Name: quantifying acute changes in renal sympathetic nerve activity in response to central nervous system manipulations in anesthetized rats
Uploaded: 2018-09-11T14:00:00
Description: 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

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 leav...

<ol>
	<li>Mancia, G., Grassi, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+autonomic+nervous+system+and+hypertension.">The autonomic nervous system and hypertension.</a> Circulation Research. 114 (11), 1804-1814 (2014).</li><li>Kannan, A., Medina, R. I., Nagajothi, N., Balamuthusamy, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+sympathetic+nervous+system+and+the+effects+of+denervation+on+renal+arteries.">Renal sympathetic nervous system and the effects of denervation on renal arteries.</a> World Journal of Cardiology. 6 (8), 814-823 (2014).</li><li>Johns, E. J., Kopp, U. C., DiBona, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+control+of+renal+function.">Neural control of renal function.</a> Comprehensive Physiology. 1 (2), 767 (2011).</li><li>Fink, A. M., Dean, C., Piano, M. R., Carley, D. 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F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Afferent+renal+denervation+impairs+baroreflex+control+of+efferent+renal+sympathetic+nerve+activity.">Afferent renal denervation impairs baroreflex control of efferent renal sympathetic nerve activity.</a> American Journal of Physiology, Regulatory, Integrative and Comparative Physiology. 295 (6), R1882-R1890 (2008).</li><li>Guild, S. J., 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=Quantifying+sympathetic+nerve+activity:+problems,+pitfalls+and+the+need+for+standardization.">Quantifying sympathetic nerve activity: problems, pitfalls and the need for standardization.</a> Experimental Physiology. 95 (1), 41-50 (2010).</li><li>Stocker, S. D., Hunwick, K. J., Toney, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypothalamic+paraventricular+nucleus+differentially+supports+lumbar+and+renal+sympathetic+outflow+in+water-deprived+rats.">Hypothalamic paraventricular nucleus differentially supports lumbar and renal sympathetic outflow in water-deprived rats.</a> Journal of Physiology. 15 (563 Pt 1), 249-263 (2005).</li><li>Stocker, S. D., Gordon, K. W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glutamate+receptors+in+the+hypothalamic+paraventricular+nucleus+contribute+to+insulin-induced+sympathoexcitation.">Glutamate receptors in the hypothalamic paraventricular nucleus contribute to insulin-induced sympathoexcitation.</a> Neurophysiology. 113 (5), 1302-1309 (2015).</li><li>Saponjic, J., Radulovacki, M., Carley, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Injection+of+glutamate+into+the+pedunculopontine+tegmental+nuclei+of+anesthetized+rat+causes+respiratory+dysrhythmia+and+alters+EEG+and+EMG+power.">Injection of glutamate into the pedunculopontine tegmental nuclei of anesthetized rat causes respiratory dysrhythmia and alters EEG and EMG power.</a> Sleep and Breathing. 9 (2), 82-91 (2005).</li><li>DiBona, G. F., Jones, S. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+analysis+of+renal+nerve+activity+responses+to+baroreceptor+denervation+in+hypertensive+rats.">Dynamic analysis of renal nerve activity responses to baroreceptor denervation in hypertensive rats.</a> Hypertension. 37 (4), 1153-1163 (2001).</li><li>Kunitake, T., Kannan, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Discharge+pattern+of+renal+sympathetic+nerve+activity+in+the+conscious+rat:+spectral+analysis+of+integrated+activity.">Discharge pattern of renal sympathetic nerve activity in the conscious rat: spectral analysis of integrated activity.</a> Journal of Neurophysiology. 84 (6), 2859-2867 (2000).</li><li>Machado, B. H., Bonagamba, L. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+L-glutamate+into+the+nucleus+tractus+solitarii+increases+arterial+pressure+in+conscious+rats.">Microinjection of L-glutamate into the nucleus tractus solitarii increases arterial pressure in conscious rats.</a> Brain Research. 576 (1), 131-138 (1992).</li><li>Murakami, M., 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=Inhalation+anesthesia+is+preferable+for+recording+rat+cardiac+function+using+an+electrocardiogram.">Inhalation anesthesia is preferable for recording rat cardiac function using an electrocardiogram.</a> Biological and Pharmaceutical Bulletin. 37 (5), 834-839 (2014).</li><li>Nakamura, T., Tanida, M., Niijima, A., Hibino, H., Shen, J., Nagai, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Auditory+stimulation+affects+renal+sympathetic+nerve+activity+and+blood+pressure+in+rats.">Auditory stimulation affects renal sympathetic nerve activity and blood pressure in rats.</a> Neuroscience Letters. 416 (2), 107-112 (2007).</li><li>Bardgett, M. E., McCarthy, J. J., Stocker, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glutamatergic+receptor+activation+in+the+rostral+ventrolateral+medulla+mediates+the+sympathoexcitatory+response+to+hyperinsulinemia.">Glutamatergic receptor activation in the rostral ventrolateral medulla mediates the sympathoexcitatory response to hyperinsulinemia.</a> Hypertension. 55 (2), 284-290 (2010).</li><li>Brozoski, D. T., Dean, C., Hopp, F. A., Seagard, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uptake+blockade+of+endocannabinoids+in+the+NTS+modulates+baroreflex-evoked+sympathoinhibition.">Uptake blockade of endocannabinoids in the NTS modulates baroreflex-evoked sympathoinhibition.</a> Brain Research. 1059 (2), 197-202 (2005).</li><li>Barman, S. 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. 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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 ...

This method can help answer key questions about the autonomic nervous system particularly about the central nervous system mechanisms that regulate sympathetic outflow to the kidney. The main advantage of this technique is that it allows measurement of the impact that central nervous system manipulations have on renal sympathetic outflow to the kidney. 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.For brain access, gently move the rat into the prone position in a stereotaxic surgery frame with the head between the ear bars and adjust the incisor bar to equalize the height of lambda and bregma. Next make a two centimeter rostral caudal incision through the midline of the scalp and use cotton tipped applicators to firmly remove any connective tissue from the skull surface. Apply hydrogen peroxide to the exposed skull to assist in visualizing the bregma, lambda and midline sutures.Using an atlas of the rat brain to guide targeting, drill a burr hole osteotomy sized for electrode access through the skull. To isolate the renal sympathetic nerves, connect a wire renal sympathetic nerve activity or RSNA electrode to a 10X pre amplifier and a microelectrode amplifier and make a scalpel incision extending from four to five 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 one to two centimeter rostral caudal 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 microdissecting 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 a 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 and 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 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.In this representative experiment, intravenous injection of phenylephrine was used to induce an increase in the mean arterial pressure and to evoke the baroreflex and transient sympathoinhibition. To quantify the RSNA the raw RSNA was rectified and averaged for non overlapping 10 second segments and the noise estimate was subtracted from each segment. When performing this procedure, it's important to remember to ensure that the rat is adequately anesthetized, that it is properly positioned in the stereotaxic frame and that the system is adequately grounded to reduce electrical noise.Following this procedure and collection of brain tissue, histology methods can be applied to characterize the types of central system nuclei that control a sympathoexcitatory response. This technique allow neuroscientists to explore autonomic regulation mechanisms in anesthetized rats which informs future studies in animals to understand the control of blood pressure and renal function. Don't forget that it is critical to assess the plane of anesthesia at least every 15 minutes.And to provide supplemental anesthesia as needed.

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Research

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Neuroscience

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 ...

Immunohistochemical Analysis in the Rat Central Nervous System and Peripheral Lymph Node Tissue Sections

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based multicolor labeling is challenging, especially when utilized for assessing interrelations between target proteins in the tissue with a high fat content such as the central nervous system (CNS).
Our protocol represents a refinement of the standard immunolabeling technique particularly adjusted for detection of both structural and soluble proteins in the rat CNS and peripheral lymph nodes (LN) affected by neuroinflammation. Nonetheless, with or without further modifications, our protocol could likely be used for detection of other related protein targets, even in other organs and species than here presented.

We here present an optimized, detailed protocol for double immunostaining in formalin-fixed, paraffin-embedded rat central nervous system (CNS) and peripheral lymph node (LN) tissue sections.

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based ...

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 ...