This study was supported by the National Institute for Nursing Research (K99/R00NR014369).

All methods described were approved by the Institutional Animal Care Committee at the University of Illinois at Chicago.
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 RSNA4,5,6. 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 Vein (for Intravenous Access)
<ol>
	<li>Add heparin to 0.9% sterile saline (to achieve 20 units/mL). Fill a 1 mL syringe with the heparinized saline through a 22G needle. Connect 15 cm of PE-50 tubing to the needle and fill tubing with the solution.</li>
	<li>With the rat lying supine, create a 2 cm horizontal incision through the inguinal area. Using cotton-tipped applicators, dissect the connective tissue to expose the femoral vein and artery. To hold the incision open, either apply single-hook elastic surgical stays secured to the surgical field with silk tape or use small hemostats.</li>
	<li>Use hemostats to bend the tip of a 22G needle at a 90&#176; angle to serve as a catheter introducer10.</li>
	<li>Visualize the vessels under the microscope. Gently separate the vein and artery using curved forceps. Place two 12 cm long 5-0 silk suture beneath the vein (one distal and one proximal); place the suture in the same manner beneath the artery.
	<ol>
		<li>Tie the distal (bottom) suture to occlude the vein; secure the edges of this suture to the surgical field using either silk tape or small hemostats. Gently pull the vein taut but not with so much force that the vessel will tear. Position the vein perpendicular to the surgeon&#39;s dominant hand.</li>
	</ol>
	</li>
	<li>Use a loose overhand half knot in the proximal suture to briefly occlude the vein. With delicate hemostatic forceps, gently clamp this suture to occlude blood flow. Hold the 22G needle with the bent tip in the non-dominant hand; clasp the tubing with forceps using the dominant hand.</li>
	<li>Puncture a small hole in the vein with the catheter introducer (Step 3.3) and insert the PE-50 tubing (pre-filled with heparinized saline) into the vessel; use the bent needle to keep the opening in the vessel open and to assist in positioning the tip of the catheter into the vessel10.
	<ol>
		<li>Release the proximal suture and gently flush 0.2 mL of heparinized saline into the vein; advance the catheter. Check the blood return from the vein to ensure proper placement. Complete the proximal knot and, with distal suture tie, secure the tubing inside the vein.</li>
	</ol>
	</li>
	<li>Use venous access during the experiments for administering supplemental anesthesia or medications and for blood collection. Incorporate a 3-way connector if regular intravenous infusions and blood sampling will be necessary. Check noxious toe-pinch reflex every 15 min to titrate anesthesia.</li>
</ol>
4. 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 (Step 3.4) 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 a pressure transducer/infusion system
	<ol>
		<li>Connect the pressure transducer/tubing to a 500 mL 0.9% saline bag. Flush the tubing with saline, removing all bubbles, and place the bag inside a pressure inducer bag to pressurize the system.</li>
		<li>As described in Step 3.1, fill a 1 mL syringe with the heparinized saline through a 22G needle and connect 15 cm of PE-50 to the needle (flush tubing with heparinized saline).</li>
		<li>Use a loose half knot in the proximal suture to briefly occlude the artery. Hold the catheter introducer (Step 3.3) in the non-dominant hand; hold the distal tip of the PE-50 with vessel cannulation forceps in the dominant hand. Puncture a hole in the artery with the bent 22G needle and insert the cannula into the vessel.</li>
		<li>Release the proximal suture, gently flush 0.2 mL of heparinized saline into the artery and advance the catheter as far as possible. Check for arterial blood return to ensure proper placement. Complete the proximal knot and, with distal suture tie, secure the tubing inside the artery. Connect the arterial line to the pressure transducer/tubing. 
		NOTE: The distal portion of tubing can be taped to the rat&#39;s hindlimb to secure the arterial line. An alternative approach to vessel cannulation is described by Jespersen et al.;11 their protocol differs by using retractors to spread the incision, glue-rather than suture-to secure the tubing, and the approach does not include the bent needle introducer.</li>
	</ol>
	</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 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 (Step 3.3) 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>
5. 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 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 targeting12, drill a burr hole osteotomy, sized for electrode access, through the skull.</li>
</ol>
6. 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 dissect 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 the 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 Hz4,5,6,7,8. Use an increased sampling rate when a CNS manipulation is hypothesized to cause rapid/brief sympathetic responses.</li>
</ol>
7. 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 sympathoinhibition4,13,14.</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.
	<ol>
		<li>If the bursts of RSNA are not occurring cyclically with the cardiac cycle, and if there is any interference in the recording, then carefully reposition the electrode.</li>
		<li>As respiratory movements can also affect the quality of the RSNA recording, improve the signal by gently moving the electrode into a position where the muscle movements do not disrupt the electrode during breathing.</li>
	</ol>
	</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 RSNA4,5,6. As an alternative approach for quantifying noise, administer a short-acting ganglionic blocker such as atropine, hexamethonium, chlorisondamine, or pentolinium tartrate8,15,16,17.</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>
8. Euthanasia (Transcardiac Perfusion)
<ol>
	<li>Identify the locations of CNS manipulations by injecting dyes or fluorochromes, creating electrolytic lesions, or by detecting of c-fos expression.</li>
	<li>If brain analyses will require fixation, prepare the rat for transcardiac perfusion. Assess the toe-pinch reflex to ensure that the rat remains deeply anesthetized. Provide supplemental anesthesia if needed. Perform transcardiac perfusion of paraformaldehyde fixative in a fume hood. 
	CAUTION: Avid skin/eye irritant.</li>
	<li>Insert tubing into the perfusion pump and prime with 0.9% normal saline.</li>
	<li>Make a 5-6 cm lateral incision through the skin and abdominal wall immediately beneath the rib cage and open the chest cavity. Carefully separate the liver from the diaphragm. Make a small incision in the diaphragm using curved blunt scissors. Inject 0.1 mL of heparin directly into the left ventricle.</li>
	<li>Pass a perfusion needle into the left ventricle (a stainless steel gavage needle works well for this step) by either puncturing it into the heart or by cutting a small incision using sharp scissors and passing the gavage needle through so the tip is visible through the wall of the aorta (but does should not reach the aortic arch). Use a surgical or electrical clip to secure the needle in place.</li>
	<li>Using an infusion pump, administer 0.9% normal saline (room temperature). Immediately create a 2-3 mm incision in the right atrium to create an outlet for the saline rinse. Do not cut the descending aorta. Continue the saline rinse until the liver changes color from red/brown to pale yellow, an infusion of approximately 400 mL over 2-3 min.</li>
	<li>Stop the pump. Switch the perfusate to the fixative (e.g., 10% formalin or 4% paraformaldehyde); infuse 400 mL over 2-3 min. Remove the brain and store the specimen in fixative solution overnight at 4 &#176;C before transferring the tissue to 30% sucrose (30 mL of sucrose dissolved in 100 mL of 0.1 M phosphate-buffered saline) for at least 3 days or until the brain sinks, for cryoprotection before cryogenetic sectioning18.</li>
</ol>
9. Analyze Data
<ol>
	<li>Full-wave rectify the raw RSNA to obtain absolute values. Full-wave rectify a 10 s segment of the raw noise signal. It is important to exclude any studies that were affected by a low signal-to-noise ratio. In studies quantifying RSNA, investigators applied a priori criteria such as requiring signal to noise ratios to exceed 2:1 to 6:117,19,20.</li>
	<li>Calculate the mean rectified RSNA for non-overlapping segments (&#181;V) and subtract the noise estimate (&#181;V). Depending on the aims of the experiment, investigators may select intervals such as 10 s (Figure 1) or 1 s. Calculate means by using waveform analysis options in the physiologic software or exporting the data into spreadsheets to calculate averages for selected time intervals.</li>
	<li>To normalize across different animals, express values for further analysis as the percent change from baseline. Use parametric statistics to conduct group comparisons.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+can+we+learn+about+neural+control+of+the+cardiovascular+system+by+studying+rhythms+in+sympathetic+nerve+activity?.">What can we learn about neural control of the cardiovascular system by studying rhythms in sympathetic nerve activity?.</a> International Journal of Psychophysiology. 103, 69-78 (2016).</li><li>Charkoudian, N., Wallin, B. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sympathetic+neural+activity+to+the+cardiovascular+system:+integrator+of+systemic+physiology+and+interindividual+characteristics.">Sympathetic neural activity to the cardiovascular system: integrator of systemic physiology and interindividual characteristics.</a> Comprehensive Physiology. 4 (2), 825-850 (2014).</li><li>Malpas, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sympathetic+nervous+system+overactivity+and+its+role+in+the+development+of+cardiovascular+disease.">Sympathetic nervous system overactivity and its role in the development of cardiovascular disease.</a> Physiological Reviews. 90 (2), 513-557 (2010).</li><li>Chen, W. W., Xiong, X. Q., Chen, Q., Li, Y. H., Kang, Y. M., Zhu, G. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+sympathetic+afferent+reflex+and+its+implications+for+sympathetic+activation+in+chronic+heart+failure+and+hypertension.">Cardiac sympathetic afferent reflex and its implications for sympathetic activation in chronic heart failure and hypertension.</a> Acta Physiologica. 213 (4), 778-794 (2015).</li><li>Linz, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulation+of+renal+sympathetic+innervation:+Recent+insights+beyond+blood+pressure+control.">Modulation of renal sympathetic innervation: Recent insights beyond blood pressure control.</a> Clinical Autonomic Research. , (2018).</li></ol>

Anne M. Fink is a member of the Customer Advisory Board for Data Sciences International.

Critical steps for measuring RSNA include: (1) avoiding stretching of the renal artery and nerves when separating the kidney from the paraspinal muscle and when placing the nerve segment on the recording electrodes, (2) carefully dissecting the renal nerve fibers from the surrounding tissue/vessel, (3) ensuring that the electrode wires are free of tissue, blood, or lymph fluid, and (4) preventing the nerve from drying out by applying mineral oil to the renal nerve and silica gel to the nerve-electrode unit. For troublesh...

Pre-clinical discoveries about autonomic control of the cardiovascular system inform strategies for managing disorders such as hypertension, heart failure, and chronic kidney disease. Over-activity of the sympathetic nervous system and reduced vagal cardiac tone contribute to elevated blood pressure (BP)1. Chronically elevated renal sympathetic outflow enhances catecholamine secretion and decreases renal blood flow, with deleterious consequences to the cardiovascular/renal systems2,3. To define the neurobiological pathways leading to autonomic dysfunction, studies in rodents are important for determining how central nervous system (CNS) neurons regulate sympathetic parameters. The purpose of this protocol is to provide technical information about measuring renal sympathetic nerve activity (RSNA) and BP and to outline the techniques for quantifying acute sympathetic changes in response to CNS manipulations in anesthetized rats.
Acute (non-survival) RSNA measurements (lasting min to h) are useful when scientists will probe the CNS pharmacologically, electrically, or optogenetically in anesthetized rats to determine the functions of specific nuclei. Using these methods, structures such as the solitary nucleus, periaqueductal gray, pedunculopontine tegmentum, and rostral ventrolateral medulla have been investigated to define neurobiological pathways regulating sympathetic parameters4,5,6,7. This approach is important for identifying CNS targets to be investigated further in chronic models of autonomic dysfunction8,9. To complete these experiments, the laboratory requires a soldering iron, surgical microscope, stereotaxic frame, microelectrode amplifier, and audio monitor. Depending on the factors present in the laboratory that contribute to electrical noise, the surgical/recording area may require a Faraday cage/grounding strap to reduce electrical noise in the RSNA recording. If brain analyses will require tissue fixation, a perfusion pump and fume hood are required. Data can be digitized and recorded using multiple physiologic software/data acquisition (analogue-digital converter) units4,5, with different analysis options and compatibilities for incorporating telemetric signals.

<table>
	<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 surgery</td>
		</tr>
		<tr>
			<td>Micro-dissecting tweezers</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>11251-10</td>
			<td>RSNA surgery</td>
		</tr>
		<tr>
			<td>Micro-hook</td>
			<td>Fine Science Tools; Foster City, CA</td>
			<td>10064-14</td>
			<td>RSNA surgery</td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Fisher Scientific; Waltham, MA</td>
			<td>8042-47-5</td>
			<td>RSNA surgery</td>
		</tr>
		<tr>
			<td>Audio monitor</td>
			<td>A-M Systems; Sequim, WA</td>
			<td>3300</td>
			<td>RSNA surgery</td>
		</tr>
		<tr>
			<td>Silica gel</td>
			<td>Wacker, Munchen; Germany</td>
			<td>RT601A-B</td>
			<td>RSNA surgery</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>

quantifying acute changes in renal sympathetic nerve activity in response to central nervous system manipulations in anesthetized rats

Renal sympathetic nerve activity (RSNA) and mean arterial pressure are important parameters in cardiovascular and autonomic research; however, there are limited resources directing scientists in the techniques for measuring and analyzing these variables. This protocol describes the methods for measuring RSNA and mean arterial pressure in anesthetized rats. The protocol also includes the approaches for accessing the brain during RSNA recordings for central nervous system (CNS) manipulations. The RSNA recording technique is compatible with pharmacologic, optogenetic, or electrical stimulation of the CNS. The approach is useful when an investigator will measure short-term (min to h) autonomic responses in non-survival experiments to correlate anatomically with CNS nuclei. The approach is not intended to be used to obtain chronic (survival) recordings of RSNA in rats. Discharges in RSNA, averaged rectified RSNA, and mean arterial pressure can be quantified and analyzed further using parametric statistical tests. Methods for obtaining venous access, recording mean arterial pressure telemetrically, and brain fixation for future histological analysis are also described in the article.

Figure 1 illustrates a sample RSNA and BP recording from a Nembutal-anesthetized rat. An intravenous injection of phenylephrine was used to induce an increase in mean arterial pressure and to evoke the baroreflex and transient sympathoinhibition4,6. To quantify RSNA, the raw RSNA was rectified and averaged for non-overlapping 10 s segments; the noise estimate was subtracted from each segment.
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Watch this Scientific Journal Video about Quantifying Acute Changes in Renal Sympathetic Nerve Activity in Response to Central Nervous System Manipulations in Anesthetized Rats at JoVE.com

Quantifying Acute Changes in Renal Sympathetic Nerve Activity in Response to Central Nervous System Manipulations in Anesthetized Rats

Methods for measuring sympathetic and cardiovascular responses to central nervous system (CNS) manipulations are important for advancing neuroscience. This protocol was developed to assist scientists with measuring and quantifying acute changes in renal sympathetic nerve activity (RSNA) in anesthetized rats (non-survival).

Renal sympathetic nerve activity (RSNA) and mean arterial pressure are important parameters in cardiovascular and autonomic research; however, there are ...

quantifying-acute-changes-renal-sympathetic-nerve-activity-response

Research

JoVE Journal

Neuroscience

7.7K Views.  University of Illinois at Chicago. Methods for measuring sympathetic and cardiovascular responses to central nervous system (CNS) manipulations are important for advancing neuroscience. This protocol was developed to assist scientists with measuring and quantifying acute changes in renal sympathetic nerve activity (RSNA) in anesthetized rats (non-survival).

Video: Quantifying Acute Changes in Renal Sympathetic Nerve Activity in Response to Central Nervous System Manipulations in Anesthetized Rats

Optic Nerve Transection: A Model of Adult Neuron Apoptosis in the Central Nervous System

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve 
transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous
 chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals 
through the vitreous fluid ensures that they act upon the entire RGC population. Moreover, RGCs can be selectively transfected by applying short interfering RNAs 
(siRNAs), plasmids, or viral vectors to the cut end of the optic nerve 5-7 or injecting vectors into their target, the superior colliculus 8. 
This allows researchers to study apoptotic mechanisms in the desired neuronal population without confounding effects on other bystander neurons or surrounding glia. 
An additional benefit is the ease and accuracy with which cell survival can be quantified after injury. The retina is a flat, layered tissue and RGCs are localized in 
the innermost layer, the ganglion cell layer. The survival of RGCs can be tracked over time by applying a fluorescent tracer (3% Fluorogold) to the cut end of the 
optic nerve at the time of axotomy, or by injecting the tracer into the superior colliculus (RGC target) one week prior to axotomy. The tracer is retrogradely transported, labeling 
the entire RGC population. Because the ganglion cell layer is a monolayer (one cell thick), RGC densities can be quantified in flat-mounted tissue, without the need 
for stereology. Optic nerve transection leads to the apoptotic death of 90% of injured RGCs within 14 days postaxotomy 9-11. RGC apoptosis has a 
characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a time window for 
experimental manipulations directed against pathways involved in apoptosis.

Optic Nerve transection is a widely used model of adult CNS injury. Ninety percent of retinal ganglion cells (RGCs) whose axons are completely transected (axotomy) die within 14 days after axotomy. This model is easily amenable to experimental manipulations and highly reproducible.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are tightly regulated, affecting the final dynamics of protein abundance. Various regulatory mechanisms exist at the translation step, rendering mRNA levels alone an unreliable indicator of gene expression. In addition, local regulation of mRNA translation has been particularly implicated in neuronal functions, shifting &#39;translatomics&#39; to the focus of attention in neurobiology. The presented method can be used to bridge transcriptomics and proteomics.
Here we describe essential modifications to the technique of polyribosome fractionation, which interrogates the translatome based on the association of actively translated mRNAs to multiple ribosomes and their differential sedimentation in sucrose gradients. Traditionally, working with in vivo samples, particularly of the central nervous system (CNS), has proven challenging due to the restricted amounts of material and the presence of fatty tissue components. In order to address this, the described protocol is specifically optimized for use with minimal amount of CNS material, as demonstrated by the use of single mouse spinal cord and brain. Briefly, CNS tissues are extracted and translating ribosomes are immobilized on mRNAs with cycloheximide. Myelin flotation is then performed to remove lipid rich components. Fractionation is performed on a sucrose gradient where mRNAs are separated according to their ribosomal loading. Isolated fractions are suitable for a range of downstream assays, including new genome wide assay technologies.

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

This protocol illustrates essential modifications of polyribosome fractionation in order to study the translatome of in vivo CNS samples. It allows global assessment of translation and transcription regulation through the isolation and comparison of total RNA to ribosome bound RNA fractions.

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the pathophysiological basis of these diseases, it is necessary to precisely characterize in which way the processing of information is disturbed between the different neuronal parts of the circuit. Using extracellular in vivo electrophysiological recordings, it is possible to accurately delineate neuronal activity within a neuronal network. The application of this method has several advantages over alternative techniques, e.g., functional magnetic resonance imaging and calcium imaging, as it allows a unique temporal and spatial resolution and does not rely on genetically engineered organisms. However, the use of extracellular in vivo recordings is limited since it is an invasive technique that cannot be universally applied. In this article, a simple and easy to use method is presented with which it is possible to simultaneously record extracellular potentials such as local field potentials and multiunit activity at multiple sites of a network. It is detailed how a precise targeting of subcortical nuclei can be achieved using a combination of stereotactic surgery and online analysis of multi-unit recordings. Thus, it is demonstrated, how a complete network such as the hyperdirect cortico-basal ganglia loop can be studied in anesthetized animals in vivo.

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

In this study, the methodology is presented on how to perform multi-site in vivo electrophysiological recordings from the hyperdirect pathway under urethane anesthesia.

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the ...

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...