We thank the staff from the Alfred Medical Research and Education Precinct Animal Services (AMREP AS) team for their assistance and care of the mice used in this study and for the support of the Operational Infrastructure Support scheme of the Victorian State Government.

All experiments described were approved by the Alfred Medical Research Education Precinct Animal Ethics Committee (AMREP AEC) and mice were provided humane care in line with the National Health and Medical Research Council (NHMRC) of Australia Guidelines on Animal experimentation. Animals were administered their prescribed diet and water ad libitum and housed in a temperature-controlled environment (~21 - 22 &#176;C) with a 12 h light and 12 h-dark cycle. Seven week old male mice (on a C57Bl/6J background) were fed either regular normal chow diet (energy content 14.3 MJ/kg, consisting of 76% of kJ from carbohydrate, 5% fat, 19% protein; see Table of Materials) or for the high fat-feeding group, a high fat diet (HFD) (energy content 19 MJ/kg, consisting of 36% of kJ from carbohydrate, 43% fat, 21% protein, Specialty Feeds) for 3 weeks. Body weight and body composition measurements using an EchoMRI machine were made weekly while the metabolic monitoring analysis took place in a CLAMS after 3 weeks of the diet.
1. Body Composition Analyzer Procedure
Note: To function optimally, the EchoMRI 4-in-1 used in this protocol should be contained within a room where the air temperature is stable and does not fluctuate. Ideally this should be constantly monitored. Moving of the machine and interruptions to power should also be avoided if possible. If the power supply has been interrupted and the system has to be restarted, allow at least 2 - 3 h for the machine to warm up before using it again. Before starting, ensure that you are wearing correct personal protective equipment.
<ol>
	<li>Prior to scanning mice, perform a system test on the body composition analyzer machine. This involves using a calibration standard (referred to as a canola oil system test sample (COSTS)) to test the precision of the instrument and to ensure there has been no drift in its accuracy.
	<ol>
		<li>Open the system software, then click the&#160;System Test toolbar button or pressing &#34;Alt + Y&#34; simultaneously.</li>
		<li>Before the system test is carried out by the computer, wait for a reminder to verify that the correct COSTS (in this case the mouse-specific COSTS) has been placed inside the gantry of the system ( Figure 1). Once confirmed that this is indeed the case, accept to proceed with the test, which will take few minutes to complete.</li>
	</ol>
	</li>
	<li>Once the system test has been passed, continue forward with scanning.
	<ol>
		<li>If the system test fails, repeat the system test.</li>
		<li>If the machine continues to be out of range (indicating a deviation has occurred), calibration may be necessary to rectify the situation. Complete this by following the prompts or as described in the user manual provided at time of purchase. If the problem persists, check the manual3 or report the issue to the manufacturer&#39;s support team and seek further instruction.</li>
	</ol>
	</li>
	<li>Place the mice in a small animal specimen holder (long cylinder) to keep them contained while in the machine. To do this, place the holder horizontally, pick up the mouse and insert it into the opening of the cylinder head first. Slowly and carefully bring the holder to the vertical position so that the mouse is at the bottom of the cylinder and ready for analysis.</li>
	<li>Once within the holder, insert a delimiter to limit the movement of the mouse during the measurement period. In some circumstances, with extremely active mice, it may be necessary to hold the delimiter in place with your fingertip. 
	NOTE: Familiarize the mice with placement in the specimen holders prior to their initial analysis to reduce stress. The use of a red colored animal specimen holder can also reduce the potential stress response, as the mice feel that they are in the dark.</li>
	<li>Within the software, select a folder (folder toolbar) to save the data to and create a file name.</li>
	<li>If necessary, reduce the amount of random noise in the fat and lean measurements by increasing the number of primary accumulations of the scan. Once the software is initiated, the primary accumulations is set to a recommended default value for general everyday use; unless there is a specific reason to change these parameters, the default settings will give the necessary level of precision to users.</li>
	<li>If not interested in obtaining data for free water and total water, turn off the water stage by selecting the tab to say no. Doing so will reduce the scan duration significantly and improve throughput.</li>
	<li>Initiate the scan by selecting &#34;start scan&#34; or by pressing F5 on the keyboard. Enter all relevant data about the animal (e.g., animal ID, body mass, etc.) and press &#34;ok&#34; or F5 to commence the scan, which will take approximately 1 min.</li>
	<li>After data have been obtained, remove the animal holder containing the mouse from the machine and place the animal back in its cage. Once all animals have been scanned, export the data for further analysis and collation.</li>
	<li>Before and after use, thoroughly clean the animal holders as per the instructions of the manufacturer. As these holders are constructed from acrylic plastic, isopropyl alcohol and ethyl alcohol should be avoided as they can cause cracking of the holders and/or rapid deterioration of the holder, thereby increasing the likelihood of breakage. Instead, either use warm dishwashing water solution, or, if further disinfectant is required, use F10 (at a 1:125 dilution) or other disinfectant or cleaning sprays (see Table of Materials) and then wipe off.</li>
</ol>
2. Metabolic Animal Monitoring System Procedure
NOTE: The system requires ~2 h to warm up and stabilize. If the machine has been turned off, it must be switched on to allow the Zirconia cell to be heated to 725 &#176;C. Also we generally place mice in the body composition analyzer a day prior to entering the animal monitoring system to avoid any issues with restraint stress.
<ol>
	<li>Ensure the computer attached to the animal monitoring system is turned on and open the control program. Select the &#34;Oxymax Utility&#34; option from the tool menu to initiate the pumps.</li>
	<li>Fill water bottles with appropriate water, weigh and inspect the health of the mice, and organize food. If measuring food intake in the system, consider powdering the food. Fill the food hoppers by depressing the spring-loaded platform and tip food into the hopper. Ensure that the food hopper and water bottle are completely full to ensure that there is enough food and water to last the allotted experimental time.</li>
	<li>Check the status of the drierite/desiccant; if using a color indicator, it should be blue and therefore dry, but if it is pink/purple, it has had significant moisture absorbance and should be replaced or topped up.</li>
	<li>Check the condition of the ammonia trap and soda lime and replace if required. If the ammonia trap is connected two at a time, when the second trap displays signs of a color change, replace the first one. An increase in the CO2 offset can also signify the need to replace the soda lime. 
	NOTE: Desiccant can be dried in an oven and reused, however we follow the recommendations from the manufacturer of the system to use fresh each time.</li>
	<li>Assemble the chambers. To do this, place the food hopper on the balance, then place the chamber on top with the perforated platform that becomes the floor of the chamber inserted. Carefully place the mouse in the chamber and attach the lid of the system with the front and back clips and secure before positioning the water bottle and fastening. As a precaution, re-check all chamber lids, mice, and water (Figure 2A-D). 
	NOTE: Depending on the size of the mice being examined, it may be necessary to adjust the height of the spaces above the food hopper so that the mice have access to the food but not enough space that they can sleep directly on top of the feeder.</li>
	<li>As it is recommended that the gas sensors be calibrated before each experiment, calibrate the system.
	<ol>
		<li>Use a gas of known composition (0.5% CO2, 20.5% O2, balance nitrogen). Connect the calibration gas tank to the system via a regulator and hose. Turn on and ensure the tank output pressure is reading 5 - 10 psi. 
		Note: Some systems will have a second tank, hose and regulator for use of pure Nitrogen as an &#34;offset&#34; gas. The system we operate instead utilizes soda lime to generate CO2 free air.</li>
		<li>Follow procedures to calibrate both the O2 and CO2 sensors. Select &#34;calibration&#34; from the tools menu and sequentially calibrate both the O2 and CO2. Before calibrating ensure that 1) sample and reference flows are 0.400LPM, 2) the Zirconia O2 sensor temperature is 725 &#176;C (&#177; 1 &#176;C), 3) the sample and reference drier and air pumps are on, and 4) the calibration gas is attached and turned on.</li>
		<li>If necessary, when calibrating the O2 sensor, slightly adjust the offset control on the front of the zirconia oxygen sensor to achieve an O2 ratio value of 1.0000 (&#177; 0.0002). This is to ensure it is within acceptable limits (highlighted in green font in the software display on the computer screen).</li>
		<li>After successful O2 and CO2 sensor calibration, turn off the calibration gas cylinder and disconnect the hose from the regulator. After calibration, O2 for reference air (atmospheric) should read 20.92 (&#177; 00.02). If calibration is out of tolerance, repeat, and refer to trouble shooting guides from the manufacturer. Failing this, contact the manufacturer for further instructions.</li>
	</ol>
	</li>
	<li>Proceed with experimental set-up. Select &#34;experimental file open&#34; from the experiment menu. Select the appropriate template (e.g., mouse). Under &#34;setup&#34; in the experiment menu define the parameters of the experiment that should be recorded (e.g., mouse ID, weight, group, etc.) De-select any chambers not in use and select the location for the experiment to be saved.</li>
	<li>Ensure the scales have been tared if measuring food intake and start the capturing of data by selecting &#34;run&#34; in the experiment menu. Data is captured for different lengths of time depending on the phenotype, institutional guidelines on animal isolation, and system usage. 
	NOTE: In our hands, the experiment is routinely run for 48 h, with the first 24 h used as acclimatisation to the new environment and the second 24 h used for data analysis. The data collection period is based on how long the investigator wishes to keep their mice singly housed and dependent on animal ethics approval. Alternatively, if provisions exist, mice may be acclimatised in the chambers prior to being placed into the system and connected. Each chamber is measured approximately once every 13 min when a 12 chamber system is in use.</li>
	<li>Regularly check and monitor the results that are obtained while the mice are in the system to ensure animal welfare and that appropriate data is being collected. Any issue may be able to be identified at this stage and rectified. Check on each mouse every morning and evening when they are in the system.</li>
	<li>Check the metabolic tab at the top of the data file page for the data collected in real time for each mouse in regards to the oxygen consumption, RER, and energy expenditure. Meanwhile, beam breaks and food consumption data can be located in the activity and feeding tabs, respectively. Check that the &#34;O2 In&#34; is reading around 20.90 - 20.94, the &#34;CO2 In&#34; is around 0.040 - 0.050, the RER is between 0.7 and 1, and the flow rate is constant at 0.5 - 0.6 L/min.</li>
	<li>At regular intervals, check that the mice have access to food and water and that they are consuming each. Ensure they are not demonstrating any signs of distress such as digging at the perforated flooring. Also, monitor the results which are displayed.</li>
	<li>At the completion of the allocated experimental time, select &#34;stop&#34; from the experiment menu and export the results (as CSV files, File&#62;Export&#62;Generate Subject CSV&#39;s) for analysis.</li>
	<li>Inspect the health of the mice, weigh them and then return to their home cages.
	<ol>
		<li>Mice may be hostile towards each other after separation, so monitor once they are housed together again.</li>
		<li>Disassemble the cages, remove excess food from hoppers, and tip out any feces, urine, and food from the cages. Submerge bottles and sippers in diluted T-bac solution, soak, and clean the other components in diluted bleach solution. Rinse with clean water and leave to air dry.</li>
	</ol>
	</li>
	<li>Calculate metabolic parameters with the software. The software utilizes a number of equations to provide the final data output4. 
	For calculation of oxygen consumption and carbon dioxide production: Oxygen consumption: VO2 (LPM)= ViO2i - VoO2o; Carbon dioxide production: VCO2 (LPM)=VoCO2o-ViCO2i 
	Where: Vi = the input ventilation rate (LPM), Vo = the output ventilation rate (LPM), O2i = the O2 concentration at input, O2o = the O2 concentration at output, CO2i = the CO2 concentration at input, CO2o = the CO2 concentration at output. 
	For calculation of RER: RER = VCO2 / VO2. Note that protein oxidation was not measured and therefore the RER was not adjusted for this. 
	For calculation of energy expenditure: Energy Expenditure: CV = 3.815 + 1.232* RER 
	Heat (Kcal/h)) = CV * VO2. Where: CV is the calorific value (the relationship between heat and the volume of oxygen consumption). This is derived from &#34;The Elements of the Science of Nutrition&#34; referred to as the Lusk Table, comprised by Graham Lusk.</li>
</ol>

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The authors have nothing to disclose.

Critical steps 
The protocols described herein provide an example of ways in which to measure body composition and various metabolic parameters in mice using a body composition analyzer and a metabolic animal monitoring system. For both techniques, it is critically important to ensure that the machines are working optimally, and to do this, it is imperative that the researcher performs a system test for the body composition analyzer and calibrates to a known gas composition fo...

Metabolism underpins many aspects of normal cellular, organ, and whole-body physiology. Consequently, in the setting of various pathologies, alterations to metabolism may directly contribute to the underlying condition or may be adversely impacted as a side-effect of the pathology. Traditionally, metabolic research and studies into energy balance have been concentrated on the field of obesity and related conditions such as insulin resistance, pre-diabetes, glucose intolerance, cardiovascular disease, and diabetes. This research is warranted given the escalating prevalence of such conditions worldwide and the individual, societal, and economic costs these conditions inflict. As such, the development of prevention strategies and new therapeutics to target obesity is a continuing goal in research laboratories around the world and preclinical mouse models are heavily relied upon for these studies.
While weighing mice provides a reliable assessment of weight gain or loss, it does not provide a breakdown of the different components that make up whole-body composition (fat mass, lean mass, free water as well as other components such as fur and claws). The weighing of fat pads at the completion of studies once the mouse is deceased provides an accurate measure of different fat depots but can only provide data for a single time point. As a consequence, it is often necessary to enroll multiple cohorts to investigate the development of obesity over time, significantly increasing animal numbers, time, and costs. The use of dual-energy X-ray absorptiometry (DEXA) provides an approach to assess body fat and lean tissue contents and allows the researcher to obtain data in a longitudinal fashion. However, the procedure requires mice to be anesthetized1, and repeated bouts of anesthesia may impact the accumulation of adipose tissue or impact other aspects of metabolic regulation. EchoMRI utilizes nuclear magnetic resonance relaxometry to measure fat and lean mass, free water, and total water content. This is achievable due to the creation of contrast between the different tissue components, with differences in the duration, amplitude and spatial distribution of generated radio frequencies allowing the delineation and quantification of each tissue type. This technique is advantageous as it is non-invasive, quick, simple, requires no anesthesia or radiation, and, importantly, has been positively validated against chemical analysis2.
A key consideration of obesity and related research is the energy balance equation. While fat accumulation is more complicated than purely energy in (food intake) versus energy out (energy expenditure), they are vital factors to be able to measure. Daily energy expenditure is the total of four different components: (1) basal energy expenditure (resting metabolic rate); (2) the energy expenditure due to the thermic effect of food consumption; (3) the energy required for thermoregulation; and (4) the energy spent on physical activity. As energy expenditure generates heat, measuring heat production by an animal (known as direct calorimetry) can be used to assess energy expenditure. Alternatively, measurement of inspired and expired concentrations of O2 and CO2, allowing for determination of whole-body O2 consumption and CO2 production, can be utilized as a way to indirectly measure (indirect calorimetry) heat production and consequently calculate energy expenditure. An increase in food intake or a decrease in energy expenditure will predispose mice to weight gain and observations of changes in these parameters can provide useful information of likely mechanisms of action in particular models of obesity. A related metabolic parameter of interest is the respiratory exchange ratio (RER), an indicator of the proportion of substrate/fuel (i.e., carbohydrate or fat) that is undergoing metabolism and being utilized to produce energy. Consequently, measurement of food intake (energy consumed) combined with physical activity levels, O2 consumption, RER, and energy expenditure can provide a broad understanding of an organism&#39;s metabolic profile. One method to gather such data is to use a comprehensive laboratory animal monitoring system (CLAMS), which is based on the indirect calorimetry method to measure energy expenditure and has the added capabilities of determining physical activity levels (beam breaks) and food intake via scales incorporated into the measurement chamber.
In this protocol we provide a straight-forward description of the use of a body composition analyzer to assess body composition in mice and a metabolic animal monitoring system to measure aspects of metabolism. Considerations and limitations for these techniques will be discussed as well as suggested methods of analysis, interpretation, and data representation.

<table border="0" cellpadding="0" cellspacing="0" width="1053">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:251px;">4 in 1 system</td>
			<td style="width:157px;">EchoMRI</td>
			<td style="width:275px;">4 in 1 system</td>
			<td style="width:371px;">Whole body composition analyser</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Canola oil test sample (COSTS)</td>
			<td>EchoMRI</td>
			<td style="width:275px;">Mouse-specific (contact company for cat number)</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Animal specimen holder&#160;</td>
			<td>EchoMRI</td>
			<td style="width:275px;">103-E56100R</td>
			<td style="width:371px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:251px;">Delimiter&#160;</td>
			<td>EchoMRI</td>
			<td style="width:275px;">600-E56100D</td>
			<td style="width:371px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">12 chamber system</td>
			<td>Columbus Instruments</td>
			<td>Custom built</td>
			<td>Metabolic Caging System; includes control program</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Drierite</td>
			<td>Fisher Scientific</td>
			<td>238988</td>
			<td>CLAMS consumable</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">Calibration gas tank</td>
			<td>Air Liquide</td>
			<td>Mixed to order</td>
			<td>Gas calibration (0.5% CO2, 20.5% O2, balance nitrogen).&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Normal chow diet</td>
			<td>Specialty Feeds</td>
			<td>Irradiated mouse and rat diet</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">High fat diet</td>
			<td>Specialty Feeds</td>
			<td>SF04-001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Balance</td>
			<td>Mettler Toledo</td>
			<td>PL202-S</td>
			<td>Balance for weighing mice</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">TexQ Disinfectant spray</td>
			<td>TexWipe</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hydrogen Peroxide cleaning solution</td>
			<td>TexWipe</td>
			<td>TX684</td>
			<td></td>
		</tr>
	</tbody>
</table>

body composition and metabolic caging analysis in high fat fed mice

Alterations to body composition (fat or lean mass), metabolic parameters such as whole-body oxygen consumption, energy expenditure, and substrate utilization, and behaviors such as food intake and physical activity can provide important information regarding the underlying mechanisms of disease. Given the importance of body composition and metabolism to the development of obesity and its subsequent sequelae, it is necessary to make accurate measures of these parameters in the pre-clinical research setting. Advances in technology over the past few decades have made it possible to derive these measures in rodent models in a non-invasive and longitudinal fashion. Consequently, these metabolic measures have proven useful when assessing the response of genetic manipulations (for example knockout or transgenic mice, viral knock-down or overexpression of genes), experimental drug/compound screening and dietary, behavioral or physical activity interventions. Herein, we describe the protocols used to measure body composition and metabolic parameters using an animal monitoring system in chow-fed and high fat diet-fed mice.

The results seen in Figure 3 display a typical change in body composition parameters upon high fat feeding, as measured via EchoMRI. At baseline there was no difference in any parameter measured (Figure 3A-F). However, after just 1 week of high fat feeding, there was a significant increase in body weight, fat mass, and fat mass percentage in the HFD group (Figure 3A,B</strong...

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Body Composition and Metabolic Caging Analysis in High Fat Fed Mice

This protocol describes the use of a body composition analyzer and metabolic animal monitoring system to characterize body composition and metabolic parameters in mice. An obesity model induced by high-fat feeding is used as an example for the application of these techniques.

Alterations to body composition (fat or lean mass), metabolic parameters such as whole-body oxygen consumption, energy expenditure, and substrate ...

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15.3K Views.  Baker Heart and Diabetes Institute. This protocol describes the use of a body composition analyzer and metabolic animal monitoring system to characterize body composition and metabolic parameters in mice. An obesity model induced by high-fat feeding is used as an example for the application of these techniques. 

Video: Body Composition and Metabolic Caging Analysis in High Fat Fed Mice

Dissection of Human Vitreous Body Elements for Proteomic Analysis

The vitreous is an optically clear, collagenous extracellular matrix that fills the inside of the eye and overlies the retina. 1,2 Abnormal interactions between vitreous substructures and the retina underlie several vitreoretinal diseases, including retinal tear and detachment, macular pucker, macular hole, age-related macular degeneration, vitreomacular traction, proliferative vitreoretinopathy, proliferative diabetic retinopathy, and inherited vitreoretinopathies. 1,2 The molecular composition of the vitreous substructures is not known. Since the vitreous body is transparent with limited surgical access, it has been difficult to study its substructures at the molecular level. We developed a method to separate and preserve these tissues for proteomic and biochemical analysis. The dissection technique in this experimental video shows how to isolate vitreous base, anterior hyaloid, vitreous core, and vitreous cortex from postmortem human eyes. One-dimensional SDS-PAGE analyses of each vitreous component showed that our dissection technique resulted in four unique protein profiles corresponding to each substructure of the human vitreous body. Identification of differentially compartmentalized proteins will reveal candidate molecules underlying various vitreoretinal diseases.

This video shows an effective technique for differentiating and dissecting the various semi-transparent structures of the human vitreous body in post mortem eyes.

The vitreous is an optically clear, collagenous extracellular matrix that fills the inside of the eye and overlies the retina. 1,2 Abnormal interactions ...

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 ...

Protocol for Long Duration Whole Body Hyperthermia in Mice

Hyperthermia is a general term used to define the increase in core body temperature above normal. It is often used to describe the increased core body temperature that is observed during fever. The use of hyperthermia as an adjuvant has emerged as a promising procedure for tumor regression in the field of cancer biology. For this purpose, the most important requirement is to have reliable and uniform heating protocols. We have developed a protocol for hyperthermia (whole body) in mice. In this protocol, animals are exposed to cycles of hyperthermia for 90 min followed by a rest period of 15 min. During this period mice have easy access to food and water. High body temperature spikes in the mice during first few hyperthermia exposure cycles are prevented by immobilizing the animal. Additionally, normal saline is administered in first few cycles to minimize the effects of dehydration. This protocol can simulate fever like conditions in mice up to 12-24 hr. We have used 8-12 weeks old BALB/Cj female mice to demonstrate the protocol.

This paper describes a protocol for whole body hyperthermia in mice that can stimulate fever like conditions up to 12-24 hr.

Hyperthermia is a general term used to define the increase in core body temperature above normal. It is often used to describe the increased core body ...

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 ...

Dissection of the Mouse Pancreas for Histological Analysis and Metabolic Profiling

We have been investigating the pancreas specific transcription factor, 1a cre-recombinase; lox-stop-lox- Kristen rat sarcoma, glycine to aspartic acid at the 12 codon (Ptf1acre/+;LSL-KrasG12D/+) mouse strain as a model of human pancreatic cancer. The goal of our current studies is to identify novel metabolic biomarkers of pancreatic cancer progression. We have performed metabolic profiling of urine, feces, blood, and pancreas tissue extracts, as well as histological analyses of the pancreas to stage the cancer progression. The mouse pancreas is not a well-defined solid organ like in humans, but rather is a diffusely distributed soft tissue that is not easily identified by individuals unfamiliar with mouse internal anatomy or by individuals that have little or no experience performing mouse organ dissections. The purpose of this article is to provide a detailed step-wise visual demonstration to guide novices in the removal of the mouse pancreas by dissection. This article should be especially valuable to students and investigators new to research that requires harvesting of the mouse pancreas by dissection for metabolic profiling or histological analyses.

This video article provides a detailed demonstration of the procedures required to successfully remove the pancreas from a mouse by dissection for histological analysis and metabolic profiling.

We have been investigating the pancreas specific transcription factor, 1a cre-recombinase; lox-stop-lox- Kristen rat sarcoma, glycine to aspartic acid at ...

Semi-automated Analysis of Mouse Skeletal Muscle Morphology and Fiber-type Composition

For years, distinctions between skeletal muscle fiber types were best visualized by myosin-ATPase staining. More recently, immunohistochemical staining of myosin heavy chain (MyHC) isoforms has emerged as a finer discriminator of fiber-type. Type I, type IIA, type IIX and type IIB fibers can now be identified with precision based on their MyHC profile; however, manual analysis of these data can be slow and down-right tedious. In this regard, rapid, accurate assessment of fiber-type composition and morphology is a very desirable tool. Here, we present a protocol for state-of-the-art immunohistochemical staining of MyHCs in frozen sections obtained from mouse hindlimb muscle in concert with a novel semi-automated algorithm that accelerates analysis of fiber-type and fiber morphology. As expected, the soleus muscle displayed staining for type I and type IIA fibers, but not for type IIX or type IIB fibers. On the other hand, the tibialis anterior muscle was composed predominantly of type IIX and type IIB fibers, a small fraction of type IIA fibers and little or no type I fibers. Several image transformations were used to generate probability maps for the purpose of measuring different aspects of fiber morphology (i.e., cross-sectional area (CSA), maximal and minimal Feret diameter). The values obtained for these parameters were then compared with manually-obtained values. No significant differences were observed between either mode of analysis with regards to CSA, maximal or minimal Feret diameter (all p &#62; 0.05), indicating the accuracy of our method. Thus, our immunostaining analysis protocol may be applied to the investigation of effects on muscle composition in many models of aging and myopathy.

Immunohistochemical staining of myosin heavy chain isoforms has emerged as the state-of-the-art discriminator of skeletal muscle fiber-type (i.e., type I, type IIA, type IIX, type IIB). Here, we present a staining protocol along with a novel semi-automated algorithm that facilitates rapid assessment of fiber-type and fiber morphology.

For years, distinctions between skeletal muscle fiber types were best visualized by myosin-ATPase staining. More recently, immunohistochemical staining of ...

Study of In Vivo Glucose Metabolism in High-fat Diet-fed Mice Using Oral Glucose Tolerance Test (OGTT) and Insulin Tolerance Test (ITT)

Obesity represents the most important single risk factor in the pathogenesis of type 2 diabetes, a disease which is characterized by a resistance to insulin-stimulated glucose uptake and a gross decompensation of systemic glucose metabolism. Despite considerable progress in the understanding of glucose metabolism, the molecular mechanisms of its regulation in health and disease remain under-investigated, while novel approaches to prevent and treat diabetes are urgently needed. Diet derived glucose stimulates the pancreatic secretion of insulin, which serves as the principal regulator of cellular anabolic processes during the fed-state and thus balances blood glucose levels to maintain systemic energy status. Chronic overfeeding triggers meta-inflammation, which leads to alterations in peripheral insulin receptor-associated signaling and thus reduces the sensitivity to insulin-mediated glucose disposal. These events ultimately result in elevated fasting glucose and insulin levels as well as a reduction in glucose tolerance, which in turn serve as important indicators of insulin resistance. Here, we present a protocol for the generation and metabolic characterization of high-fat diet (HFD)-fed mice as a frequently used model of diet-induced insulin resistance. We illustrate in detail the oral glucose tolerance test (OGTT), which monitors the peripheral disposal of an orally administered glucose load and insulin secretion over time. Additionally, we present a protocol for the insulin tolerance test (ITT) to monitor whole-body insulin action. Together, these methods and their downstream applications represent powerful tools to characterize the general metabolic phenotype of mice as well as to specifically assess alterations in glucose metabolism. They may be especially useful in the broad research field of insulin resistance, diabetes and obesity to provide a better understanding of pathogenesis as well as to test the effects of therapeutic interventions.

Study of In Vivo Glucose Metabolism in High-fat Diet-fed Mice Using Oral Glucose Tolerance Test OGTT and Insulin Tolerance Test ITT

The current article describes the generation and metabolic characterization of high-fat diet-fed mice as a model of diet-induced insulin resistance and obesity. It further features detailed protocols to perform the oral glucose tolerance test and the insulin tolerance test, monitoring whole-body alterations of glucose metabolism in vivo.

Obesity represents the most important single risk factor in the pathogenesis of type 2 diabetes, a disease which is characterized by a resistance to ...

Incorporation of a Survivable Liver Biopsy Procedure in Mice to Assess Non-alcoholic Steatohepatitis (NASH) Resolution

Clinical trials assessing therapies for the treatment of non-alcoholic steatohepatitis (NASH) involve a baseline and end of study liver biopsy, and assessment of improvement in disease endpoints, often reflected as a percent of each treatment arm that improved, worsened or remained unchanged. Traditional preclinical rodent studies for putative NASH therapies are often limited by not knowing the level of liver disease/NASH present at the start of therapeutic intervention, instead of randomizing treatment groups on easily measurable endpoints such as body weight, metabolic status or similar. Here, we describe a liver biopsy technique in a diet-induced NASH mouse model, for the assessment of baseline liver disease in order to exclude mice that do not exhibit fibrosis and to equally distribute animals with similar fibrosis between treatment groups. These levels can then be compared to the terminal, post-intervention levels for a truer understanding of in vivo pharmacological effects and thus more accurately reflect clinical trial design strategies. The mouse is properly anesthetized and prepared for the surgery using sterile conditions. A small incision is made in the upper abdomen and the left lateral lobe of the liver is exposed. A wedge of the liver is surgically removed, and a similar-sized piece of absorbable gelatin is put in its place to stop any bleeding. The mouse is surgically sutured and stapled closed and will recover back to normal within 1 day. The entire process takes 5-10 min per mouse. Here we exemplify the utility of this procedure by leveraging the pre-study biopsy to assess the impact of the glucagon-like peptide-1 (GLP-1) receptor agonist liraglutide on NASH endpoints in mice.

Incorporation of a Survivable Liver Biopsy Procedure in Mice to Assess Non-alcoholic Steatohepatitis NASH Resolution

In clinical settings, a liver biopsy is used to assess stages of non-alcoholic steatohepatitis (steatosis, inflammation, hepatocyte ballooning, and fibrosis). Here, we illustrate a survivable liver biopsy in mice which can be used for histological analyses to enable assessment of therapeutic agents in a manner more aligned with clinical trials.

Clinical trials assessing therapies for the treatment of non-alcoholic steatohepatitis (NASH) involve a baseline and end of study liver biopsy, and ...

Quantitative Analysis of Cellular Composition in Advanced Atherosclerotic Lesions of Smooth Muscle Cell Lineage-Tracing Mice

Atherosclerosis remains the leading cause of death worldwide and, despite countless preclinical studies describing promising therapeutic targets, novel interventions have remained elusive. This is likely due, in part, to a reliance on preclinical prevention models investigating the effects of genetic manipulations or pharmacological treatments on atherosclerosis development rather than the established disease. Also, results of these studies are often confounding because of the use of superficial lesion analyses and a lack of characterization of lesion cell populations. To help overcome these translational hurdles, we propose an increased reliance on intervention models that employ investigation of changes in cellular composition at a single cell level by immunofluorescent staining and confocal microscopy. To this end, we describe a protocol for testing a putative therapeutic agent in a murine intervention model including a systematic approach for animal dissection, embedding, sectioning, staining, and quantification of brachiocephalic artery lesions. In addition, due to the phenotypic diversity of cells within late-stage atherosclerotic lesions, we describe the importance of using cell-specific, inducible lineage tracing mouse systems and how this can be leveraged for unbiased characterization of atherosclerotic lesion cell populations. Together, these strategies may assist vascular biologists to more accurately model therapeutic interventions and analyze atherosclerotic disease and will hopefully translate into a higher rate of success in clinical trials.

We propose a standardized protocol to characterize the cellular composition of late-stage murine atherosclerotic lesions including systematic methods of animal dissection, tissue embedding, sectioning, staining, and analysis of brachiocephalic arteries from atheroprone smooth muscle cell lineage tracing mice.

Atherosclerosis remains the leading cause of death worldwide and, despite countless preclinical studies describing promising therapeutic targets, novel ...

Vinyl Chloride and High-Fat Diet as a Model of Environment and Obesity Interaction

Vinyl chloride (VC), an abundant environmental contaminant, causes steatohepatitis at high levels, but is considered safe at lower levels. Although several studies have investigated the role of VC as a direct hepatotoxicant, the concept that VC modifies sensitivity of the liver to other factors, such as nonalcoholic fatty liver disease (NAFLD) caused by high-fat diet (HFD) is novel. This protocol describes an exposure paradigm to evaluate the effects of chronic, low-level exposure to VC. Mice are acclimated to low-fat or high-fat diet one week prior to the beginning of the inhalation exposure and remain on these diets throughout the experiment. Mice are exposed to VC (sub-OSHA level: &#60;1 ppm) or room air in inhalation chambers for 6 hours/day, 5 days/week, for up to 12 weeks. Animals are monitored weekly for body weight gain and food consumption. This model of VC exposure causes no overt liver injury with VC inhalation alone. However, the combination of VC and HFD significantly enhances liver disease. A technical advantage of this co-exposure model is the whole-body exposure, without restraint. Moreover, the conditions more closely resemble a very common human situation of a combined exposure to VC with underlying nonalcoholic fatty liver disease and therefore support the novel hypothesis that VC is an environmental risk factor for the development of liver damage as a complication of obesity (i.e., NAFLD). This work challenges the paradigm that the current exposure limits of VC (occupational and environmental) are safe. The use of this model can shed new light and concern on the risks of VC exposure. This model of toxicant-induced liver injury can be used for other volatile organic compounds and to study other interactions that may impact the liver and other organ systems.

The goal of this protocol was to develop a murine model of low-level toxicant exposure that does not cause overt liver injury but rather exacerbates pre-existing liver damage. This paradigm better recapitulates human exposure and the subtle changes that occur upon exposure to toxicant concentrations that are considered safe.

Vinyl chloride (VC), an abundant environmental contaminant, causes steatohepatitis at high levels, but is considered safe at lower levels. Although ...