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><tbody><tr><td>4 in 1 system</td><td>EchoMRI</td><td>4 in 1 system</td><td>Whole body composition analyser</td></tr><tr><td>Canola oil test sample (COSTS)</td><td>EchoMRI</td><td>Mouse-specific (contact company for cat number)</td><td></td></tr><tr><td>Animal specimen holder&amp;nbsp;</td><td>EchoMRI</td><td>103-E56100R</td><td></td></tr><tr><td>Delimiter&amp;nbsp;</td><td>EchoMRI</td><td>600-E56100D</td><td></td></tr><tr><td>12 chamber system</td><td>Columbus Instruments</td><td>Custom built</td><td>Metabolic Caging System; includes control program</td></tr><tr><td>Drierite</td><td>Fisher Scientific</td><td>238988</td><td>CLAMS consumable</td></tr><tr><td>Calibration gas tank</td><td>Air Liquide</td><td>Mixed to order</td><td>Gas calibration (0.5% CO&lt;sub&gt;2&lt;/sub&gt;, 20.5% O&lt;sub&gt;2&lt;/sub&gt;, balance nitrogen).&amp;nbsp;</td></tr><tr><td>Normal chow diet</td><td>Specialty Feeds</td><td>Irradiated mouse and rat diet</td><td></td></tr><tr><td>High fat diet</td><td>Specialty Feeds</td><td>SF04-001</td><td></td></tr><tr><td>Balance</td><td>Mettler Toledo</td><td>PL202-S</td><td>Balance for weighing mice</td></tr><tr><td>TexQ Disinfectant spray</td><td>TexWipe</td><td></td><td></td></tr><tr><td>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...

Watch this Scientific Journal Video about Body Composition and Metabolic Caging Analysis in High Fat Fed Mice at JoVE.com

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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15.4K Views.  Baker Heart and Diabetes Institute. Dit protocol beschrijft het gebruik van een lichaam samenstelling analyzer en metabole dierlijke controlesysteem te karakteriseren lichaamssamenstelling en metabole parameters in muizen. Een model van de obesitas geïnduceerd door vetrijke voeding wordt gebruikt als een voorbeeld voor de toepassing van deze technieken.

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

A Protocol for Housing Mice in an Enriched Environment

Environmental enrichment (EE) is a housing environment for mice that boosts mental and physical health compared to standard laboratory housing. Our recent studies demonstrate that environmental enrichment decreases adiposity, increases energy expenditure, resists diet induced obesity, and causes cancer remission and inhibition in mice. EE typically consists of larger living space, a variety of &#8216;toys&#8217; to interact with, running wheels, and can include a number of other novel environmental changes. All of this fosters a more complex social engagement, cognitive and physical stimulations. Importantly, the toy location and type of toy is changed regularly, which encourages the mice to adapt to a frequently changing and novel environment. Many variables can be manipulated in EE to promote health effects in mice. Thus these approaches are difficult to control and must be properly managed to successfully replicate the associated phenotypes. Therefore, the goal of this video is to demonstrate how EE is properly set up and maintained to assure a complex, challenging, and controlled environment so that other researchers can easily reproduce the protective effects of EE against obesity and cancer.

Environmental enrichment for mice requires a complex and challenging setup, as well as comprehensive husbandry and handling techniques to assure robust metabolic and anti-cancer effects in the mice. This protocol provides detailed procedures to reproduce the above mentioned effects in mice.

Environmental enrichment (EE) is a housing environment for mice that boosts mental and physical health compared to standard laboratory housing. Our recent ...

Behavior

Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Establishing a system of procedures to qualitatively and quantitatively characterize in vivo and in vitro hepatic steatosis is important for metabolic study in the liver. Here, numerous assays are described to comprehensively measure the phenotype and parameters of hepatic steatosis in mouse and hepatocyte models. 
Combining the physiological, histological, and biochemical methods, this system can be used to assess the progress of hepatic steatosis. In vivo, the measurements of body weight and nuclear magnetic resonance (NMR) provide a general understanding of mice in a non-invasive manner. Hematoxylin and Eosin (H&amp;E) and Oil Red O staining determine the histological morphology and lipid deposition of liver tissue under nutrient overload conditions, such as high-fat diet feeding. Next, the total lipid contents are isolated by chloroform/methanol extraction, which are followed by a biochemical analysis for triglyceride and cholesterol. Moreover, mouse primary hepatocytes are treated with high glucose plus insulin to stimulate lipid accumulation, an efficient in vitro model to mimic diet-induced hyperglycemia and hyperinsulinemia in vivo. Then, the lipid deposition is measured by Oil Red O staining and chloroform/methanol extraction. Oil Red O staining determines intuitive hepatic steatotic phenotypes, while the lipid extraction analysis determines the parameters that can be analyzed statistically. The present protocols are of interest to scientists in the fields of fatty liver diseases, insulin resistance, and type 2 diabetes.

Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Here, optimized methods to generate in vivo and in vitro models of hepatic steatosis and to analyze the steatotic phenotypes and related physiological parameters are described.

Establishing a system of procedures to qualitatively and quantitatively characterize in vivo and in vitro hepatic steatosis is important for metabolic ...

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 ...

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

The nematode C. elegans has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans metabolic research.

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

We present robust biochemical and microscopic methods for studying Caenorhabditis elegans lipid stores. A rapid, simple, fixing-staining procedure for fluorescent lipid droplet imaging leverages the spectral properties of the lipophilic dye Nile red. We then present biochemical measurement of triglycerides and phospholipids using solid phase extraction and gas chromatography-mass spectrometry.

The nematode C. elegans has emerged as an important  model for the study of conserved genetic pathways regulating fat metabolism as  it relates to human ...

Biology

Palatable Western-style Cafeteria Diet as a Reliable Method for Modeling Diet-induced Obesity in Rodents

Obesity is rapidly increasing in incidence in developed and developing countries and is known to induce or exacerbate many diseases.&#160;The health burden of obesity and its comorbid conditions highlight the need for better understanding of its pathogenesis, yet ethical constraints limit studies in humans. To this end externally valid models of obesity in laboratory animals are essential for the understanding of being overweight and obesity.&#160;While many species have been used to model the range of changes that accompany obesity in humans, rodents are most commonly used. Our laboratory has developed a western-style cafeteria diet that consistently leads to considerable weight gain and markers of metabolic disease in rodents. The diet exposes rodents to a variety of highly palatable foods to induce hyperphagia, modeling the modern western food environment. This diet rapidly induces weight gain and body fat accumulation in rats allowing for the study of effects of overeating and obesity. While the cafeteria diet may not provide the same control over macronutrient and micronutrient profile as purified high-fat or high-fat, high-sugar diets, the cafeteria diet typically induces a more severe metabolic phenotype than that observed with purified diets and is more in line with metabolic disturbances observed in the overweight and obese human population.

This protocol describes the use of a highly palatable, western-style cafeteria diet to model overeating and obesity in rodents. Here, we provide a detailed outline of food selection, preparation and measurement, and explain methodological factors that assist in generating a robust and reproducible phenotype.

Obesity is rapidly increasing in incidence in developed and developing countries and is known to induce or exacerbate many diseases.&#160;The health ...

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 ...

Assessing Whole-Body Lipid-Handling Capacity in Mice

Assessing lipid metabolism is a cornerstone of evaluating metabolic function, and it is considered essential for in vivo metabolism studies. Lipids are a class of many different molecules with many pathways involved in their synthesis and metabolism. A starting point for evaluating lipid hemostasis for nutrition and obesity research is needed. This paper describes three easy and accessible methods that require little expertise or practice to master, and that can be adapted by most labs to screen for lipid-metabolism abnormalities in mice. These methods are (1) measuring several fasting serum lipid molecules using commercial kits (2) assaying for dietary lipid-handling capability through an oral intralipid tolerance test, and (3) evaluating the response to a pharmaceutical compound, CL 316,243, in mice. Together, these methods will provide a high-level overview of lipid handling capability in mice.

This paper provides three easy and accessible assays for assessing lipid metabolism in mice.

Assessing lipid metabolism is a cornerstone of evaluating metabolic function, and it is considered essential for in vivo metabolism studies. Lipids are a ...

Identification and Dissection of Diverse Mouse Adipose Depots

Adipose tissues are complex organs with a wide array of functions, including storage and mobilization of energy in response to local and global needs, uncoupling of metabolism to generate heat, and secretion of adipokines to regulate whole-body homeostasis and immune responses. Emerging research is identifying important regional differences in the developmental, molecular, and functional profiles of adipocytes located in discrete depots throughout the body. Different properties of the depots are medically relevant since metabolic diseases often demonstrate depot-specific effects. This protocol will provide investigators with a detailed anatomic atlas and dissection guide for the reproducible and accurate identification and excision of diverse mouse adipose tissues. Standardized dissection of discrete adipose depots will allow detailed comparisons of their molecular and metabolic characteristics and contributions to local and systemic pathologic states under various nutritional and environmental conditions.

Adipocytes exist in discrete depots and have diverse roles within their unique microenvironments. As regional differences in adipocyte character and function are uncovered, standardized identification and isolation of depots is crucial for advancement of the field. Herein, we present a detailed protocol for the excision of various mouse adipose depots.

Adipose tissues are complex organs with a wide array of functions, including storage and mobilization of energy in response to local and global needs, ...

Assessment of the Metabolic Effects of Isocaloric 2:1 Intermittent Fasting in Mice

Intermittent fasting (IF), a dietary intervention involving periodic energy restriction, has been considered to provide numerous benefits and counteract metabolic abnormalities. So far, different types of IF models with varying durations of fasting and feeding periods have been documented. However, interpreting the outcomes is challenging, as many of these models involve multifactorial contributions from both time- and calorie-restriction strategies. For example, the alternate day fasting model, often used as a rodent IF regimen, can result in underfeeding, suggesting that health benefits from this intervention are likely mediated via both caloric restriction and fasting-refeeding cycles. Recently, it has been successfully demonstrated that 2:1 IF, comprising 1 day of fasting followed by 2 days of feeding, can provide protection against diet-induced obesity and metabolic improvements without a reduction in overall caloric intake. Presented here is a protocol of this isocaloric 2:1 IF intervention in mice. Also described is a pair-feeding (PF) protocol required to examine a mouse model with altered eating behaviors, such as hyperphagia. Using the 2:1 IF regimen, it is demonstrated that isocaloric IF leads to reduced body weight gain, improved glucose homeostasis, and elevated energy expenditure. Thus, this regimen may be useful to investigate the health impacts of IF on various disease conditions.

The current article describes a detailed protocol for isocaloric 2:1 intermittent fasting to protect and treat against obesity and impaired glucose metabolism in wild-type and ob/ob mice.

Intermittent fasting (IF), a dietary intervention involving periodic energy restriction, has been considered to provide numerous benefits and counteract ...

Determining Basal Energy Expenditure and the Capacity of Thermogenic Adipocytes to Expend Energy in Obese Mice

Energy expenditure measurements are necessary to understand how changes in metabolism can lead to obesity. Basal energy expenditure can be determined in mice by measuring whole-body oxygen consumption, CO2 production, and physical activity using metabolic cages. Thermogenic brown/beige adipocytes (BA) contribute significantly to rodent energy expenditure, particularly at low ambient temperatures. Here, measurements of basal energy expenditure and total BA capacity to expend energy in obese mice are described in two detailed protocols: the first explaining how to set up the assay to measure basal energy expenditure using analysis of covariance (ANCOVA), a necessary analysis given that energy expenditure co-varies with body mass. The second protocol describes how to measure BA energy expenditure capacity in vivo in mice. This procedure involves anesthesia, needed to limit expenditure caused by physical activity, followed by the injection of beta3-adrenergic agonist, CL-316,243, which activates energy expenditure in BA. These two protocols and their limitations are described in sufficient detail to allow a successful first experiment.

This manuscript describes a protocol to measure the basal metabolic rate and the oxidative capacity of thermogenic adipocytes in obese mice.

Energy expenditure measurements are necessary to understand how changes in metabolism can lead to obesity. Basal energy expenditure can be determined in ...

Body Composition and Metabolic Caging Analysis in High Fat Fed Mice

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