recording changes in renal sympathetic nerve activity in response to arterial pressure in rats

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.

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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. Create Bipolar RSNA Electrodes 
<ol>
	<li>To create the electrode, cut two pieces of stainless steel wire each approximately 18 mm long. Cut one piece of polyethylene (PE-50) tubing approximately 15 mm long. Feed both pieces of wire into the tubing, leaving the wire protruding from both ends.</li>
	<li>Remove the insulation from the ends of the wires; trim the wires leaving 2-3 mm of exposed wire. On one end, crimp male pins over the exposed wire. Solder the pins securely to the wire, secure the pins inside a connector strip, and cover the connection with epoxy. 
	NOTE: An alternative approach that avoids soldering is to use quick connect/release alligator clips.</li>
	<li>On the opposite end of the electrode, remove the insulation from the ends of the wires, leaving 2-3 mm of exposed wire. Bend this portion of the wires to create small &#34;V&#34; shaped hooks in the uninsulated wire. 
	NOTE: This is the portion of the electrode that will be in contact with the renal sympathetic nerve. It is important to seal this end to prevent fluids from entering the tubing. Silicone or epoxy effectively can be used.</li>
</ol>
2. Administer Anesthesia and Prepare Surgical Sites 
<ol>
	<li>Administer anesthesia to a male Sprague Dawley rat (age 9-11 weeks, weighing 150-400 g). Administer pentobarbital sodium 50 mg/kg via an intraperitoneal (IP) injection. To assess a stable plane of anesthesia during surgery, check toe-pinch reflex every 15 min and re-dose anesthesia as needed. 
	NOTE: Pentobarbital sodium (Nembutal) was used in previous studies to achieve a sustained plane of anesthesia without interfering with the modulation of RSNA. This protocol is for non-survival surgery, so there is no recovery/post-operative monitoring period.</li>
	<li>Prepare the surgical site according to institutional animal care guidelines (i.e., shave the rat&#39;s abdomen, back, and head; cleanse the skin with 10% povidone-iodine solution; apply eye lubricant; and place the rat on a heating pad). Maintain the body temperature at 37 &#176;C during the experiments.</li>
</ol>
3. Cannulate the Femoral Artery for Mean Arterial Pressure Monitoring
<ol>
	<li>Visualize the artery under the microscope. Similar to the method used for venous cannulation, tie the distal suture to occlude the artery; secure the edges of this suture to the surgical field with silk tape and position the artery perpendicular to the surgeon&#39;s dominant hand.</li>
	<li>Arterial access if using telemetry
	<ol>
		<li>Inspect the pressure-sensing catheter under high magnification before arterial cannulation. Ensure that the catheter is free of bubbles/ debris and has an intact meniscus between the fluid-filled (proximal) and gel-filled (distal) components. Prior to each implantation, refill the gel at the distal tip of the catheter. Turn on the transmitter using a magnet; monitor blood pressure (BP) during the surgery to endure perfect placement.</li>
		<li>Use a loose overhand half knot in the proximal suture to briefly occlude the femoral artery. Hold the catheter introducer in the non-dominant hand. Hold the tip of the cannula of the telemetry unit with vessel cannulation forceps to avoid displacing gel from the tip.</li>
		<li>Puncture a hole in the artery with the bent 22G needle and insert the cannula into the artery using vessel cannulation forceps to avoid displacing gel from the tip. Advance the cannula as far as possible. Using the proximal and distal suture ties, secure the pressure catheter.</li>
		<li>Tuck the body of the telemetry implant inside the flank adjacent to the incision and close this incision using 4-0 nylon suture on a cutting needle. Turn the telemetry devices off by magnet at the conclusion of the recording period to conserve battery life.</li>
	</ol>
	</li>
</ol>
4. Position the Rat in the Stereotaxic Surgery Frame to Access the Brain
<ol>
	<li>Gently move the rat into the prone position in the stereotaxic surgery frame.</li>
	<li>Position the rat between the ear bars and adjust the incisor bar to equalize the height of lambda and bregma. Positioning may depend on the rat strain, weight, and locations of central nervous system (CNS) targets.</li>
	<li>Make a 2 cm rostrocaudal scalpel incision through the midline of the scalp. Using cotton-tipped applicators, firmly remove connective tissue from the skull surface. Apply hydrogen peroxide to the skull to assist in visualizing the bregma, lambda, and midline sutures.</li>
	<li>Using an atlas of the rat brain to guide targeting, drill a burr hole osteotomy, sized for electrode access, through the skull.</li>
</ol>
5. Isolating the Renal Sympathetic Nerves
<ol>
	<li>Connect the wire RSNA electrode (Steps 1.1-1.3) to a 10X pre-amplifier and microelectrode amplifier.</li>
	<li>Isolate the renal nerves through a retroperitoneal incision prior to, or after, the rat is secured in the stereotaxic frame. Position the RSNA electrodes once the rat is in the stereotaxic frame. Make a scalpel incision extending from 4-5 cm below the ribs in the caudal direction, slightly lateral to the spine. Blunt dissects the incision to visualize the paraspinal muscles.</li>
	<li>Use scissors to make a very superficial 1-2 cm rostrocaudal incision where the fat meets the muscle. Using cotton-tipped applicators, spread the fat away from the muscle to visualize the kidney. It is important not to enter peritoneal space.</li>
	<li>Use retractors to gently separate the kidney from the paraspinal muscles to visualize the renal artery and abdominal aorta. Do not stretch the vessels excessively to avoid damaging the renal nerves. Use a 2&#34; x 2&#34; cotton gauze pad soaked in saline to protect the kidney from injury.</li>
	<li>Under high magnification, 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. The renal nerves closely follow the renal artery from the aorta to the kidneys.</li>
	<li>Select a segment of the nerve bundle that will be placed on the recording electrode. Gently dissect the nerve fibers from the surrounding tissue/vessel using micro-dissecting tweezers.</li>
	<li>Secure the wire RSNA electrode in a holder (e.g., an alligator clip attached to a support stand). 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. 
	NOTE: The nerve should rest inside both &#34;V&#34; shaped hooks in the uninsulated wire, parallel to the nerve. The electrode wires must not touch any other tissue, blood, or lymph fluid.</li>
	<li>Fill the incision with mineral oil to prevent the exposed renal sympathetic nerve from becoming dry. Use a grounding clip with one end on the skin of the incision and the other attached to the Faraday cage.</li>
	<li>Direct the signal to the amplifiers using high- and low-pass filtering (10 Hz and 3 kHz). Adjust the gain up to 10 K. Include an audio monitor to assess the bursting pattern of the RSNA. Use sampling rates ranging between 2,000-10,000 Hz. Use an increased sampling rate when a CNS manipulation is hypothesized to cause rapid/brief sympathetic responses.</li>
</ol>
6. Record Data
<ol>
	<li>Assess the quality of the RSNA recording by evoking the baroreflex with a bolus injection of 1 mL of saline or 10 &#181;g/mL phenylephrine (in 0.1 mL) intravenously. As illustrated in Figure 1, the infusion should increase BP and inhibit RSNA. An increase in mean arterial pressure of 60-80 mmHg is sufficient for renal sympathoinhibition.</li>
	<li>Adjust the position of the electrodes to improve the signal if necessary. Repositioning is required if the nerve is not in contact with both hooks on the electrode or if any tissue, blood, or lymph fluid is in contact with the wires. 
	NOTE: The need for repositioning is based on the auditory characteristics of the nerve discharges.</li>
	<li>Once a clear signal is obtained, secure the RSNA electrode in place by withdrawing mineral oil and applying a silica gel to cover the nerve/ electrode connection in the incision pocket. Do not move the rat before the gel has set completely.</li>
	<li>Perform CNS manipulation protocols while continuously recording RSNA and mean arterial pressure. If a microinjector/pulser is used for brainstem manipulations, a logic signal from this device can be introduced into the RSNA/BP recordings to document the timing of CNS manipulations.</li>
	<li>When the experiment is complete, determine the noise level by crushing the nerve proximal to the recording electrodes between the silica gel and the spinal muscle. Record at least 30 s of this &#34;zero&#34; value for RSNA. As an alternative approach for quantifying noise, administer a short-acting ganglionic blocker such as atropine, hexamethonium, chlorisondamine, or pentolinium tartrate.</li>
	<li>Carefully remove the RSNA electrode and remove any traces of silica gel from the wire electrodes. Save the electrode for re-use. Turn off the telemetry transmitter and remove it, taking care not to damage the tip of the catheter.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Stainless steel wire</td>
			<td>A-M Systems; Sequim, WA</td>
			<td>791000</td>
			<td>RSNA electrode</td>
		</tr>
		<tr>
			<td>Polyethylene (PE-50) tubing</td>
			<td>VWR; Radnor, PA</td>
			<td>63019-048</td>
			<td>RSNA electrode; vessel cannulation</td>
		</tr>
		<tr>
			<td>Miniature pin connector</td>
			<td>A-M Systems; Sequim, WA</td>
			<td>520200</td>
			<td>RSNA electrode</td>
		</tr>
		<tr>
			<td>Crimping tool</td>
			<td>Daniels Manufacturing Corp.; Orlando, FL</td>
			<td>M22520</td>
			<td>RSNA electrode</td>
		</tr>
		<tr>
			<td>Connector strip</td>
			<td>Amphenol; Clinton Township, MI</td>
			<td>221-2653</td>
			<td>RSNA electrode</td>
		</tr>
		<tr>
			<td>J-B Kwik Epoxy</td>
			<td>J-B Weld, Sulphur Springs, TX</td>
			<td>8270</td>
			<td>RSNA electrode</td>
		</tr>
		<tr>
			<td>Silicone</td>
			<td>Permatex; Hartford, CT</td>
			<td>2222</td>
			<td>RSNA electrode</td>
		</tr>
		<tr>
			<td>Heparin sodium; Injectable (10 mL vial, 1000 U/mL)</td>
			<td>KV Veterinary Supply; David City, NE</td>
			<td>P03466</td>
			<td>Venous line patency</td>
		</tr>
		<tr>
			<td>Phenylephrine HCl; Injectable (1 mL vial; 10 mg/mL)</td>
			<td>ACE Surgical Supply; Brockton, MA</td>
			<td>950-6312</td>
			<td>Testing renal sympathoinhibition</td>
		</tr>
		<tr>
			<td>Single-hook elastic surgical stays</td>
			<td>Harvard Apparatus; Holliston, MA</td>
			<td>72-2595</td>
			<td>Incision</td>
		</tr>
		<tr>
			<td>Silk surgical tape</td>
			<td>3M, Minneapolis, MN</td>
			<td>1538-0</td>
			<td>Secure surgical stays</td>
		</tr>
		<tr>
			<td>Needles, 20 G</td>
			<td>Sigma-Aldrich; St. Louis, MO</td>
			<td>Z192554-100EA</td>
			<td>Vessel cannulation</td>
		</tr>
		<tr>
			<td>Dumont #7 curved forceps</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>11274-20</td>
			<td>Vessel cannulation</td>
		</tr>
		<tr>
			<td>5-0 silk suture ties</td>
			<td>Braintree Scientific; Braintree, MA</td>
			<td>SUT-S 106</td>
			<td>Vessel cannulation</td>
		</tr>
		<tr>
			<td>Delicate hemostatic forceps</td>
			<td>Roboz Surgical Instrument Co.; Gaithersburg, MD</td>
			<td>RS-7117</td>
			<td>Vessel cannulation and RSNA surgery</td>
		</tr>
		<tr>
			<td>Crile Hemostatic forceps</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>13004-14</td>
			<td>Needle bending</td>
		</tr>
		<tr>
			<td>Telemetry transmitter</td>
			<td>Data Sciences International; Minneapolis, MN</td>
			<td>PA-10</td>
			<td>Mean arterial pressure monitoring (telemetry)</td>
		</tr>
		<tr>
			<td>Re-gel syringe</td>
			<td>Data Sciences International; Minneapolis, MN</td>
			<td>276-0038-001</td>
			<td>Transmitter reuse (telemetry)</td>
		</tr>
		<tr>
			<td>Disposable pressure transducer</td>
			<td>Transpac; San Clemente, CA</td>
			<td>MI-1224</td>
			<td>Mean arterial pressure monitoring</td>
		</tr>
		<tr>
			<td>Clear-Cuff pressure infuser</td>
			<td>MILA International Inc.; Florence, KY</td>
			<td>2281339</td>
			<td>Mean arterial pressure monitoring</td>
		</tr>
		<tr>
			<td>Vessel cannulation forceps</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>00574-11</td>
			<td>Catheter insertion</td>
		</tr>
		<tr>
			<td>Black monofilament nylon 4-0 suture on reverse cutting needle</td>
			<td>McKesson Medical-Surgical; San Francisco, CA</td>
			<td>S661GX</td>
			<td>Secure telemetry transmitter</td>
		</tr>
		<tr>
			<td>Telemetry receiver</td>
			<td>Data Sciences International; Minneapolis, MN</td>
			<td>RPC-1</td>
			<td>Mean arterial pressure monitoring (telemetry)</td>
		</tr>
		<tr>
			<td>LabChart Pro (software), PowerLab (acquisition hardware)</td>
			<td>AD Instruments; Colorado Springs, CO</td>
			<td>ML846, MX2 matrix 2.0 (Compatible with Data Science International telemetry)</td>
			<td>3 options for software/acquisition hardware</td>
		</tr>
		<tr>
			<td>SciWorks (software), DataWave (acquisition hardware)</td>
			<td>DataWave Technologies, Loveland, CO</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Spike 2 (software), Micro1401-3</td>
			<td>Cambridge Electronic Design Ltd., London UK</td>
			<td>1401-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-drill</td>
			<td>Roboz Surgical Instrument Co.; Gaithersburg, MD</td>
			<td>RS-6300</td>
			<td>CNS surgery</td>
		</tr>
		<tr>
			<td>Stereotaxic surgery frame</td>
			<td>Stoelting; Wood Dale, IL</td>
			<td>51600</td>
			<td>CNS surgery</td>
		</tr>
		<tr>
			<td>Microelectrode amplifier with 10X pre-amplifier</td>
			<td>A-M Systems; Sequim, WA</td>
			<td>1800-2</td>
			<td>RSNA recording</td>
		</tr>
		<tr>
			<td>Retractors</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>17009-07</td>
			<td>RSNA recording</td>
		</tr>
		<tr>
			<td>Micro-dissecting tweezers</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>11251-10</td>
			<td>RSNA recording</td>
		</tr>
		<tr>
			<td>Micro-hook</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>10064-14</td>
			<td>RSNA recording</td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Fisher Scientific; Waltham, MA</td>
			<td>8042-47-5</td>
			<td>RSNA recording</td>
		</tr>
		<tr>
			<td>Audio monitor</td>
			<td>A-M Systems; Sequim, WA</td>
			<td>3300</td>
			<td>RSNA recording</td>
		</tr>
		<tr>
			<td>Silica gel</td>
			<td>Wacker, Munchen; Germany</td>
			<td>RT601A-B</td>
			<td>RSNA recording</td>
		</tr>
		<tr>
			<td>Electrical clips</td>
			<td>Tyco Electronics; Schaffhausen, Switzerland</td>
			<td>EB0283-000</td>
			<td>Grounding or securing perfusion needle</td>
		</tr>
		<tr>
			<td>Bonn scissors, straight/sharp points</td>
			<td>Roboz Surgical Instrument Co; Gaithersburg, MD</td>
			<td>RS-5840</td>
			<td>Perfusion</td>
		</tr>
		<tr>
			<td>Gavage needle</td>
			<td>Harvard Apparatus; Holliston, MA</td>
			<td>75-0286</td>
			<td>Perfusion</td>
		</tr>
		<tr>
			<td>Masterflex perfusion pump</td>
			<td>Cole-Parmer; Vernon Hills, IL</td>
			<td>7524-10</td>
			<td>Perfusion</td>
		</tr>
		<tr>
			<td>Masterflex platinum-cured silicone tubing</td>
			<td>Cole-Parmer; Vernon Hills, IL</td>
			<td>96410-15</td>
			<td>Perfusion</td>
		</tr>
		<tr>
			<td>Formalin (10% buffered solution; 4 L)</td>
			<td>Sigma-Aldrich; St. Louis, MO</td>
			<td>HT501128</td>
			<td>Perfusion</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich; St. Louis, MO</td>
			<td>S0389</td>
			<td>Cryoprotection</td>
		</tr>
	</tbody>
</table>

Source: Fink, A. M., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/58205">Quantifying Acute Changes in Renal Sympathetic Nerve Activity in Response to Central Nervous System Manipulations in Anesthetized Rats.</a> J. Vis. Exp. (2018).
This video demonstrates a technique of studying the activity of renal sympathetic nerves (RSNs) in response to changes in arterial pressure in rats. Upon attaching a pressure-sensing catheter to the femoral artery, the renal sympathetic nerves are exposed. The nerves are then lifted onto a recording electrode, and their electrical activity is recorded. Next, a vasoconstricting drug is injected to elevate blood pressure, and corresponding changes in RSN activity are monitored.

<img alt="Figure 1" src="/files/ftp_upload/22747/22747_Figure1.jpg" /> 
Figure 1: RSNA and BP in response to phenylephrine injection IV.&#160;The raw RSNA (A) was full-wave rectified (B); rectified crushed &#34;zero&#34; RSNA is shown in inset&#160;C. Non-overlapping 10 s averages (minus noise) were calculated (D). To evoke the baroreflex, 0.1 mL of phenylephrine (1 &#181;g/mL) was injected intraveno...

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

Take an anesthetized rat in the supine position and make an incision in the inguinal region to expose the femoral artery.
Occlude the artery to temporarily restrict blood flow. Create an opening, insert a pressure-sensing catheter to measure arterial blood pressure, and secure the catheter.
Position the rat in the prone position in a stereotaxic frame and make a lateral incision below the ribs.
Using a microscope, identify the renal sympathetic nerves, or RSNs, that regulate blood flow to the kidney.
Lift the nerve onto a recording electrode, add oil to prevent nerve dehydration and record electrical activity.
Intravenously inject a drug that causes arterial narrowing, which elevates systemic blood pressure.
Baroreceptors detect elevated blood pressure and signal the brainstem to inhibit sympathetic nerves, including RSNs.
Remove the oil, apply a gel to secure the electrode for stable recording, and monitor the RSN activity.&#160;
A transient inhibition of RSN activity coinciding with increased arterial pressure confirms drug-induced RSN inhibition.

86 Aufrufe. Aufzeichnung von Veränderungen der Aktivität des sympathischen Nervus der Niere in der Niere als Reaktion auf den arteriellen Druck bei Ratten

Serie: Aufzeichnung von Veränderungen der Aktivität des sympathischen Nervus der Niere in der Niere als Reaktion auf den arteriellen Druck bei Ratten - Experiment

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.

Neuer Ansatz zur gleichzeitigen Erfassung von renalen sympathischen Nerven-Aktivität und Blutdruck mit intravenösen Infusion in dem Bewusstsein, hemmungslose Mäuse.

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

Forschung

JoVE Journal

Medizin

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain death (BD) still represents the major source for solid transplants. Unfortunately, the irreversible loss of brain function is known to induce multiple pathophysiological changes, including hemodynamic as well as hormonal modifications, finally leading to a systemic inflammatory response. Models that allow a systematic investigation of these effects in vivo are scarce. We present a murine model of BD induction, which could aid investigations into the devastating effects of BD on allograft quality. After implementing intra-arterial blood pressure measurement via the common carotid artery and reliable ventilation via a tracheostomy, BD is induced by steadily increasing intracranial pressure using a balloon catheter. Four hours after BD induction, organs may be harvested for analysis or for further transplantation procedures. Our strategy enables the comprehensive analysis of donor BD in a murine model, therefore allowing an in-depth understanding of BD-related effects in solid organ transplantation and potentially paving the way to optimized organ preconditioning.

We present a murine model of brain death induction in order to evaluate the influence of its pathophysiological effects on organs as well as on consecutive grafts in the context of solid organ transplantation.

Hirntodinduktion bei Mäusen mit intraarterieller Blutdrucküberwachung und -beatmung mittels Tracheostomie

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain ...

Neurowissenschaften

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.

Gleichzeitige Elektrokardiographie Aufnahme und Invasive Blut Druckmessung bei Ratten

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

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

Transkript

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