinary bladder catheter combined with external urethral sphincter electrodes, and second, the measurement of the function of the urinary bladder and external urethral sphincter in implanted awake animals.

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually ...

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/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

Neuroscience

Neurophysiology

Stimulation and Modulation Techniques

Recording Changes in Renal Sympathetic Nerve Activity in Response to Arterial Pressure in Rats

Before arterial cannulation, inspect the pressure-sensing catheter under high magnification to confirm that the catheter is free of bubbles and debris and has an intact meniscus between the fluid and gel-filled components. Prior to each implantation, refill the gel at the distal tip of the catheter, and use a magnet to turn on the transmitter. Place a loose overhand half knot in the proximal suture to briefly occlude the femoral artery, and holding the catheter introducer in the non-dominant hand, use vessel cannulation forceps to grasp the tip of the cannula of the telemetry unit to avoid displacing the gel from the tip.
Use a bent 22-gauge needle tip to puncture a hole in the artery and insert the cannula into the artery as far as possible. Then, secure the pressure catheter with the proximal and distal suture ties, and tuck the body of the telemetry implant inside the flank adjacent to the incision. Gently move the rat into the prone position in a stereotaxic surgery frame with the head between the ear bars.
To isolate the renal sympathetic nerves, connect a wire Renal Sympathetic Nerve Activity or RSNA electrode to a 10X preamplifier in a microelectrode amplifier, and make a scalpel incision extending from 4 to 5 centimeters below the ribs in the caudal direction, slightly lateral to the spine.
Blunt-dissect to visualize the paraspinal muscles and make a very superficial 1 to 2-centimeter rostrocaudal incision where the fat meets the muscle. Using cotton-tipped applicators, spread the fat away from the muscle to visualize the kidney without entering the peritoneal space, and use retractors to gently separate the kidney from the paraspinal muscles to visualize the renal artery and abdominal aorta, and use high magnification to identify the renal nerves in the incision pocket.
The nerve bundles are most easily visible in the right angle formed by the aorta and the renal artery, closely following the renal artery from the aorta to the kidneys. Then, use micro-dissecting tweezers to gently dissect a segment of the nerve bundle to be placed on the recording electrode from the surrounding tissue. Secure the wire RSNA electrode in a holder and lower the electrode to the level of the nerve segment.
Use a nerve hook to gently lift the segment of the renal nerve onto the electrode without stretching the nerve and fill the incision with mineral oil to keep the exposed renal sympathetic nerve hydrated. Attach a grounding clip to the animal, in the other end to the Faraday cage, and use high and low pass filtering to direct the signal to the amplifiers. Then, adjust the gain up to 10 kilohertz, including an audio monitor, to assess the bursting pattern of the RSNA.
To assess the quality of the RSNA recording, evoke the baroreflex with a 100-microliter bolus intravenous injection of 10 micrograms per milliliter of phenylephrine to increase the blood pressure and to inhibit the renal sympathetic nerve activity, adjusting the position of the electrodes to improve the signal as necessary. Once a clear signal has been obtained, aspirate the mineral oil, and secure the electrode in place with silica gel.

Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents

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