Name: Video: إجراء تحفيز الأعصاب الكهفية في الفئران لقياس الضغط داخل الكهف - Concept
Uploaded: 2024-10-31T13:46:41.000+00:00
Duration: 1 min 13 s
Description: 101 Views. المصدر: Hox, M., et al. تحفيز الأعصاب الكهفية وتسجيل الضغط داخل الكهف في الفئران. J. Vis. Exp. (2018). يوضح هذا الفيديو عزل وتحفيز العصب الكهفي في فأر مخدر لدراسة تغيرات ضغط الدم في الجسم الكهفي.

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Preparation of Tubing, Electrode, and Instruments for the Surgical Procedure
<ol>
	<li>Use the following microsurgical instruments: surgical scissors, angled micro scissors, tissue forceps, a pair of Dumont #7 and #5 curved micro forceps, a micro needle holder, and retractors. 
	NOTE: This is an acute procedure, so the instruments need not 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 piece of polyethylene (PE)-50 tubing to make a catheter for the intracavernous pressure (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 &#956;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. Then, 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: When 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 system&#39;s 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 a scalpel first 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 cavernous nerve (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 insertion point. Hold the needle with straight micro forceps in the other dominant hand and 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>When running the recording program (e.g., Spike 2), continuously record 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 a diminished response (Figure 10).</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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</td>
			<td>Science Products GmbH, Heidelberg, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon Glue</td>
			<td>Kwik-Sil, World Precision Instruments, Sarasota, FL, USA</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

<img alt="Figure 1" src="/files/ftp_upload/22752/22752_Figure1.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.
<img alt="Figure 2" src="/files/ftp_upload/22752/22752_Figure2.jpg" /> 
Figure 2. Dissection of the ...

Performing Cavernous Nerve Stimulation in Rats to Measure Intracavernous Pressure

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

Take an anesthetized rat with surgically exposed tunica albuginea, a connective tissue layer surrounding the cavernous body &#8212; the erectile tissue of the penis.&#160;
Next, make a midline abdominal incision to expose the prostate, retract it, and visualize the cavernous nerve.
Slide a suture under the nerve to elevate it and place the hooks of a bipolar electrode around the nerve.
Dry the nerve and secure it to the electrode with glue.
Take a heparinized saline-filled tubing connected to a needle and insert it through the tunica into the cavernous body.
Apply electrical impulses to stimulate the cavernous nerve, triggering neurotransmitter release.
These neurotransmitters stimulate the endothelial cells of the cavernous artery to produce nitric oxide, which diffuses into the smooth muscles of the cavernous body, inducing muscle relaxation.
This causes blood vessel dilation, increasing blood flow and pressure within the cavernous body with each impulse, recorded by the saline-filled tubing.

performing-cavernous-nerve-stimulation-rats-to-measure-intracavernous

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Stimulation and Modulation Techniques

101 Views. المصدر: Hox, M., et al. تحفيز الأعصاب الكهفية وتسجيل الضغط داخل الكهف في الفئران. J. Vis. Exp. (2018). يوضح هذا الفيديو عزل وتحفيز العصب الكهفي في فأر مخدر لدراسة تغيرات ضغط الدم في الجسم الكهفي. 

Video: إجراء تحفيز الأعصاب الكهفية في الفئران لقياس الضغط داخل الكهف - Concept

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.

نهج جديد للتسجيل المتزامن لنشاط العصب متعاطفة مع الكلي وضغط الدم مع التسريب الوريدي في واعية، الفئران غير المقيد.

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

JoVE Journal

Medicine

Subcutaneous Trigeminal Nerve Field Stimulation for Refractory Facial Pain

Chronic or neuropathic trigeminal facial pain can be challenging to treat. Neurosurgical procedures should be applied when conservative treatment fails. Neuromodulation techniques for chronic facial pain include deep brain stimulation and motor cortex stimulation, which are complex to perform. Subcutaneous nerve field stimulation is certified for chronic back pain and is the least invasive form of neuromodulation. We applied this technique to treat chronic and neuropathic trigeminal pain as an individual therapy concept. First, trial stimulation is performed. Subcutaneous leads are placed in the painful trigeminal dermatome under local anesthesia. The leads are connected to an external neurostimulator that applies constant stimulation. Patients undergo a 12 day outpatient trial to assess the effect of the stimulation. Electrodes are removed after the trial. If the patient reports pain reduction of at least 50% in intensity and/or attack frequency, a reduction in medication or increase in quality of life, permanent implantation is scheduled. New electrodes are implanted under general anesthesia and are subcutaneously tunneled to an infraclavicular internal pulse generator. Patients are able to turn stimulation on and off and to increase or decrease the stimulation amplitude as needed.
This technique represents a minimal invasive alternative to other more invasive means of neuromodulation for trigeminal pain such as motor cortex stimulation or deep brain stimulation.

Treating chronic or neuropathic facial pain can be challenging when medical or standard treatment fails. Subcutaneous nerve field stimulation is the least invasive form of neuromodulation and is used for chronic back pain. We applied this technology to treat chronic and neuropathic trigeminal facial pain.

تحت الجلد العصب الثلاثي التوائم تحفيز لآلام الوجه حرارية

Chronic or neuropathic trigeminal facial pain can be challenging to treat. Neurosurgical procedures should be applied when conservative treatment fails. ...

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.

تسجيل كهربية متزامنة والدم الغازية الضغط القياس في الفئران

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

Performing Cavernous Nerve Stimulation in Rats to Measure Intracavernous Pressure

Transcript

Performing Cavernous Nerve Stimulation in Rats to Measure Intracavernous Pressure