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

Samenvatting

Source: Fink, A. M., et al. Quantifying Acute Changes in Renal Sympathetic Nerve Activity in Response to Central Nervous System Manipulations in Anesthetized Rats. 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.

Protocol

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

1. 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.
2. 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.
3. 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 "V" 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.

2. Administer Anesthesia and Prepare Surgical Sites

1. 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.
2. Prepare the surgical site according to institutional animal care guidelines (i.e., shave the rat'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 °C during the experiments.

3. Cannulate the Femoral Artery for Mean Arterial Pressure Monitoring

1. 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's dominant hand.
2. Arterial access if using telemetry
   1. 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.
   2. 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.
   3. 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.
   4. 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.

4. Position the Rat in the Stereotaxic Surgery Frame to Access the Brain

1. Gently move the rat into the prone position in the stereotaxic surgery frame.
2. 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.
3. 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.
4. Using an atlas of the rat brain to guide targeting, drill a burr hole osteotomy, sized for electrode access, through the skull.

5. Isolating the Renal Sympathetic Nerves

1. Connect the wire RSNA electrode (Steps 1.1-1.3) to a 10X pre-amplifier and microelectrode amplifier.
2. 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.
3. 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.
4. 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" x 2" cotton gauze pad soaked in saline to protect the kidney from injury.
5. 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.
6. 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.
7. 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 "V" shaped hooks in the uninsulated wire, parallel to the nerve. The electrode wires must not touch any other tissue, blood, or lymph fluid.
8. 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.
9. 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.

6. Record Data

1. Assess the quality of the RSNA recording by evoking the baroreflex with a bolus injection of 1 mL of saline or 10 µg/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.
2. 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.
3. 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.
4. 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.
5. 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 "zero" value for RSNA. As an alternative approach for quantifying noise, administer a short-acting ganglionic blocker such as atropine, hexamethonium, chlorisondamine, or pentolinium tartrate.
6. 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.

Resultaten

Figure 1: RSNA and BP in response to phenylephrine injection IV. The raw RSNA (A) was full-wave rectified (B); rectified crushed "zero" RSNA is shown in inset C. Non-overlapping 10 s averages (minus n...

Materialen

Lijst van materialen gebruikt in dit artikel

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