This study was supported by the Canadian Institutes of Health Research and Genome Canada.JS holds salary support awards from the Alberta Heritage Foundationfor Medical Research, Heart and Stroke Foundation of Canadaand the Canadian Diabetes Association. Special thanks to Dr. David Wasserman and Bingle Bracy for teaching this procedure to the Shearer laboratory.

A: Surgical Catheterization of Arterial and Venous Circulation
Part 1: Arterial and Venous Catheter Preparation
<ol>
	<li>Cut 15 cm of PE-50 (internal diameter of .58mm (.023") x an outer diameter of .965mm (.038"). Cut a 1mm section of silastic tubing (0.76mm (0.030") internal diameter x 1.65mm (0.065") outer diameter) for use as restraining bead. The restraining bead prevents the rat from pulling out the catheter once it is in place.</li>
	<li>Insert the tips of micro dissecting forceps into the lumen of the restraining bead and gently hold the tips of the forceps apart to stretch the opening wider. Using another pair of micro dissecting forceps, slide the silastic tubing into the lumen of the restraining bead. Secure with strong adhesive glue (Loctite Super Glue). The bead must lie flat around the catheter. Allow catheter to dry (24h). </li>
	<li>For a 300 gram rat, the artery line is ~2.7cm from the restraining bead while the jugular line is ~3.2cm. Catheter lengths from the restraining are adjusted by 0.5cm for every 100g increase in body weight. Do not bevel the edges of the catheter as this may puncture the vessel during insertion.</li>
	<li>Immediately prior to surgery, fill the lines with heparinized saline (10U/ml), seal both ends with a stainless steel tubing plug and place in ethanol (70%) to sterilize. Air dry prior to insertion. </li>
</ol>
Part 2: Surgical Preparation
<ol>
	<li>Procedures were approved by the University of Calgary Animal Care and Use Committee and abide by theCanadian Association for Laboratory Animal Science guidelines for experimentation. The procedures outlined below were performed on adult, male Sprague-Dawley rats (~300g). All procedures are performed under isoflurane, although it is possible to use injectable anesthetics. All surgical procedures are performed ensuring aseptic techniques. Surgical equipment, beakers and heparinized saline are autoclave sterilized. Surgical gloves, syringes and cotton tipped applicators are purchased sterilized from the supplier.</li>
	<li>Weigh the rat and record the result. The weight will be important in the post-surgical monitoring of animals. Only animals that regain weight to pre-surgery levels should be used for experimentation. Anesthetize rat (3% isoflurane) in an anesthetic box. Keep the rat at ~2% isoflurane during surgery using a nose cone. Surgery must be conducted in a disinfected area that promotes asepsis.</li>
	<li>Prepare the animal by removing hair from the surgical site (neck and shoulder blades). A small hair clipper or chemical depilatory can be used. Perform this procedure in an area separate from where the surgery is to be conducted.</li>
	<li>Secure animal to surgical table. Ensure animal is fully anesthetized by checking for the presence of foot/eye reflexes. Prepare the surgical sites with an appropriate skin disinfectant (70% ethanol followed by betadine scrub followed by 70% ethanol).</li>
</ol>
Part 3: Surgery
<ol>
	<li>Make small vertical midline incision 1cm superior to the sternum (Pick up skin longitudinally along medial axis with forceps and cut with scissor or scalpel).</li>
	<li>Blunt dissect using micro dissecting forceps to expose the left sternomastoid muscle. Reflect this muscle to expose approximately 1cm of the left carotid artery. Use micro dissecting forceps under carotid artery to hold the artery in place. Gently tease off connective tissue from the carotid artery. It is important to isolate the vagus nerve from the artery without damaging either the artery or the nerve. Isolate the artery then ligate the cephalad end with 4-0 silk suture (this will be used to manipulate the carotid artery during surgery). Note: 4-0 sutures should be sterilized with 70% ethanol prior to use.</li>
	<li>Clamp the vessel with serrated, micro dissecting forceps. Puncture the ligated end with a 21 gauge VENOJECT Multi-Sample Luer Adapter. Remove the stainless steel tubing plug and carefully insert catheter with the aid of a sterile catheter introducer to admit it into the artery. Ensuring the catheter is secure with forceps, partially release micro dissecting forceps clamping the artery and continue inserting the catheter to the bead. At this point, the tip of the catheter should be in the aortic arch. Take care not to release catheter, pressure of the vessel will force it out. </li>
	<li>Tie two 4-0 sutures securely below bead and one above and confirm that the catheter will sample. Flush the line with 10U/ml heparinized saline. Re-plug the external end of the catheter with a stainless steel tubing plug.</li>
	<li>Using the same incision, blunt dissect to expose right external jugular vein. Isolate carefully and ligate the cephalad end with silk suture. Puncture the vein with a 21 gauge VENOJECT Multi-Sample Luer Adapter. Remove the stainless steel tubing plug and insert catheter with sterile catheter introducer to the bead and tie two sutures below the bead. Tie a third suture above bead and confirm that it samples. Flush with the venous line 10U/mL heparinized saline. Re-plug the external catheter end with stainless steel tubing plug.</li>
	<li>Tunnel 14-gauge blunt needle under skin and make incision in the back between the shoulder blades. Thread the catheters through the needle to exteriorize them at the back of the rat. Approximately 4cm of line is visible. Cut 0.5cm of Tygon S-50-HL Medical Tubing, place it around both the exteriorized catheters and secure with a tape as required (blue for vein, red for artery). Retest catheters to ensure patency, flush and fill with 150U/ml heparinized saline to prevent clotting.</li>
	<li>Close all incisions with 3-0 silk suture. Place rat prone, in pre-warmed, clean cage with food in the bottom of the cage. </li>
</ol>
B: Postoperative Care
Part 1: Immediate Postoperative Care and Monitoring
<ol>
	<li>Once the rat regains full ambulatory ability and alertness return the rat to animal housing.</li>
	<li>Allow the rat to recover for 3-5 days. </li>
	<li>Monitor daily for infection, pain and changes in weight. Infection may be of concern if discharge from incision sites, general lethargy and/or pain is observed. Pain is indicated by hunched posture, ruffled back fur and absence of ambulatory and/or eating behavior. A loss in weight may be experienced immediately following surgery up to three days post-surgery, however, weight should stabilize and/or increase to within 10% of the pre-surgical weight within 5 days. Severe weight loss may indicate infection, stroke and/or pain. </li>
	<li>Retest catheters daily to ensure patency. </li>
</ol>
Part 2: Maintaining Catheter Patency
<ol>
	<li>Each day during post-op recovery, fill 1ml Slip Tip Syringes with 150U/ml heparinized saline and cap it with a 22 gauge blunt needle. The blunt needle is inserted into 20cm of PE-50 tubing with 23 gauge stainless steel tubing coupler at the other end.</li>
	<li>Remove air bubbles by placing the end with the tubing coupler higher than the rest of the 1ml syringe and pushing the bubbles out of the line. </li>
	<li>Clamp off the arterial line externalized from the rat with hemostats just below the plug.</li>
	<li>Remove the steel catheter plug using the second pair of hemostats. </li>
	<li>Insert the tubing coupler connected to the syringe into the arterial line and release the hemostats occluding the externalized arterial catheter.</li>
	<li>Aspirate the arterial blood of the catheter into syringe. If the catheter does not draw easily it may be necessary to very gently push in a small amount of flush solution through the catheter to dislodge the tip of the catheter in case it is wedged against the vessel wall. The catheter has been completely cleared when the blood reaches the syringe.</li>
	<li>Clamp the PE-50 tubing close to the 1ml syringe and dispose of the syringe. Replace with a new syringe filled with fresh 150U/ml heparinized saline. Unclamp the tubing and holding the new syringe upright, flick the new syringe with a finger to dislodge any possible new air bubbles upwards and away from the line. Note: Care must be ensured as air bubbles can cause a stroke.</li>
	<li>Inject the 150U/ml heparinzed saline until the catheter line is clear and free of blood. </li>
	<li>Clamp off the arterial line externalized from the rat with hemostats just below the tubing coupler. Remove the syringe connector from the arterial catheter line and replace it with the stainless steel tubing plug. </li>
	<li>Release the hemostats occluding the externalized arterial catheter. </li>
	<li>Repeat for venous line. However, because of the low venous pressure sampling is often not possible. If the 150U/ml heparinzed saline can be infused with minimal resistance it is probable that the catheter is well positioned in the vein.</li>
</ol>
C: Hyperinsulinemic-Euglycemic Clamp
Part 1: Clamp Set-up and Preparation
<ol>
	<li>Weigh the rat and record the weight. This will be required to determine insulin and glucose infusion rates. Fast the rat for at least 5h prior to experiment. This ensures the animal is in a postprandial state and the contribution of glucose from dietary sources is minimized. Place the rat in a small container/cage with bedding that limits excessive movement. The animal should be able to turn around and groom freely. </li>
	<li>Set up lines and infusion pumps as seen in Figure 1. P50 connection tubing lengths are as follows: syringe to Y connector is 8cm, Y connector to rat is 15cm, arterial line is 20cm, and venous line is 10cm in length. Remove stainless steel tubing plugs and flush jugular and arterial lines to ensure infusion and sampling respectively with 10U/ml heparinized saline. </li>
	<li>Determine insulin volume required. This will vary depending on the insulin level and weight of the rat. Here, 4mU/kg/min is administered at a rate of 2uL/min. This is a high physiological dose. Where possible, attempts should be made to limit infusion volumes. Insulin should be appropriately diluted and be administered in the presence of plasma. A 3% rat plasma solution in saline is used to dilute the insulin in this laboratory. Insulin is HumulinR 100U/ml (Eli Lily); although other fast acting insulin can be employed. Ensure diluted insulin is well mixed prior to putting in infusion syringe. </li>
	<li>Prepare 50% dextrose. This can be placed directly into syringe on the infusion pump. Mark pump with 'glucose' to avoid future confusion. Remember to ensure sufficient volumes of insulin and glucose for the study. Clamps generally last ~2h. In the present protocol, 3ml syringes are used. </li>
	<li>Place glucose and insulin syringes onto Harvard Apparatus Model 11 Plus Syringe Pumps. Advance glucose to Y-Connector and clamp line with a hemostat. Advance insulin to animal, clamp line with hemostat. Clamp lines and allow rat to relax for 30min prior to starting the experiment. NOTE: Hemostat tip guards are suggested in order to avoid permanent crimping of the lines.</li>
	<li>Prepare centrifuge and EDTA-coated tubes for plasma collection. In the present study, additional blood samples will be obtained once the animal is clamped. </li>
</ol>
Part 2: Experimental Protocol
<ol>
	<li>A baseline insulin sample and hematocrit sample should be acquired. Hematocrit sampling ensures blood volume is maintained throughout the experiment. Generally, hematocrit levels should not fall more than 10% of the initial value. </li>
	<li>Obtain a baseline glucose sample (One Touch Ultra, LifeScan, Inc.) and determine the whole blood glucose level to be clamped. Here, we clamp at euglycemia, or 5.0-5.5mM (Figure 2). After each blood sample, flush arterial line with a small volume of 10U/ml heparinized saline to prevent clotting. NOTE: Ensure no clots or air bubbles are in the line prior to infusing. These can give the animal a stroke. </li>
	<li>Start insulin infusion. Take blood glucose at 5-10min intervals, monitoring glucose at each time point. Adjust glucose infusion rate as required until a steady state is achieved. This process is generally a trial and error process and can take 30min to &gt;2h. The levels of glucose required to maintain glycemia are dependent upon the experimental protocol, species and conditions present (Figure 2).</li>
	<li>Steady state glucose levels are three consecutive readings within a defined range. Here, three readings within ~1mM is considered clamped (e.g. 4.8, 5.2, 5.6mM). Once steady state is achieved, record the glucose infusion rate required to maintain glycemia for a 30min period. During this time additional blood samples can be obtained. At minimum, a second plasma insulin sample and hematocrit sample should be obtained.</li>
	<li>Once clamp is completed, animals can be used for further testing or euthanized and tissues collected for further analysis. Depending on the animal, catheter lines can stay patent for 5-7 days.</li>
</ol>
D: Hyperinsulinemic-Euglycemic (Insulin) Clamp Results
When performed correctly, the clamp procedure assesses the steady state insulin sensitivity of the rat. In presenting data obtained from the clamp, it is essential to document glucose levels and glucose infusion rates. Stable glucose levels over a time course of a minimum of 30min (Figure 3) are indicative of a steady state. Glucose is considered stable only if whole blood glucose is maintained within ~1mM. Glucose infusion rates show the levels of exogenous glucose required to maintain glycemia. Where possible, these figures should be shown as a time course rather than single, averaged value (Figure 4). 
Other recommended measures to be reported are plasma insulin and hematocrit. The determination of both fasting and clamped insulin levels confirm that insulin has been successfully administered and will detect any differences in levels between treatment groups (Figure 5). Obtaining hematocrit measures at baseline and at the conclusion of the insulin clamp are suggested (Figure 6). This is to ensure hematocrit levels do not fall more than 5% during the experiment and the accompanying alterations in blood volume and flow do not influence glucose disposal.
<img src="/files/ftp_upload/2432/2432fig1.jpg" alt="Figure 1" /> Figure 1. Experimental set-up. Figure 1 shows unrestrained, conscious rat during the clamp procedure. Catheters allow blood sampling and infusions without handling the animal. Pumps to the left contain insulin and glucose.
<img src="/files/ftp_upload/2432/2432fig2.jpg" alt="Figure 2" /> Figure 2. Expected data during the clamp procedure. To obtain a clamp at baseline levels (euglycemia, 5mM), the levels of exogenous glucose (D50) are manipulated until the baseline or 'clamp' is achieved.
<img src="/files/ftp_upload/2432/2432fig3.jpg" alt="Figure 3" /> Figure 3. Expected plasma glucose results of the clamp procedure. When the animal is 'clamped', blood glucose is relatively stable across time and experimental groups.
<img src="/files/ftp_upload/2432/2432fig4.jpg" alt="Figure 1" /> Figure 4. Representative glucose infusion rates of the clamp procedure. The amount of exogenous glucose required to maintain euglycemia differs. This is illustrated with a control (chow fed) and high fat fed (insulin resistant) animals. The high fat fed animal requires less glucose infused to maintain glycemia, primarily because it is insensitive to the insulin infused.
<img src="/files/ftp_upload/2432/2432fig5.jpg" alt="Figure 5" /> Figure 5. Representative plasma insulin of the rat. The plasma insulin during the insulin clamp should be higher than the fasted, baseline plasma insulin. This ensures insulin was properly administered to the animal during the insulin clamp.
<img src="/files/ftp_upload/2432/2432fig6.jpg" alt="Figure 6" /> Figure 6. Reporting hematocrit. The baseline hematocrit and hematocrit following the experiment must be obtained and reported. This ensures hematocrit levels do not fall more than 5% of baseline levels resulting from excessive arterial blood sampling.

<ol>
	<li>Halseth, A. E., Bracy, D. P., Wasserman, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Limitations+to+basal+and+insulin-stimulated+skeletal+muscle+glucose+uptake+in+the+high-fat-fed+rat.">Limitations to basal and insulin-stimulated skeletal muscle glucose uptake in the high-fat-fed rat.</a> Am J Physiol Endocrinol Metab. 279, E1064-E1071 (2000).</li><li>Halseth, A. E., Bracy, D. P., Wasserman, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Limitations+to+muscle+glucose+uptake+due+to+high+fat+feeding.">Limitations to muscle glucose uptake due to high fat feeding.</a> Am. J. Physiol. 279, E1064-E1071 (2000).</li><li>Muniyappa, R., Lee, S., Chen, H., Quon, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Current+approaches+for+assessing+insulin+sensitivity+and+resistance+in+vivo:+advantages,+limitations,+and+appropriate+usage.">Current approaches for assessing insulin sensitivity and resistance in vivo: advantages, limitations, and appropriate usage.</a> Am J Physiol Endocrinol Metab. 294, 15-26 (2008).</li><li>Cho, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insulin+resistance+and+a+diabetes+mellitus-like+syndrome+in+mice+lacking+the+protein+kinase+Akt2+(PKB+beta).">Insulin resistance and a diabetes mellitus-like syndrome in mice lacking the protein kinase Akt2 (PKB beta).</a> Science. 292, 1728-1731 (2001).</li><li>Dubois, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+SHP-1+protein+tyrosine+phosphatase+negatively+modulates+glucose+homeostasis.">The SHP-1 protein tyrosine phosphatase negatively modulates glucose homeostasis.</a> Nat Med. 12, 549-556 (2006).</li><li>Pacini, G., Thomaseth, K., Ahren, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contribution+to+glucose+tolerance+of+insulin-independent+vs.+insulin-dependent+mechanisms+in+mice.">Contribution to glucose tolerance of insulin-independent vs. insulin-dependent mechanisms in mice.</a> Am J Physiol Endocrinol Metab. 281, 693-703 (2001).</li><li>Herbach, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dominant-negative+effects+of+a+novel+mutated+Ins2+allele+causes+early-onset+diabetes+and+severe+beta-cell+loss+in+Munich+Ins2C95S+mutant+mice.">Dominant-negative effects of a novel mutated Ins2 allele causes early-onset diabetes and severe beta-cell loss in Munich Ins2C95S mutant mice.</a> Diabetes. 56, 1268-1276 (2007).</li><li>Maeda, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diet-induced+insulin+resistance+in+mice+lacking+adiponectin/ACRP30.">Diet-induced insulin resistance in mice lacking adiponectin/ACRP30.</a> Nat Med. 8, 731-737 (2002).</li><li>Potenza, M. A., Marasciulo, F. L., Tarquinio, M., Quon, M. J., Montagnani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Treatment+of+spontaneously+hypertensive+rats+with+rosiglitazone+and/or+enalapril+restores+balance+between+vasodilator+and+vasoconstrictor+actions+of+insulin+with+simultaneous+improvement+in+hypertension+and+insulin+resistance.">Treatment of spontaneously hypertensive rats with rosiglitazone and/or enalapril restores balance between vasodilator and vasoconstrictor actions of insulin with simultaneous improvement in hypertension and insulin resistance.</a> Diabetes. 55, 3594-3603 (2006).</li><li>Ayala, J. E., Bracy, D. P., McGuinness, O. P., Wasserman, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Considerations+in+the+design+of+hyperinsulinemic-euglycemic+clamps+in+the+conscious+mouse.">Considerations in the design of hyperinsulinemic-euglycemic clamps in the conscious mouse.</a> Diabetes. 55, 390-397 (2006).</li><li>Wasserman, D. H., Ayala, J. E., McGuinness, O. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lost+in+translation.">Lost in translation.</a> Diabetes. 58, 1947-1950 (2009).</li></ol>

Procedures were approved by the University of Calgary Animal Care and Use Committee and abide by the Canadian Association for Laboratory Animal Science guidelines for experimentation. The authors have no conflicting interests or disclosures.

Initially developed for investigation of insulin sensitivity in humans, the clamp procedure has now been adapted to other species including laboratory rats and mice. Investigating animal models of insulin resistance provides a significant aid in understanding the pathophysiology of insulin sensitivity and associated pathologies as well as identifying therapeutic interventions that have clinical value1,2. Several methods to evaluate insulin sensitivity in animals have been employed3. Such techniques ...

<table border="1">
	<tbody>
		<tr valign="top">
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<tr valign="top">
			<td>Intramedic Polyethylene Tubing (PE-50)</td>
			<td>Fisher Scientific</td>
			<td>14-170-12B</td>
			<td>Internal diameter of .58mm (.023") x Outer diameter of .965mm (.038")</td>
		</tr>
		<tr valign="top">
			<td>Dow Corning Silastic Laboratory Tubing</td>
			<td>Fisher Scientific</td>
			<td>11-189-15C</td>
			<td>Internal diameter of .76mm (0.030") x Outer diameter of 1.65mm (0.065") </td>
		</tr>
		<tr valign="top">
			<td>Tygon S-50-HL Medical Tubing</td>
			<td>Harvard Apparatus</td>
			<td>PY2 72-1251</td>
			<td>Internal diameter of 3.2mm (0.125") x Outer diameter of 4.7mm (0.1875")</td>
		</tr>
		<tr valign="top">
			<td>Loctite Super Glue</td>
			<td>Grand &amp; Toy</td>
			<td>32237</td>
			<td>Gel Control</td>
		</tr>
		<tr valign="top">
			<td>Sterile Surgical Blade</td>
			<td>VWR</td>
			<td>BD371610</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Curved Micro Dissecting Forceps</td>
			<td>George Tiemann &amp; Co.</td>
			<td>160-20</td>
			<td>x 2</td>
		</tr>
		<tr valign="top">
			<td>Straight Micro Dissecting Forceps</td>
			<td>George Tiemann &amp; Co.</td>
			<td>160-15</td>
			<td>x 2</td>
		</tr>
		<tr valign="top">
			<td>Curved Hemostat</td>
			<td>George Tiemann &amp; Co.</td>
			<td>105-1135</td>
			<td>x 2</td>
		</tr>
		<tr valign="top">
			<td>Straight Hemostat</td>
			<td>George Tiemann &amp; Co.</td>
			<td>105-1130</td>
			<td>x 2</td>
		</tr>
		<tr valign="top">
			<td>Hemostat Tip Guards</td>
			<td>Robbins Instruments, Inc.</td>
			<td>15.09-2-004</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Straight Surgery Scissors</td>
			<td>George Tiemann &amp; Co.</td>
			<td>105-402</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>VENOJECT Multi-Sample Luer Adapter</td>
			<td>Terumo Medical Products</td>
			<td>810127A</td>
			<td>21 guage, 1 in.</td>
		</tr>
		<tr valign="top">
			<td>Sterile Catheter Introducer</td>
			<td>Becton Dickinson</td>
			<td>406999</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>14-gauge Blunt Needle</td>
			<td>Becton Dickinson</td>
			<td>511310</td>
			<td>14 guage, 2 in.</td>
		</tr>
		<tr valign="top">
			<td>Sterile Surgical Suture</td>
			<td>Johnson &amp; Johnson Medical Products</td>
			<td>1679H</td>
			<td>Silk, size 3-0</td>
		</tr>
		<tr valign="top">
			<td>Non-Sterile Surgical Suture </td>
			<td>Angiotech Pharmaceuticals, Inc.</td>
			<td>SP116</td>
			<td>Silk, size 4-0</td>
		</tr>
		<tr valign="top">
			<td>Cotton Swabs</td>
			<td>VWR</td>
			<td>10806-005</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>4ply Gauze Pads</td>
			<td>VWR</td>
			<td>CA43845-062</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Small Animal Cordless Clippers</td>
			<td>Harvard Apparatus</td>
			<td>729063</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Isoflurane</td>
			<td>Halocarbon Products Corp.</td>
			<td>IPN-45</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Anesthetic Cart</td>
			<td>Benson Medical Industries, Inc.</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>70 % Ethanol</td>
			<td>Fisher Scientific</td>
			<td>HC-1000</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Betadine Antiseptic Solution</td>
			<td>Western Drug Distribution Centre Ltd.</td>
			<td>105267</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Model 11 Plus Syringe Pump</td>
			<td>Harvard Apparatus</td>
			<td>702208</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Stainless Steel Tubing Couplers</td>
			<td>Harvard Apparatus</td>
			<td>72-4434</td>
			<td>23 gauge, 0.3 in.</td>
		</tr>
		<tr valign="top">
			<td>Stainless Steel Tubing Plugs</td>
			<td>Harvard Apparatus</td>
			<td>72-4436</td>
			<td>23 gauge, 0.5 in.</td>
		</tr>
		<tr valign="top">
			<td>Stainless Steel Blunt Needles</td>
			<td>Instech Laboratories, Inc.</td>
			<td>LS22</td>
			<td>22 gauge</td>
		</tr>
		<tr valign="top">
			<td>60 Degree Y-Connectors</td>
			<td>Small Parts</td>
			<td>STCY-22-05</td>
			<td>22 gauge</td>
		</tr>
		<tr valign="top">
			<td>CritSpin Micro-hematocrit Centrifuge</td>
			<td>Iris Sample Processing</td>
			<td>CS12</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Mini Centrifuge</td>
			<td>Fisher Scientific</td>
			<td>05-090-100</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Micro Centrifuge Tubes</td>
			<td>VWR</td>
			<td>53550-778</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>50ml polypropylene centrifuge tubes</td>
			<td>VWR</td>
			<td>89004-364</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>1ml Plastic Slip Tip Syringes</td>
			<td>Becton Dickinson</td>
			<td>309602</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>3ml Plastic Luerlok Tip Syringes</td>
			<td>Becton Dickinson</td>
			<td>309585</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Heparin Anticoagulant Injection</td>
			<td>Western Drug Distribution Centre Ltd.</td>
			<td>102824</td>
			<td>Manufacturer: LEO Pharma Inc.
				Conc. 1000 IU</td>
		</tr>
		<tr valign="top">
			<td>EDTA Solution</td>
			<td>Promega Corp.</td>
			<td>V4231</td>
			<td>0.5 M, pH 8.0</td>
		</tr>
		<tr valign="top">
			<td>Saline</td>
			<td>Western Drug Distribution Centre Ltd.</td>
			<td>ABB7983154</td>
			<td>Manufacturer: Hospira
				0.9% Sodium Chloride</td>
		</tr>
		<tr valign="top">
			<td>50% Dextrose</td>
			<td>Vetoquinol</td>
			<td>8DEX012D</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Humulin-R </td>
			<td>Eli Lilly</td>
			<td>HI-210</td>
			<td>100U/ml</td>
		</tr>
		<tr valign="top">
			<td>1ml Insulin Syringes </td>
			<td>Becton Dickinson</td>
			<td>309311</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Fisherbrand* Hemato-Seal Sealant</td>
			<td>Fisher Scientific</td>
			<td>02-678</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Fisherbrand* Microhematocrit Capillary Tubes</td>
			<td>Fisher Scientific</td>
			<td>22-362-574</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>One Touch Ultra Test Strips</td>
			<td>LifeScan, Inc.</td>
			<td>AW 085-314H</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>One Touch Ultra Blood Glucose Meter</td>
			<td>LifeScan, Inc.</td>
			<td>AW 085-314B</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Sodium Pentobarbitol</td>
			<td>Ceva Sante Animale</td>
			<td>1715 138</td>
			<td>Conc. 54.7mg/ml</td>
		</tr>
		<tr valign="top">
			<td>Red Laboratory Labeling Tape</td>
			<td>VWR</td>
			<td>89097-932</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Blue Laboratory Labeling Tape</td>
			<td>VWR</td>
			<td>89097-936</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Weigh Scale</td>
			<td>Fisher Scientific</td>
			<td>01-913-88</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Vortex</td>
			<td>VWR</td>
			<td>58815-234</td>
			<td>&nbsp;</td>
		</tr>
		<tr valign="top">
			<td>Timer</td>
			<td>VWR</td>
			<td>62344-641</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

hyperinsulinemic-euglycemic clamp in the conscious rat

Type 2 diabetes (T2D) is rapidly rising in prevalence. Characterized by either inadequate insulin production or the inability to utilize insulin produced, T2D results in elevated blood glucose levels. The "gold-standard" in assessing insulin sensitivity is a hyperinsulinemic-euglycemic clamp or insulin clamp. In this procedure, insulin is infused at a constant rate resulting in a drop in blood glucose. To maintain blood glucose at a constant level, exogenous glucose (D50) is infused into the venous circulation. The amount of glucose infused to maintain homeostasis is indicative of insulin sensitivity. Here, we show the basic clamp procedure in the chronically catheterized, unrestrained, conscious rat. This model allows blood to be collected with minimal stress to the animal. Following the induction of anesthesia, a midline incision is made and the left common carotid artery and right jugular vein are catheterized. Inserted catheters are flushed with heparinized saline, then exteriorized and secured. Animals are allowed to recover for 4-5 days prior to experiments, with weight gain monitored daily. Only those animals who regain weight to pre-surgery levels are used for experiments. On the day of the experiment, rats are fasted and connected to pumps containing insulin and D50. Baseline glucose is assessed from the arterial line and used a benchmark throughout the experiment (euglycemia). Following this, insulin is infused at a constant rate into the venous circulation. To match the drop in blood glucose, D50 is infused. If the rate of D50 infusion is greater than the rate of uptake, a rise in glucose will occur. Similarly, if the rate is insufficient to match whole body glucose uptake, a drop will occur. Titration of glucose continues until stable glucose readings are achieved. Glucose levels and glucose infusion rates during this stable period are recorded and reported. Results provide an index of whole body insulin sensitivity. The technique can be refined to meet specific experimental requirements. It is further enhanced by the use of radioactive tracers that can determine tissue specific insulin-stimulated glucose uptake as well as whole body glucose turnover.

Watch this Scientific Journal Video about Hyperinsulinemic-Euglycemic Clamp in the Conscious Rat at JoVE.com

Hyperinsulinemic-Euglycemic Clamp in the Conscious Rat

The hyperinsulinemic-euglycemic clamp is the "gold standard" for the assessment of insulin action. Insulin is infused at a constant rate stimulating glucose uptake. The amount of exogenous glucose infused to counter this drop is indicative of insulin sensitivity. Here the procedure is performed on a conscious, unrestrained rat.

Type 2 diabetes (T2D) is rapidly rising in prevalence. Characterized by either inadequate insulin production or the inability to utilize insulin produced, ...

hyperinsulinemic-euglycemic-clamp-in-the-conscious-rat

Research

JoVE Journal

Medicine

51.9K Views.  University of Calgary. The hyperinsulinemic-euglycemic clamp is the "gold standard" for the assessment of insulin action. Insulin is infused at a constant rate stimulating glucose uptake. The amount of exogenous glucose infused to counter this drop is indicative of insulin sensitivity. Here the procedure is performed on a conscious, unrestrained rat.

Video: Hyperinsulinemic-Euglycemic Clamp in the Conscious Rat

Hyperinsulinemic-euglycemic Clamps in Conscious, Unrestrained Mice

Type 2 diabetes is characterized by a defect in insulin action. The hyperinsulinemic-euglycemic clamp, or insulin clamp, is widely considered the "gold standard" method for assessing insulin action in vivo. During an insulin clamp, hyperinsulinemia is achieved by a constant insulin infusion. Euglycemia is maintained via a concomitant glucose infusion at a variable rate. This variable glucose infusion rate (GIR) is determined by measuring blood glucose at brief intervals throughout the experiment and adjusting the GIR accordingly. The GIR is indicative of whole-body insulin action, as mice with enhanced insulin action require a greater GIR. The insulin clamp can incorporate administration of isotopic 2[14C]deoxyglucose to assess tissue-specific glucose uptake and [3-3H]glucose to assess the ability of insulin to suppress the rate of endogenous glucose appearance (endoRa), a marker of hepatic glucose production, and to stimulate the rate of whole-body glucose disappearance (Rd).
The miniaturization of the insulin clamp for use in genetic mouse models of metabolic disease has led to significant advances in diabetes research. Methods for performing insulin clamps vary between laboratories. It is important to note that the manner in which an insulin clamp is performed can significantly affect the results obtained. We have published a comprehensive assessment of different approaches to performing insulin clamps in conscious mice1 as well as an evaluation of the metabolic response of four commonly used inbred mouse strains using various clamp techniques2. Here we present a protocol for performing insulin clamps on conscious, unrestrained mice developed by the Vanderbilt Mouse Metabolic Phenotyping Center (MMPC; URL: www.mc.vanderbilt.edu/mmpc). This includes a description of the method for implanting catheters used during the insulin clamp. The protocol employed by the Vanderbilt MMPC utilizes a unique two-catheter system3. One catheter is inserted into the jugular vein for infusions. A second catheter is inserted into the carotid artery, which allows for blood sampling without the need to restrain or handle the mouse. This technique provides a significant advantage to the most common method for obtaining blood samples during insulin clamps which is to sample from the severed tip of the tail. Unlike this latter method, sampling from an arterial catheter is not stressful to the mouse1. We also describe methods for using isotopic tracer infusions to assess tissue-specific insulin action. We also provide guidelines for the appropriate presentation of results obtained from insulin clamps.

The hyperinsulinemic-euglycemic clamp, or insulin clamp, is the gold standard for assessing insulin action in vivo. A method for performing insulin clamps in mice is described. This includes a method for arterial catheterization that permits experiments to be performed in conscious, unrestrained mice with minimal stress.

Type 2 diabetes is characterized by a defect in insulin action. The hyperinsulinemic-euglycemic clamp, or insulin clamp, is widely considered the "gold ...

Angiogenesis in the Ischemic Rat Lung

The adult lung is perfused by both the systemic bronchial artery and the entire venous return flowing through the pulmonary arteries. In most lung pathologies, it is the smaller systemic vasculature that responds to a need for enhanced lung perfusion and shows robust neovascularization. Pulmonary vascular ischemia induced by pulmonary artery obstruction has been shown to result in rapid systemic arterial angiogenesis in man as well as in several animal models. Although the histologic assessment of the time course of bronchial artery proliferation in rats was carefully described by Weibel 1, mechanisms responsible for this organized growth of new vessels are not clear. We provide surgical details of inducing left pulmonary artery ischemia in the rat that leads to bronchial neovascularization. Quantification of the extent of angiogenesis presents an additional challenge due to the presence of the two vascular beds within the lung. Methods to determine functional angiogenesis based on labeled microsphere injections are provided.

The lung is perfused by both the systemic bronchial artery and pulmonary arteries. In most lung pathologies, it is the smaller systemic vasculature that shows robust neovascularization. Cessation of pulmonary blood flow promotes brisk bronchial angiogenesis. We provide surgical details of inducing left pulmonary artery ischemia that promotes bronchial neovascularization.

The  adult lung is perfused by both the systemic bronchial artery and the entire  venous return flowing through the pulmonary arteries. In most lung ...

Slow-release Drug Delivery through Elvax 40W to the Rat Retina: Implications for the Treatment of Chronic Conditions

Diseases of the retina are difficult to treat as the retina lies deep within the eye. Invasive methods of drug delivery are often needed to treat these diseases. Chronic retinal diseases such as retinal oedema or neovascularization usually require multiple intraocular injections to effectively treat the condition. However, the risks associated with these injections increase with repeated delivery of the drug. Therefore, alternative delivery methods need to be established in order to minimize the risks of reinjection. Several other investigations have developed methods to deliver drugs over extended time, through materials capable of releasing chemicals slowly into the eye. In this investigation, we outline the use of Elvax 40W, a copolymer resin, to act as a vehicle for drug delivery to the adult rat retina. The resin is made and loaded with the drug. The drug-resin complex is then implanted into the vitreous cavity, where it will slowly release the drug over time. This method was tested using 2-amino-4-phosphonobutyrate (APB), a glutamate analogue that blocks the light response of the retina. It was demonstrated that the APB was slowly released from the resin, and was able to block the retinal response by 7 days after implantation. This indicates that slow-release drug delivery using this copolymer resin is effective for treating the retina, and could be used therapeutically with further testing.

This paper details how Elvax 40W can be used as a slow-release method for drug delivery to the adult rat retina. The protocol for preparing, loading, and delivering the drug-resin complex to the eye is described.

Diseases of the retina are difficult to treat as the retina lies deep within the eye. Invasive methods of drug delivery are often needed to treat these ...

Catheterization of the Carotid Artery and Jugular Vein to Perform Hemodynamic Measures, Infusions and Blood Sampling in a Conscious Rat Model

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. Intra-tracheal inoculation of a known amount of bacteria has been successfully used to reproduce the pathogenesis of pneumonia which then develops into sepsis. Monitoring hemodynamic parameters and providing standard clinical treatment including infusion of antibiotics, fluids and drugs to maintain blood pressure is critical to simulate routine supportive care in this model but to do so requires both arterial and venous vascular access. The video details the surgical technique for implanting carotid artery and common jugular vein catheters in an anesthetized rat. Following a 72 hr recovery period, the animals will be re-anesthetized and connected to a tether and swivel setup attached to the rodent housing which connects the implanted catheters to the hemodynamic monitoring system. This setup allows free movement of the rat during the study while continuously monitoring pressures, infusing fluids and drugs (antibiotics, vasopressors) and performing blood sampling.

Vascular accesses to measure hemodynamics, provide fluids and perform blood sampling are important to any small animal model study. We present a technique for implanting catheters into the carotid artery and the common jugular vein in an anesthetized rat for connecting to a system to perform monitoring, infusions and sampling.

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. ...

Transcutaneous Assessment of Renal Function in Conscious Rodents

Glomerular filtration rate (GFR) is the gold standard to assess overall kidney function. However, traditional methods to evaluate GFR are cumbersome and time-consuming. In addition, serial blood or urine samples are required, with the associated stress for the experimental animals. A recent technique significantly reduces the investment in time and resources, minimizing the invasiveness and the animal stress, but being equally valid as the traditional approaches. The method measures transcutaneously renal function. Using an optical device and the exogenous renal marker fluorescein isothiocyanate (FITC)-sinistrin, this technique is capable of measuring the elimination kinetics of the marker through the skin. With neither blood nor urine samples nor the associated laboratory assays needed, the results of the transcutaneous measurement are almost instantaneously available. The method has been already validated in different species and successfully applied in several models of renal pathology. Moreover, due to its minimally invasive characteristics, it is suitable for sequential measurements within the same animal. Here is provided a detailed protocol to carry out the transcutaneous assessment of renal function in rodents.

Determination of glomerular filtration rate (GFR) is the gold standard to assess overall kidney function. However, traditional procedures to measure this parameter are cumbersome and require a large investment of time. Here we describe a faster and minimally invasive method to determinate GFR transcutaneously.

Glomerular filtration rate (GFR) is the gold standard to assess overall kidney function. However, traditional methods to evaluate GFR are cumbersome and ...

The Dimethylnitrosamine Induced Liver Fibrosis Model in the Rat

Four to six week old, male Wistar rats were used to produce animal models of liver fibrosis. The process requires four weeks of administration of 10 mg/kg dimethylnitrosamine (DMN), given intraperitoneally for three consecutive days per week. Intraperitoneal injections were performed in the fume hood as DMN is a known hepatoxin and carcinogen. The model has several advantages. Firstly, liver changes can be studied sequentially or at particular stages of interest. Secondly, the stage of liver disease can be monitored by measurement of serum alanine aminotransferase (ALT) and aspartate aminotransferase (AST) enzymes. Thirdly, the severity of liver damage at different stages can be confirmed by sacrifice of animals at designated time points, followed by histological examination of Masson&#39;s Trichome stained liver tissues. After four weeks of DMN dosing, the typical fibrosis score is 5 to 6 on the Ishak scale. The model can be reproduced consistently and has been widely used to assess the efficacy of potential anti-fibrotic agents.

We describe a method to produce an animal model of liver fibrosis in the rat, and assess the degree of fibrosis by histological examination of the liver. The model can be used to study the development of liver disease as well as to test the efficacy of potential anti-fibrotic agents.

Four to six week old, male Wistar rats were used to produce animal models of liver fibrosis. The process requires four weeks of administration of 10 mg/kg ...

Method of Direct Segmental Intra-hepatic Delivery Using a Rat Liver Hilar Clamp Model

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ischemia/reperfusion (I/R) injury with myriad negative consequences. Potential I/R injury in marginal organs destined for liver transplantation contributes to the current donor shortage secondary to a decreased organ utilization rate. A significant need exists to explore hepatic I/R injury in order to mediate its impact on graft function in transplantation. Rat liver hilar clamp models are used to investigate the impact of different molecules on hepatic I/R injury. Depending on the model, these molecules have been delivered using inhalation, epidural infusion, intraperitoneal injection, intravenous administration or injection into the peripheral superior mesenteric vein. A rat liver hilar clamp model has been developed for use in studying the impact of pharmacologic molecules in ameliorating I/R injury. The described model for rat liver hilar clamp includes direct cannulation of the portal supply to the ischemic hepatic segment via a side branch of the portal vein, allowing for direct segmental hepatic delivery. Our approach is to induce ischemia in the left lateral and median lobes for 60 min, during which time the substance under study is infused. In this case, pegylated-superoxide dismutase (PEG-SOD), a free radical scavenger, is infused directly into the ischemic segment. This series of experiments demonstrates that infusion of PEG-SOD is protective against hepatic I/R injury. Advantages of this approach include direct injection of the molecule into the ischemic segment with consequent decrease in volume of distribution and reduction in systemic side effects.

A unique rat liver hilar clamp model was developed for studying the impact of pharmacologic molecules in ameliorating ischemia-reperfusion injury. This model includes direct cannulation of the portal supply to the ischemic liver segment via a branch of the portal vein, allowing for direct hepatic delivery.

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ...

Re-Arterialized Rat Partial Liver Transplantation with an in vivo Vessel-Oriented 70% Hepatectomy

Split liver transplantation and living liver donor liver transplantation were developed in the clinic to utilize liver organs in a more efficient manner. To better understand the mechanism behind these surgical procedures, a rat partial liver transplantation (PLTx) model was established for relevant surgical studies. Because of the complexity of the rat PLTx model, a protocol with detailed descriptions is required. An article published previously reported a protocol in which ex vivo hepatectomy was used to achieve 50% rat PLTx. In contrast to this protocol, we introduced a re-arterialized PLTx with an in vivo 70% hepatectomy. An updated vessel-oriented hepatectomy was incorporated into the rat PLTx to refine the microsurgical procedure. The portal veins and hepatic arteries of the left lateral lobe and the median lobe were individually dissected and ligated before removal of the liver parenchyma, thereby decreasing the probability of bleeding in the remnant liver stump. Furthermore, an end-to-side vessel anastomosis between the common hepatic artery and the enlarged proper hepatic artery was introduced to re-arterialize the hepatic artery. By using this end-to-side vessel anastomosis technique, the diameter of the anastomosis was enlarged, thereby decreasing the difficulty of hand suture and maintaining a high rate of anastomotic patency. Moreover, the cuff anastomosis of the infrahepatic vena cava was slightly modified. A section of circumferential liver parenchyma around the vena cava of a recipient was preserved during cuff anastomosis to maintain the three-dimensional shape of the vascular lumen. This section of liver parenchyma was removed after completing the anastomosis. With this modification, the step involving placement of stay sutures was omitted, thereby further shortening the cuff anastomosis time. By using this protocol of rat PLTx, a low liver enzyme level, an intact liver lobular architecture and a high survival rate were achieved after microsurgery.

Re-Arterialized Rat Partial Liver Transplantation with an in vivo Vessel-Oriented 70% Hepatectomy

Here, a protocol involving re-arterialized rat partial liver transplantation is presented. Specifically, 70% liver was resected in vivo by using an updated technique of vessel-oriented hepatectomy. The hepatic artery was reconstructed in an end-to-side manner. The cuff technique was modified to shorten the anastomosis time of the infrahepatic vena cava.

Split liver transplantation and living liver donor liver transplantation were developed in the clinic to utilize liver organs in a more efficient manner. ...

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to non-invasively assess some features of the respiratory function in humans as well as in laboratory animals. The present article focuses on the application of the double-chamber plethysmography (DCP) in small animals. It provides background information on the methodology as well as a detailed step-by-step procedure to successfully assess respiratory function in conscious, spontaneously breathing animals in a non-invasive manner. The DCP can be used to monitor the respiratory function of multiple animals in parallel, as well as to identify changes induced by aerosolized substances over a chosen time period and in a repeated manner. Experiments on control and allergic mice are used herein to demonstrate the utility of the technique, explain the associated outcome parameters, as well as to discuss the related advantages and shortcomings. Overall, the DCP provides valid and theoretically sound readouts that can be trusted to evaluate the respiratory function of conscious small animals both at baseline and after challenges with aerosolized substances.

The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to ...

Multi-system Monitoring for Identification of Seizures, Arrhythmias and Apnea in Conscious Restrained Rabbits

Patients with ion channelopathies are at a high risk of developing seizures and fatal cardiac arrhythmias. There is a higher prevalence of heart disease and arrhythmias in people with epilepsy (i.e., epileptic heart.) Additionally, cardiac and autonomic disturbances have been reported surrounding seizures. 1:1,000 epilepsy patients/year die of sudden unexpected death in epilepsy (SUDEP). The mechanisms for SUDEP remain incompletely understood. Electroencephalograms (EEG) and electrocardiograms (ECG) are two techniques routinely used in the clinical setting to detect and study the substrates/triggers for seizures and arrhythmias. While many studies and descriptions of this methodology are in rodents, their cardiac electrical activity differs significantly from humans. This article provides a description of a non-invasive method for recording simultaneous video-EEG-ECG-oximetry-capnography in conscious rabbits. As cardiac electrical function is similar in rabbits and humans, rabbits provide an excellent model of translational diagnostic and therapeutic studies. In addition to outlining the methodology for data acquisition, we discuss the analytical approaches for examining neuro-cardiac electrical function and pathology in rabbits. This includes arrhythmia detection, spectral analysis of EEG and a seizure scale developed for restrained rabbits.

Using simultaneous video-EEG-ECG-oximetry-capnography, we developed a methodology to evaluate the susceptibility of rabbit models to develop provoked arrhythmias and seizures. This novel recording system establishes a platform to test the efficacy and safety of therapeutics and can capture the complex cascade of multi-system events that culminate in sudden death.

Patients with ion channelopathies are at a high risk of developing seizures and fatal cardiac arrhythmias. There is a higher prevalence of heart disease ...