This work was supported by the Leading Initiative for Excellent Young Researchers (from MEXT); a Grant-in-Aid for Scientific Research (B) (Grant Number JP21H02352); Japan Agency for Medical Research and Development (AMED-RPIME, Grant Number JP21gm6510009h0001, JP22gm6510009h9901); the Uehara Memorial Foundation; Astellas Foundation for Research on Metabolic Disorders; Suzuken Memorial Foundation, Akiyama Life Science Foundation, and Narishige Neuroscience Research Foundation. We also thank Nur Farehan Asgar, Ph.D, for editing a draft of this manuscript.

All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at Kumamoto University.
NOTE: For pain relief, ibuprofen was given in drinking water (0.11 mg/mL) for 48 h, and buprenorphine (0.05-0.1 mg/kg i.p.) was given 30 min before surgery. Sterile conditions include gloves, masks, and autoclaved instruments sterilized with ethylene oxide between animals. The surgery was performed on a heating pad set at 37 &#176;C and covered by a new lab mat for each animal. Before the surgery, the surgical area was cleaned with a betadine&#160;solution and alcohol. All surgical instruments were sterilized with an autoclave (for no more than two surgeries). Before making the incision, mice were checked to ensure they were fully anesthetized. The depth of anesthesia for each mouse was assessed prior to and during surgery by a toe pinch.&#160;The acclimatization period was no more than 5 min each time. Follow the instructions of the IACUC at the respective institution.
1. Preparation of tubings for the jugular vein and carotid artery
<ol>
	<li>Sterilize all tubings, polypropylene supplies (e.g., pipet tips), and sutures with an autoclave or ethylene oxide. For the jugular vein, connect 8 cm of tubing1 (see Materials) and 3 cm of tubing2 using glue (Figure 2A). Put 2 mm of tubing1 on the top because tubing 2 (polyethylene) is too hard and may damage blood vessels (Tubing1.1).</li>
	<li>Extension tube for the jugular vein (Tubing1.2) to infuse glucose and insulin
	<ol>
		<li>Prepare two 30 cm tubes and one 10 cm tube with tubing2 (Figure 2A). Cut off the sharp end of a 20 &#956;L pipette tip (or anything that has a narrow tip end with OD &#60; 0.5 mm to connect with tubing 1.1) and place three tubing2 (30 cm, 30 cm, and 10 cm) together in it. Seal with adhesive glue.</li>
		<li>Place 5 cm of tubing1 at the other end of tubing2.</li>
	</ol>
	</li>
	<li>For carotid artery&#160;(Tubing1.3): Stretch tubing2 to make it thinner. Connect 8 cm of tubing1 and 3 cm of the stretched tubing2 using glue (Figure 2A). Make a mark at 9 mm from the tip.</li>
	<li>Needle with non-sharp end that connects tubing to the syringe: Make a scratch near the sharp end of the needle (23 G) with a metal file, gently bend it back and forth a few times with pliers, and break it. Do the same cut in the middle of the needle to make a piece of metal connector (Figure 2A).</li>
	<li>Extension tube for the artery to draw blood (Tubing set1.4): Connect 10 cm of tubing1 with the metal connector and the non-sharp needle that was made in step 1.4. 
	&#8203;NOTE: All tubings and sutures are sterilized with an autoclave or ethylene oxide</li>
</ol>
2. Surgery
<ol>
	<li>Anesthetize a mouse with isoflurane (1.5-2.0%) or ketamine/xylazine (ketamine 10 mg/mL, xylazine 1 mg/mL in 0.9% sterile saline, 0.1 mL/10 g of body weight (BW) via intraperitoneal [ip] injection). Keep the mouse on the warm pad (37 &#176;C) to reduce physical stress. Wait for deep anesthesia for 5-10 min. Confirm the depth of anesthesia by pedal reflex, respiration, and heart rate, and response to stimuli by toe pinching.&#160;During surgery, surgical tape should be&#160;used to secure the nose cap to the operating table for continuous inhalation of anesthesia.&#160;Apply ophthalmic ointment to eyes to prevent dryness while under anesthesia.</li>
	<li>Fill heparinized saline (100 U/mL) in tubing1.1 and 1.3 and connect them with a 1 mL syringe with a non-sharp needle (Figure 2B). Close the end of tubing3 (see Table of Materials) by melting it with a soldering iron, which will be used as caps for tubing1.1 and tubing1.3.</li>
	<li>Shave and wipe the first (interscapular on the back) and second (neck in the front) incision area with three cycles of 10% betadine followed by a preoperative preparation with sterile 70% alcohol. Make a small vertical midline incision 5 mm cephalic to the sternum, blunt dissect the tissues, and expose the artery. Separate the vagus nerve from the artery. This reduces the negative effects of removing the vagus nerve on glucose metabolism. 
	NOTE: The step from the ventral incision to the step where the catheter is inserted into the vein and secured with silk sutures is performed under a microscope.</li>
	<li>Place two silk sutures under the artery. Stop blood flow by tying one suture tightly on the cranial side (Figure 2C-1) and the other loosely at the caudal side (Figure 2C-2), which is enough to stop blood flow but enough to open again later. Place one more suture under the artery (Figure 2C-3).</li>
	<li>Cut the artery near Figure 2C-1 with spring scissors and place tubing1.3 into the artery. Make a loose tie on both the artery and tubing (Figure 2C-3, do not tie firmly. The tubing will be inserted deeper into the artery). Open the knot on the caudal side (Figure 2C-2) to insert the tube until the 9 mm mark reaches the knot in the middle (Figure 2C-3). Securely tie all ligatures and flush with heparinized sterile saline.</li>
	<li>Expose the right jugular vein from the same incision as the right carotid artery for the jugular catheter. Isolate the cranial end and ligate it with silk suture (Figure 2D-1, Table of Materials). Place another piece of suture at the caudal end of the exposed vein (Figure 2D-2). Cut the vein near the mark Figure 2D-1&#160;with spring scissors.</li>
	<li>Insert the catheter (not too deep to prevent penetrating vessels), tie it, and confirm visually that it samples blood. Flush with heparinized sterile saline (0.2 mL) and visually confirm that no blood remains in the catheter.</li>
	<li>Place the mouse on the new sterile surgical drape to prevent infection from the first wound. Turn over the mouse, wipe with three cycles of betadine followed by a preoperative preparation with sterile alcohol, and make a small incision between the shoulder blades.</li>
	<li>Pass a needle holder under the skin from the incision on the back to the ventral side. Clamp the catheters with the needle holder, pass them under the skin, and bring them back. Clean the incision site and close the ventral incisions with a synthetic&#160;suture (diameter 0.15-0.2 mm). Clamp the venous catheter with micro serrefine at the incision site between the shoulder blades.</li>
	<li>Cut the catheter 1 cm above the clamp, flush it with heparinized saline, and close it with a cap (step 2.4). Follow the same procedure for the arterial catheter. Close the dorsal incision with a synthetic&#160;suture (diameter 0.15-0.2 mm).</li>
	<li>Place the mouse in a warm, clean cage (Figure 2E). Perform postoperative care daily.</li>
</ol>
3. Recovery
<ol>
	<li>Single-house the mouse because another mouse may bite the catheter in the group housing.
	<ol>
		<li>To reduce stress by social isolation, keep mice with environmental enrichments (e.g., shelters).&#160;Perform postoperative care daily. To relieve pain, distress, and discomfort, provide postoperative care, including analgesia (ibuprofen in drinking water (0.11 mg/mL).</li>
		<li>Observe mice for signs of infection, such as festering, lethargy, or pain at the incision site. Most healthy mice begin walking and feeding about 2 h after surgery. A hunched posture, ruffled fur, and decreased food intake may indicate pain. If these signs are observed, immediately euthanize the mice by decapitating them under deep anesthesia or carbon dioxide asphyxiation.</li>
	</ol>
	</li>
	<li>Blood will enter the catheter in the artery and form a clot. To maintain catheter lines, remove the clot daily, following steps 3-6.</li>
	<li>Fill a 1 mL syringe and 23 G needle (non-sharp end) with heparinized saline (100 U/mL). Using an induction chamber,&#160;anesthetize the mouse lightly with isoflurane (1.0%-1.5%). Then, remove the mouse from the chamber and perform clot removal under isoflurane anesthesia with a nose cap.</li>
	<li>Clamp the tubing for the artery on the back of the mouse with micro serrefine, remove the cap, and extract the blood and clots. Flush the catheter with heparinized saline using another 1 mL syringe with a 23 G needle (non-sharp end) and re-cap it. Clean the venous line like the arterial line in the case of severe clotting.</li>
	<li>Do the same procedure to clean up the catheter once a day for 3-5 days.</li>
	<li>Check the body weight of the mouse. If the body weight reduces by more than 10% from the day of the surgery, apply supportive care and try to improve the normal body condition score, and use the mouse for another experiment.&#160; 
	&#8203;NOTE: The weight loss of animals was less than 10% in the experiments in this study. Weight loss will strongly affect systemic glucose metabolism. Thus, it is recommended to wait for the body weight recovery or remove from the clamp experiment.</li>
</ol>
4. Set up the pump system (for hypoglycemic clamp)
<ol>
	<li>Prepare 1 U/mL insulin in 0.1% BSA saline, 30% glucose in saline, and heparinized saline.</li>
	<li>Measure the body weight of the mouse. Calculate the volume for 1 U/mL insulin to make the insulin infusate (10 mU/kg/min). Infuse 1.7 &#956;L/min of insulin solution into the mice in the clamp experiment. To make 300 &#956;L of insulin infusate, the required volume for 1 U/mL insulin (&#956;L) is 2.647 (&#956;L/g) x Body weight (g). A volume of 300 &#956;L of insulin infusate is enough to finish the clamp experiment on 1 mouse. Table 1 shows an example of making insulin infusate.</li>
	<li>Fill insulin infusate and 30% glucose in each Hamilton syringe, connect them with tubing 1.2, and set the syringe on the syringe pump (Figure 3A). Fill each solution in tubing 1.2. Fill saline in a 1 mL syringe and tubing set 1.4 (Figure 2A).</li>
	<li>Anesthetize the mouse with isoflurane (1.0%-1.5%) and connect tubing1.2 to the venous catheter and tubing set1.4 to the arterial catheter (Figure 3A). Use cellophane tape to hold the tubes together.</li>
	<li>Place the mouse in an empty 500 mL beaker. 
	&#8203;NOTE: For hyperglycemic clamp, use saline instead of 1 U/mL insulin.</li>
</ol>
5. Hypoglycemic clamp
<ol>
	<li>Measure blood glucose levels every 5-10 min, as shown in Figure 3B, and collect blood samples for hormone measurements at -15 min, 10 min, 20 min, 40 min, 60 min, 80 min, 100 min, and 120 min. Follow step 5.2 to measure blood glucose. 
	NOTE: It is not necessary to measure blood glucose at all time points, but it is recommended to measure it at least every 10 min. Measure blood glucose every 5 min when its level and glucose infusion rate are not steady.</li>
	<li>Clamp the top end of the tubing set1.4 and connect a new syringe, extract 50 &#956;L of blood, and place it in a 1.5 mL tube for washing.&#160;Measure blood glucose level. 
	NOTE: This is the blood diluted by the saline solution in the tube because of catheter washing after the previous sampling. A 50 &#956;L volume is enough to replace diluted blood with pure blood.</li>
	<li>Store the diluted blood (red blood cells). Wash the pooled blood (~500 &#956;L) with saline (see steps 5.11-5.12) and return to the body to prevent hypoxia.</li>
	<li>The catheter is connected to the artery with high blood pressure, so when the clamp is loosened, blood will flow out and be used to measure glucose with a handy glucose meter. Infuse 50 &#956;L of saline to keep enough amount of blood fluid.</li>
	<li>For blood sampling, extract an additional 50 &#956;L of blood and place it in a 1.5 mL tube on ice. Infuse 100 &#956;L (50 &#956;L of replacement + 50 &#956;L of sampling) of saline.</li>
	<li>After 0 min of blood glucose measurement, start the insulin syringe pump. First, infuse 30 mU/kg/min in bolus for 2 min (5.1 &#956;L /min), then 10 mU/kg/min (1.7 &#956;L /min) infusion for the rest of the duration.</li>
	<li>Measure blood glucose level and change the glucose infusion rate every 5-10 min for 120 min.</li>
	<li>Create a steady state where the glucose infusion rate does not change, and the blood glucose level is 50 mg/dL.</li>
	<li>After collecting the blood sample at t = 120 min, infuse a euthanasia agent into the jugular vein and collect tissues for RNA or protein analysis.</li>
	<li>Centrifuge blood at 1000 x g for 5 min and measure hormone concentration such as insulin or c-peptide using an ELISA kit according to the manufacturer&#39;s protocol.</li>
	<li>For washing blood, centrifuge the blood at 1000 x g for 3 min. Remove the supernatant, add 500 &#956;L of heparinized saline, and perform pipetting.</li>
	<li>Repeat the wash again and place the washed red blood cells in 1 mL syringe for the artery. The blood cells will be reinfused automatically when 50 or 100 &#956;L of saline is infused in steps 5.2 and 5.3. Monitor hematocrit carefully to prevent hypoxia. 
	&#8203;NOTE: For blood glucose measurement, a commercial handy glucose meter was used.</li>
</ol>
6. Hyperglycemic clamp
<ol>
	<li>Follow the steps as the hypoglycemic clamp technique, but without insulin infusion and with blood glucose adjusted to 250-300 mg/dL. Measure blood glucose levels every 5-10 min, as shown in Figure 3B, and collect blood samples for hormone measurements at -15 min, 10 min, 20 min, 40 min, 60 min, 80 min, 100 min, and 120 min.</li>
	<li>Connect a new syringe to the tubing set1.4, extract 50 &#956;L of blood, and place it in a 1.5 mL tube for washing.</li>
	<li>For blood sampling, extract an additional 50 &#956;L of blood and place it in a 1.5 mL tube on ice. Infuse 100 &#956;L (50 &#956;L of replacement + 50 &#956;L of sampling) of saline.</li>
	<li>After 0 min of blood glucose measurement, start the glucose syringe pump. First, infuse 30 g/kg/min in bolus for 2 min (5.1 &#956;L /min), then 10 g/kg/min (1.7 &#956;L /min) for the rest of the duration.</li>
	<li>Measure the blood glucose level and change the glucose infusion rate every 5 -10 min for 120 min. Create a steady state where the glucose infusion rate does not change, and the blood glucose level is 250-300 mg/dL, a clamped hyperglycemic state.</li>
</ol>

Surgical Installation of Hyperglycemic and Hypoglycemic Clamps in Mice

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The authors declare no competing interests.

The method described here is a simple one that can be done with pipette tips, syringes, and other items found in ordinary laboratories. Although researchers may need to purchase additional tubes and pumps, expensive equipment is not needed. Thus, this protocol of catheterization and clamp is easier to start compared to previous reports12,13,14.
The clamp technique was developed around 1970 and has bee...

Glucose is an important source of energy for cells, and a lack of glucose can lead to a variety of symptoms and complications. In the event of low glucose (hypoglycemia, generally less than 70 mg/dL in fasting blood glucose level, but should not be determined by a single value1), the most common symptoms include weakness, confusion, sweating, and headache. It can also disrupt cerebral function and increase the risk of cardiovascular events and mortality2. Conversely, hyperglycemia is a medical condition in which the plasma glucose concentration exceeds normal levels (generally &#62; 126 mg/dL in fasting blood glucose level3). This can occur in individuals with diabetes who have either a deficit in insulin production or utilization. Hyperglycemia can lead to diabetic ketoacidosis, which occurs when the body cannot use glucose for energy but instead breaks down fatty acids for fuel. The hyperglycemic hyperosmolar state also increases mortality4. Long-term hyperglycemia can cause damage to blood vessels, nerves, and organs, leading to the development of several chronic complications such as cardiovascular disease, retinopathies, and kidney diseases. Thus, the blood glucose concentration must be maintained in a tight range between 100 mg/dL and 120 mg/dL.
Blood glucose is regulated by the balance between glucose input and output in a one-compartment model (Figure 1A). Glucose input includes absorbed glucose from food and glucose production from the liver, kidneys, and small intestine. Glucose output comprises glucose uptake in tissues and glucose disposal from the kidneys. Both the amount of glucose input and output are regulated by endocrine hormones. For example, glucagon, corticosterone, and catecholamines, known as counter-regulatory hormones, are released when blood glucose levels decrease5. They stimulate the breakdown of glycogen and the synthesis of glucose, mainly from the liver; these processes are known as glycogenolysis and gluconeogenesis, respectively. Hyperglycemia increases insulin release from pancreatic &#946;-cells and stimulates glucose uptake in the muscles, adipose tissues, and heart6,7,8,9. Exercise increases insulin-independent glucose uptake10. The sympathetic nervous system increases glucose uptake in muscles and brown adipose tissue6,11. To measure the ability to regulate glucose metabolism in peripheral tissues, researchers typically use the glucose tolerance test (GTT) and the insulin tolerance test (ITT) (Figure 1B,C). In GTT, two factors must be considered: insulin release and insulin sensitivity (Figure 1B). However, the glucose concentration curve during the 120 min test is different in each mouse, which may affect different amounts of hormone release. In ITT, blood glucose is regulated by both insulin sensitivity and the release of counter-regulatory hormones. Therefore, it is difficult to determine the precise meaning of glucose metabolism, insulin release, and insulin sensitivity in GTT and ITT, in situations where blood glucose levels are not constant.
To overcome these problems, it is desirable to keep blood glucose at a constant level (or &#34;clamp&#34;). In hyperglycemic clamp, glucose is infused into the bloodstream to raise blood glucose levels to a specific level and then maintained at that level for a period of time. The amount of infused glucose is adjusted based on measurements of blood glucose levels every 5-10 min to maintain a steady state. This technique is particularly useful for understanding the parameters of insulin secretion at a clamped glucose level. Hypoglycemic clamp is a method to maintain low blood glucose levels by infusing insulin. Glucose is infused at a variable rate to maintain a specific blood glucose level. If the mouse cannot recover from hypoglycemia, more glucose should be infused.
Although there are many advantages to performing hyperglycemic and hypoglycemic clamps, the surgical and experimental procedures are considered technically difficult. Thus, few research groups have been able to do them. We aimed to describe these methods for researchers with financial and workforce constraints to start these experiments at a lower budget.

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			<td>Adhesive glue</td>
			<td>Henkel AG &#38; Co. KGaA</td>
			<td>LOCTITE 454</td>
			<td></td>
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		<tr>
			<td>ELISA kit (C-peptide)</td>
			<td>Morinaga Institute of Bilogical Science Inc</td>
			<td>M1304</td>
			<td>Mouse C-peptide ELISA Kit</td>
		</tr>
		<tr>
			<td>ELISA kit (insulin)</td>
			<td>FUJIFILM Wako Pure Chemical Corporation</td>
			<td>633-03411</td>
			<td>LBIS Mouse Insulin ELISA Kit (U-type)</td>
		</tr>
		<tr>
			<td>Handy glucose meter</td>
			<td>Nipro Co.</td>
			<td>11-777</td>
			<td>Free Style Freedom Lite</td>
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		<tr>
			<td>Insulin (100U/ml)</td>
			<td>Eli Lilly &#38; Co.</td>
			<td>428021014</td>
			<td>Humulin R (100U/ml)</td>
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			<td>Mouse</td>
			<td>Japan SLC Inc.</td>
			<td>C57BL/6NCrSlc</td>
			<td>C57BL</td>
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			<td>Suture</td>
			<td>Natsume seisakusho</td>
			<td>C-23S-560 No.2</td>
			<td>Sterilized</td>
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			<td>Syringe Pump</td>
			<td>Pump Systems Inc.</td>
			<td>NE-1000</td>
			<td></td>
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			<td>Synthetic suture</td>
			<td>V&#214;MEL</td>
			<td>HR-17</td>
			<td></td>
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			<td>Tubing1</td>
			<td>AS ONE Corporation</td>
			<td>9-869-01</td>
			<td>LABORAN(R) Silicone Tube</td>
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			<td>Tubing2</td>
			<td>Fisher Scientific</td>
			<td>427400</td>
			<td>BD Intramedic PE Tubing</td>
		</tr>
		<tr>
			<td>Tubing3</td>
			<td>IGARASHI IKA KOGYO CO., LTD.</td>
			<td>size5</td>
			<td>Polyethylene tubing size5</td>
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hyperglycemic clamp and hypoglycemic clamp in conscious mice

Diabetes mellitus (DM) is caused by insufficient insulin release from the pancreatic &#946;-cells (Type1 DM) and insulin sensitivity in muscles, liver, and adipose tissues (Type2 DM). Insulin injection treats DM patients but leads to hypoglycemia as a side effect. Cortisol and catecholamines are released to activate glucose production from the liver to recover hypoglycemia, called counter-regulatory responses (CRR). In DM research using rodent models, glucose tolerance tests and 2-deoxy-glucose injection are used to measure insulin release and CRR, respectively. However, blood glucose concentrations change persistently during experiments, causing difficulties in assessing net insulin release and CRR. This article describes a method in which blood glucose is kept at 250 mg/dL or 50 mg/dL in conscious mice to compare the release of insulin and CRR hormones, respectively.
Polyethylene tubing is implanted in the mice's carotid artery and jugular vein, and the mice are allowed to recover from the surgery. The jugular vein tubing is connected to a Hamilton syringe with a syringe pump to enable insulin or glucose infusion at a constant and variable rate. The carotid artery tubing is for blood collection. For the hyperglycemic clamp, 30% glucose is infused into the vein, and blood glucose levels are measured from the arterial blood every 5 min or 10 min. The infusion rate of 30% glucose is increased until the blood glucose level becomes 250 mg/dL. Blood is collected to measure insulin concentrations. For hypoglycemic clamp, 10 mU/kg/min insulin is infused together with 30% glucose, whose infusion rate is variable to maintain 50 mg/dL of blood glucose level. Blood is collected to measure counter-regulatory hormones when both glucose infusion and blood glucose reach a steady state. Both hyperglycemic and hypoglycemic clamps have the same surgical procedure and experimental setups. Thus, this method is useful for researchers of systemic glucose metabolism.

The hypoglycemic clamp study was performed in male C57BL/6N mice (8 weeks old, more than 25 g BW) 3 h fasted at the start of the experiment (Figure 4A,B). The initial blood glucose level was 136 mg/dL (t = -15 min). If it is less than 90 mg/dL, it may be either because the surgery did not go well, or the arterial catheter was inserted too deep, or blood clots have entered the blood flow. The mouse condition after surgery affects the energy metabolism in the mouse. Physiologi...

Watch this Scientific Journal Video about Hyperglycemic Clamp and Hypoglycemic Clamp in Conscious Mice at JoVE.com

Author Spotlight: Investigating the Blood Glucose Homeostasis in Murine Brain Using a Cost-Effective Hyperglycemic And Hypoglycemic Clamp Technique

A hyperglycemic clamp is used for measuring insulin release with a maintained higher blood glucose concentration. A hypoglycemic clamp is for measuring glucose production induced by counter-regulatory responses. Both methods use the same surgical procedure. Here, we present a clamp technique to assess systemic glucose metabolism.

Hyperglycemic Clamp and Hypoglycemic Clamp in Conscious Mice

Diabetes mellitus (DM) is caused by insufficient insulin release from the pancreatic &#946;-cells (Type1 DM) and insulin sensitivity in muscles, liver, ...

hyperglycemic-clamp-and-hypoglycemic-clamp-in-conscious-mice

Research

JoVE Journal

Medicine

1.2K Views.  Kumamoto University. A hyperglycemic clamp is used for measuring insulin release with a maintained higher blood glucose concentration. A hypoglycemic clamp is for measuring glucose production induced by counter-regulatory responses. Both methods use the same surgical procedure. Here, we present a clamp technique to assess systemic glucose metabolism.

Video: Author Spotlight: Investigating the Blood Glucose Homeostasis in Murine Brain Using a Cost-Effective Hyperglycemic And Hypoglycemic Clamp Technique

Transthoracic Echocardiography in Mice

In recent years, murine models have become the primary avenue for studying the molecular mechanisms of cardiac dysfunction resulting from changes in gene expression. Transgenic and gene targeting methods can be used to generate mice with altered cardiac size and function,1-3 and as a result, in vivo techniques are needed to evaluate their cardiac phenotype. Transthoracic echocardiography, pulse wave Doppler (PWD), and tissue Doppler imaging (TDI) can be used to provide dimensional measurements of the mouse heart and to quantify the degree of cardiac systolic and diastolic performance. Two-dimensional imaging is used to detect abnormal anatomy or movements of the left ventricle, whereas M-mode echo is used for quantification of cardiac dimensions and contractility.4,5 In addition, PWD is used to quantify localized velocity of turbulent flow,6 whereas TDI is used to measure the velocity of myocardial motion.7 Thus, transthoracic echocardiography offers a comprehensive method for the noninvasive evaluation of cardiac function in mice.

Transthoracic echocardiography offers a noninvasive method for the evaluation of cardiac function in mice. A combination of ultrasound and Doppler imaging modalities can be used to obtain dimensional measurements of the heart and intracardiac blood flow, which together provide an assessment of cardiac systolic and diastolic performance.

In recent years, murine models have become the primary avenue for studying the molecular mechanisms of cardiac dysfunction resulting from changes in gene ...

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.

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 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 ...

Carotid Artery Infusions for Pharmacokinetic and Pharmacodynamic Analysis of Taxanes in Mice

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study model. As the use of mouse models are often a vital step in preclinical drug discovery and drug development1-8, it is necessary to design a system to introduce drugs into mice in a uniform, reproducible manner. Ideally, the system should permit the collection of blood samples at regular intervals over a set time course. The ability to measure drug concentrations by mass-spectrometry, has allowed investigators to follow the changes in plasma drug levels over time in individual mice1, 9, 10. In this study, paclitaxel was introduced into transgenic mice as a continuous arterial infusion over three hours, while blood samples were simultaneously taken by retro-orbital bleeds at set time points. Carotid artery infusions are a potential alternative to jugular vein infusions, when factors such as mammary tumors or other obstructions make jugular infusions impractical. Using this technique, paclitaxel concentrations in plasma and tissue achieved similar levels as compared to jugular infusion. In this tutorial, we will demonstrate how to successfully catheterize the carotid artery by preparing an optimized catheter for the individual mouse model, then show how to insert and secure the catheter into the mouse carotid artery, thread the end of the catheter out through the back of the mouse&#8217;s neck, and hook the mouse to a pump to deliver a controlled rate of drug influx. Multiple low volume retro-orbital bleeds allow for analysis of plasma drug concentrations over time.

This method was developed with the goal of delivering a steady drug solution via the carotid artery, to assess the pharmacokinetics of novel drugs in mouse models.

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study ...

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 ...

Novel Approach for Simultaneous Recording of Renal Sympathetic Nerve Activity and Blood Pressure with Intravenous Infusion in Conscious, Unrestrained Mice.

Renal sympathetic nerves contribute significantly to both physiological and pathophysiological phenomena. Evaluating renal sympathetic nerve activity (RSNA) is of great interest in many areas of research such as chronic kidney disease, hypertension, heart failure, diabetes and obesity. Unequivocal assessment of the role of the sympathetic nervous system is thus imperative for proper interpretation of experimental results and understanding of disease processes. RSNA has been traditionally measured in anesthetized rodents, including mice. However, mice usually exhibit very low systemic blood pressure and hemodynamic instability for several hours during anesthesia and surgery. Meaningful interpretation of RSNA is confounded by this non-physiological state, given the intimate relationship between sympathetic nervous tone and cardiovascular status. To address this limitation of traditional approaches, we developed a new method for measuring RSNA in conscious, freely-moving mice. Mice were chronically instrumented with radio-telemeters for continuous monitoring of blood pressure as well as a jugular venous infusion catheter and custom-designed bipolar electrode for direct recording of RSNA. Following a 48-72 hour recovery period, survival rate was 100% and all mice behaved normally. At this time-point, RSNA was successfully recorded in 80% of mice, with viable signals acquired up to 4 and 5 days post-surgery in 70% and 50% of mice, respectively. Physiological blood pressures were recorded in all mice (116&plusmn;2 mmHg; n=10). Recorded RSNA increased with eating and grooming, as well-established in the literature. Furthermore, RSNA was validated by ganglionic blockade and modulation of blood pressure with pharmacological agents. Herein, an effective and manageable method for clear recording of RSNA in conscious, freely-moving mice is described.

Anesthetized mice exhibit non-physiological systemic blood pressure, which precludes meaningful assessment of autonomic tone given the intimate relationship between blood pressure and the autonomic nervous system. Thus, a novel method to simultaneously record renal sympathetic nerve activity and blood pressure with intravenous infusion in conscious mice is outlined.

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

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 ...

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 ...

Implantation of Combined Telemetric ECG and Blood Pressure Transmitters to Determine Spontaneous Baroreflex Sensitivity in Conscious Mice

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. The baroreflex is a key homeostatic mechanism to counteract acute, short-term changes in arterial BP and to maintain BP in a relatively narrow physiological range. BP is sensed by baroreceptors located in the aortic arch and carotid sinus. When BP changes, signals are transmitted to the central nervous system and are then communicated to the parasympathetic and sympathetic branches of the autonomic nervous system to adjust HR. A rise in BP causes a reflex decrease in HR, a drop in BP causes a reflex increase in HR.
Baroreflex sensitivity (BRS) is the quantitative relationship between changes in arterial BP and corresponding changes in HR. Cardiovascular diseases are often associated with impaired baroreflex function. In various studies reduced BRS has been reported in e.g., heart failure, myocardial infarction, or coronary artery disease. 
Determination of BRS requires information from both BP and HR, which can be recorded simultaneously using telemetric devices. The surgical procedure is described beginning with the insertion of the pressure sensor into the left carotid artery and positioning of its tip in the aortic arch to monitor arterial pressure followed by the subcutaneous placement of the transmitter and ECG electrodes. We also describe postoperative intensive care and analgesic management. After a two-week period of post-surgery recovery long-term ECG and BP recordings are performed in conscious and unrestrained mice. Finally, we include examples of high-quality recordings and the analysis of spontaneous baroreceptor sensitivity using the sequence method.

The baroreflex is a heart-rate regulation mechanism by the autonomic nervous system in response to blood-pressure changes. We describe a surgical technique to implant telemetry transmitters for continuous and simultaneous measurement of electrocardiogram and blood pressure in mice. This can determine spontaneous baroreflex sensitivity, an important prognostic marker for cardiovascular disease.

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. ...

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 ...