ML is funded by the Department of Medicine at UCLA, pilot grants from P30 DK 41301 (UCLA:DDRC NIH) and P30 DK063491 (UCSD-UCLA DERC).

All experiments were approved by the Institutional Animal Care and Use Committee at the University of California, Los Angeles (UCLA). Mice were administered their diet and water ad libitum in the metabolic cage, housed in a temperature-controlled environment (~21-22 or 30 &#176;C) with a 12h light/dark cycle. 8 week-old female mice fed a high-fat diet or chow diet for 8 weeks were used for this study.
1. Measurement of the Basal Metabolic Rate (BMR)
<ol>
	<li>Measure the total body weight of the mouse using a weight scale with accuracy in the range of 0.1 g. 
	NOTE: This must be done before housing the mice in metabolic cages and after the 2-3 days of acclimation period to the metabolic cages.</li>
	<li>Measure the body composition, including fat and lean mass in non-anesthetized mice, using an appropriate body composition analysis system (see Table of Materials). 
	NOTE: These measurements are needed to determine energy expenditure and are executed in parallel to total body weight measurements (step 1.1).</li>
	<li>Set up the metabolic cages and start the acclimation period. 
	NOTE: The metabolic cages system includes an enclosure that allows the user to control the housing temperature and light of 12 cages (Figure 1A,B). Each cage has a water bottle, a feeder, and a grid (Figure 1C). The grid separates the mouse from the bottom of the cage, allowing feces collection. Once the cage is installed on each pre-determined space, a lid that seals the cage includes the water bottle slot, the tubing sampling air, the airflow system, and the physical activity sensor (Figure 1D).
	<ol>
		<li>Turn on the temperature enclosure, the airflow system, and computer 2 h before starting the assay.</li>
		<li>After 2 h, open the software controlling the enclosure (see Table of Materials) and the airflow and let the software test the computer&#39;s communication with the equipment. 
		NOTE: Oxymax software was used for the present work.</li>
		<li>Once communication is established, click File, then Open Experiment Configuration (Figure 2A) and select the experiment configuration predesigned by the vendor (or set up from a previous assay).</li>
		<li>Click on Experiment, then click on Properties, which will open the Experiment Properties window (Figure 2B).</li>
		<li>In the Properties window, set up the parameters of the environmental enclosure, including the Ambient Temperature (21 &#176;C) and the 12 h Light cycles. 
		NOTE: Keeping the software open and running allows air to flow into the cages and the enclosure to maintain the selected temperature and light cycles. Thus, the entire system can operate with mice inside the cages for several days, even without measuring oxygen and CO2.</li>
		<li>Click on Experiment, then click on Setup, and the Experiment Setup window will open, where the parameters of each metabolic cage are defined.</li>
		<li>Assign each mouse ID to the individual cage where the mouse is housed (Figure 2C).</li>
		<li>Include the lean mass or total body weight for each mouse only if no differences in body weight are observed between groups. 
		NOTE: Obtaining raw values of oxygen consumption and energy expenditure facilitates ANCOVA analyses.</li>
		<li>Set the airflow rate to the metabolic cage at 0.5-0.6 L/min.</li>
		<li>On the Experiment Setup window, select the file saving path and name. Select the backup directory (Figure 2D).</li>
		<li>Add a pre-weighted amount of food to the feeders that covers at least food intake for 1 day. 
		NOTE: If the cages have integrated scales, the food can be added directly, and the software will record it.</li>
		<li>Add the water bottles. Check that the bottle is sealed correctly and does not leak.</li>
		<li>24 h after adding the food, weigh the food that is left on the cage. 
		NOTE: The grams of food added minus the grams of food left will measure food intake.</li>
		<li>Start the oxygen, CO2, and activity measurements (step 1.4.10) once the food intake values are the same as those of mice housed in regular cages. 
		NOTE: Here, the acclimation period (usually 2-3 days) is completed, and energy expenditure measurements can start.</li>
	</ol>
	</li>
	<li>Indirect calorimetry and activity measurements to assess energy expenditure
	<ol>
		<li>Measure the body weight, fat, and lean mass of all mice before starting the measurements. 
		NOTE: These are the body weight and lean mass values used to perform ANCOVA analyses.</li>
		<li>Calibrate the CLAMS system&#39;s O2 and CO2 Zirconia-based detector (see Table of Materials) with the recommended oxygen concentration; always re-calibrate the detector before starting a new experiment.</li>
		<li>Use a calibration gas of known composition (20.50% oxygen and 0.50% CO2). 
		NOTE: Gas suppliers often refer to this gas as &#34;Primary Standard Grade.&#34;</li>
		<li>Turn on and ensure that the tank output pressure is at 5-10 psi.</li>
		<li>Open the Calibration Utility software for calibrating and testing the gas sensors (Figure 2E). Click on Experiment, then Calibrate.</li>
		<li>Press Start. Then, wait for the sensors to be tested and for the software to ask the user to turn the knobs of the gas sensor (Figure 2F) until the value of O2 identity is 1 (Figure 2G-H). Click Next when the step is complete. 
		NOTE: If the Calibration Utility performs all current steps, the calibration will automatically proceed to the next step when the progress bar is filled.</li>
		<li>Verify the calibration results when all steps have been completed, and the results are presented.</li>
		<li>Turn off the calibration gas.</li>
		<li>Change food and add sufficient food&#160;for a 48-72 h period. 
		NOTE: Although cages could be opened during the energy expenditure measurements to monitor the body weight and change the food daily, mice can be stressed by these manipulations, and measurements are lost when the cages are opened. Thus, it is recommended to avoid any manipulation during the measurement period.</li>
		<li>In the software, click on Experiment and then Run to start the oxygen, CO2, and activity measurements (Figure 3A). 
		NOTE: The execution of the measurements can be tracked in real-time in a box located at the bottom left section of the software (red rectangle, Figure 3B). The red rectangle in Figure 3B shows that the system measures cage #1 at interval #3, namely the 3rd measurement. One measurement in one cage can take about 1 min. Thus, with 12 cages connected, oxygen consumption can be measured approximately every 12 min. Continuous measurements for a minimum of 48 h are recommended.</li>
		<li>Stop the experiment by clicking Experiment and&#160;then Stop (Figure 3C).</li>
		<li>Open the cages, weigh the mice and the food. Collect the feces to calculate the number of calories and lipids excreted over the 48-72 h measurements period. 
		NOTE: Feces can be stored at -20 &#176;C for later analyses. These cages cannot be effectively used to collect urine.</li>
		<li>Click on Experiments, then Export, and Export all subjects as a CSV file (Figure 3D). 
		NOTE: To facilitate ANCOVA analyses, it is essential to export raw Oxygen consumption (VO2) and CO2 production (VCO2)&#160;values without being normalized by the body weight.</li>
	</ol>
	</li>
	<li>Data analysis and quality control
	<ol>
		<li>In the exported CSV sheet (Figure 3D) (from step 1.4.13), use the raw values of the oxygen consumption (VO2) and CO2&#160;production (VCO2) measured every 12 min in the 2-3 day period that are automatically listed by the software and include a timestamp, namely the hour and date when they were measured. 
		NOTE: VO2 and VCO2 values will be automatically corrected if body weight or lean mass values are added.</li>
		<li>In the exported CSV sheet, use the raw values of the respiratory exchange ratio (RER: VCO2/VO2) that are automatically calculated and listed by the software according to their timestamp. 
		NOTE: Values close to 1 show that the mouse primarily oxidizes carbohydrates, while values closer to 0.7 represent that the mouse is mainly oxidizing fat. RER above 1 can occur during anaerobic exercise, as the body expels more CO2 to compensate for the acidosis caused by lactate. RER higher than 1 can indicate stress. The exported CSV file also contains the raw values from Energy expenditure (EE) or Heat production in calories per minute per mouse, measured every 12 min in the 2-3 days. Here, all the listed values include a timestamp.</li>
		<li>As single EE values per mouse are needed for an ANCOVA, average the EE values recorded between 09:00-16:00 for the light (day) phase and 19:00-04:00 for the dark (night) phase per mouse and day. 
		NOTE: This can be done manually using Excel or Graph Pad. Selecting these two-time windows avoids averaging the intermediate, gradual, and unstable EE values associated with the light-dark phase transition.</li>
		<li>For a 48 h period, calculate the average of the two daylight values and the two dark phase values per mouse using Excel or Graph Pad.</li>
		<li>To quantify the total physical activity, use Excel or Graph Pad to sum the x, y, and z beam break counts measured in the metabolic cages and listed in the CSV file for each mouse. 
		NOTE: The x,y,z total activity is calculated by doing first the average value of each X, Y, Z per mouse and cycle. Then, the sum of each average X, Y, Z values per mouse and cycle is determined to plot the data as in Figure 5E&#160;(being the average of 2 days).</li>
		<li>Alternatively, represent the data showing each measurement value over time, which generates curves that illustrate the changes in EE during the transition from light to dark cycles. 
		&#8203;NOTE: Please refer to the discussion section on how and when to perform ANCOVA analyses, and the different formulas used to calculate VO2, VCO2, and EE are provided in Supplementary File 1.</li>
	</ol>
	</li>
</ol>
2. Measurement of the capacity of thermogenic adipocytes to expend energy
<ol>
	<li>Set up the measurements and mouse treatments. See step 1 for details on the experimental preparations to monitor oxygen consumption, as thermogenic capacity in adipocytes is determined indirectly by oxygen consumption following steps 2.1.1-2.2.2. 
	NOTE: This protocol requires mouse anesthesia and acute treatment with the beta-3 receptor agonist CL-316,243 (see Table of Materials), giving a rapid assessment of BA thermogenic capacity.
	<ol>
		<li>Perform body composition analysis and weigh the mice. Turn on the CLAMS, set up the temperature at 30 &#176;C (thermoneutrality), and wait for 2 h for the whole system to warm up.</li>
		<li>Set up the rest of the assay conditions, including light, assign mouse ID to each cage and add body weight value of each mouse in the corresponding cage if no difference in body weight is observed between groups.</li>
		<li>Calibrate the oxygen/CO2 detector as in steps 1.4.2-1.4.7.</li>
		<li>Start the Experiment in the software.</li>
		<li>Inject each mouse with pentobarbital (60-120 mg/kg) and place each mouse in their assigned metabolic cage (step 2.1.2). 
		NOTE: The dose of pentobarbital required to maintain mice asleep at thermoneutrality (30 &#176;C) varies with the mouse strain and genotype. It is recommended to test different pentobarbital doses from 50 to 120 mg/kg and chose the one keeping the mouse anesthetized during 2-3 h at 30 &#176;C. Efficient anesthesia is essential to remove the contribution of physical activity to energy expenditure.</li>
		<li>To ensure anesthesia, observe mice after pentobarbital injection until they are completely asleep and their decreased oxygen consumption rates become steady.</li>
		<li>Wait to obtain at least 3 stable consecutive oxygen consumption rates before injecting CL-316,243. 
		NOTE: While waiting for the stabilization of oxygen consumption, prepare the syringes with CL-316,243 for each mouse (1 mg/kg).</li>
		<li>Open cage #1 and inject CL-316,243 subcutaneously immediately after one VO2 and VCO2 measurement occurred in cage #1. Return the mouse to cage #1 immediately after injection. 
		NOTE: Measurements are being indicated in real-time in the bottom-left section of the software (Figure 3B, red rectangle).</li>
		<li>Wait for cage #2 to be measured (Figure 3B, red rectangle) and then proceed as in step 2.1.8 for cage #2. 
		NOTE: Injecting CL-316,243 immediately after a measurement allows maintaining the time constant between injections. For instance, if there are 12 mice/cages running, with measurements collected in individual cages sequentially and the collection lasting 55 s per cage, then you should inject one mouse every minute. With these injection rates, the first measurement will occur after 12 min after injection in all 12 cages.</li>
		<li>Continue the energy expenditure measurements until the energy expenditure values plateau for 5-6 consecutive measurements, usually 90-180 min after injection. 
		NOTE: Mice could wake up from the anesthesia during the experiments. These mice need to be removed from the analysis. Therefore, testing pentobarbital doses beforehand will increase the efficiency of the studies.</li>
		<li>Stop the energy expenditure measurements, but keep mice at their cages at 30 &#176;C, until they wake up.</li>
		<li>After mice are fully awake, inspect the health of the mice and return them to their initial cages.</li>
		<li>Export the data of each mouse as a CSV file using the equipment software, as described in section 1.4.13.</li>
	</ol>
	</li>
	<li>Data analysis 
	NOTE: The data analysis was performed by Excel or Graphpad
	<ol>
		<li>Plot the 3-5 consecutive values of VO2, VCO2, and EE that are stable and constant over time, as these are the values representing the metabolic rate when mice are fully anesthetized.</li>
		<li>Then, plot the first and the following consecutive VO2, VCO2, and EE measurements obtained after injection. 
		NOTE: The absolute values of EE and the fold increase in EE induced by injection indicate BA thermogenic function7.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Rolfe, D. F., Brown, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+energy+utilization+and+molecular+origin+of+standard+metabolic+rate+in+mammals.">Cellular energy utilization and molecular origin of standard metabolic rate in mammals.</a> Physiological Reviews. 77 (3), 731-758 (1997).</li><li>Heymsfield, S. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+energy+expenditure:+advances+in+organ-tissue+prediction+models.">Human energy expenditure: advances in organ-tissue prediction models.</a> Obesity Reviews. 19 (9), 1177-1188 (2018).</li><li>Kummitha, C. M., Kalhan, S. C., Saidel, G. M., Lai, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relating+tissue/organ+energy+expenditure+to+metabolic+fluxes+in+mouse+and+human:+experimental+data+integrated+with+mathematical+modeling.">Relating tissue/organ energy expenditure to metabolic fluxes in mouse and human: experimental data integrated with mathematical modeling.</a> Physiological Reports. 2 (9), 12159 (2014).</li><li>Tschop, M. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+to+analysis+of+mouse+energy+metabolism.">A guide to analysis of mouse energy metabolism.</a> Nature. 9 (1), 57-63 (2011).</li><li>Mina, A. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CalR:+A+Web-Based+Analysis+Tool+for+Indirect+Calorimetry+Experiments.">CalR: A Web-Based Analysis Tool for Indirect Calorimetry Experiments.</a> Cell Metabolism. 28 (4), 656-666 (2018).</li><li>Shum, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ABCB10+exports+mitochondrial+biliverdin,+driving+metabolic+maladaptation+in+obesity.">ABCB10 exports mitochondrial biliverdin, driving metabolic maladaptation in obesity.</a> Science Translational Medicine. 13 (594), (2021).</li><li>Assali, E. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NCLX+prevents+cell+death+during+adrenergic+activation+of+the+brown+adipose+tissue.">NCLX prevents cell death during adrenergic activation of the brown adipose tissue.</a> Nature Communication. 11 (1), 3347 (2020).</li><li>Clark, J. D., Gebhart, G. F., Gonder, J. C., Keeling, M. E., Kohn, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Special+Report:+The+1996+Guide+for+the+Care+and+Use+of+Laboratory+Animals.">Special Report: The 1996 Guide for the Care and Use of Laboratory Animals.</a> ILAR Journal. 38 (1), 41-48 (1997).</li><li>Schena, G., Caplan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Everything+You+Always+Wanted+to+Know+about+beta3-AR+*+(*+But+were+afraid+to+ask).">Everything You Always Wanted to Know about beta3-AR * (* But were afraid to ask).</a> Cells. 8 (4), 357 (2019).</li><li>Granneman, J. G., Burnazi, M., Zhu, Z., Schwamb, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=White+adipose+tissue+contributes+to+UCP1-independent+thermogenesis.">White adipose tissue contributes to UCP1-independent thermogenesis.</a> American Journal of Physiology-Endocrinology and Metabolism. 285 (6), 1230-1236 (2003).</li><li>Szentirmai, E., Kapas, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+the+brown+adipose+tissue+in+beta3-adrenergic+receptor+activation-induced+sleep,+metabolic+and+feeding+responses.">The role of the brown adipose tissue in beta3-adrenergic receptor activation-induced sleep, metabolic and feeding responses.</a> Scientific Reports. 7 (1), 958 (2017).</li></ol>

The authors declare no conflict of interest to this protocol paper. M.L. is a co-founder and consultant for Enspire Bio LLC.

Indirect calorimetry has been used for years to assess whole-body energy expenditure4. This protocol described herein provides a straightforward method of measuring the basal metabolic rate and determining BA thermogenic capacity in vivo using metabolic cages.
The indirect calorimetry method described here confirms that dividing energy expenditure values by body weight values can be misleading. For example, it can conclude that energy expenditure is systematica...

Metabolism can be defined as the integration of the biochemical reactions responsible for nutrient uptake, storage, transformation, and breakdown that cells use to grow and perform their functions. Metabolic reactions transform the energy contained in nutrients into a form that can be used by cells to synthesize new molecules and execute work. These biochemical reactions are inherently inefficient in transforming this energy into a usable form to sustain life1. Such inefficiency results in energy dissipation in the form of heat, with this heat production being used to quantify the Standard Metabolic Rate (SMR) of an organism1. The Standard condition was classically defined as heat production occurring in an awake but resting adult, not ingesting or digesting food, at thermoneutrality and without any stress1. The Basal Metabolic Rate (BMR) or basal energy expenditure in mice is referred to as the SMR but in individuals ingesting and digesting food under mild thermal stress (ambient temperatures 21-22 &#176;C)1. The challenges and difficulties of directly measuring heat production made indirect calorimetry, namely calculating heat production from oxygen consumption measurements, to become the most popular approach to determine the BMR. Calculating the BMR from oxygen consumption is possible because the oxidation of nutrients by mitochondria to synthesize ATP is responsible for 72% of the total oxygen consumed in an organism, with 8% of total oxygen consumption also occurring in mitochondria but without generating ATP (uncoupled respiration)1. The majority of the remaining 20% of oxygen consumed can be attributed to nutrient oxidation in other subcellular locations (peroxisomal fatty acid oxidation), anabolic processes, and reactive oxygen species formation1. Thus, in 1907, Lusk established an equation, based on empirical measurements, widely used to transform oxygen consumption and CO2 production into energy dissipation as heat. In humans, the brain accounts for ~25% of the BMR, the musculoskeletal system for ~18.4%, the liver for ~20 %, the heart for ~10%, and the adipose tissue for ~3-7%2. In mice, the tissue contribution to BMR is slightly different, with the brain representing ~6.5%, the skeletal muscle ~13%, the liver ~52%, the heart ~3.7%, and adipose tissue ~5%3.
Remarkably, the biochemical reactions defining the BMR are not fixed and change in response to different needs, such as external work (physical activity), development (tissue growth), internal stresses (counteracting infections, injuries, tissue turnover), and changes in ambient temperature (cold defense)1. Some organisms actively recruit processes to generate heat in cold exposure, implying that heat produced by metabolism is not just an accidental byproduct. Instead, evolution selected regulatory mechanisms that could specifically upregulate heat production by changing the rate of metabolic reactions1. Thus, these same oxygen consumption measurements can be used to determine the capacity of an organism to generate heat in response to cold.
Two major processes contribute to heat generation upon cold exposure. The first one is shivering, which generates heat by increasing mitochondrial oxidative phosphorylation and glycolysis in muscle to cover the physical work done by involuntary muscle contraction. Therefore, cold exposure will increase oxygen consumption in muscles1. The second is Non-Shivering Thermogenesis, which occurs through an increase in oxygen consumption in brown and beige adipocytes (BA). Dissipation of energy into heat in BA is mediated by the mitochondrial uncoupling protein 1 (UCP1), which allows proton re-entry into the mitochondrial matrix, decreasing the mitochondrial proton gradient. The dissipation of the mitochondrial proton gradient by UCP1 increases heat production by the elevation in electron transfer and oxygen consumption and the energy released by proton dissipation per se without generating ATP (uncoupled). Moreover, thermogenic BA can recruit additional mechanisms that elevate oxygen consumption without causing a large dissipation in the proton gradient, by activating futile oxidative ATP synthesis and consumption cycles. The metabolic cages described here, namely the CLAMS-Oxymax system from Columbus Instruments, offer the possibility to measure energy expenditure at different ambient temperatures. However, to determine BA thermogenic capacity using whole-body oxygen consumption measurements, one needs to: (1) eliminate the contribution of shivering, and other non-BA metabolic processes to energy expenditure, and (2) specifically activate BA thermogenic activity in vivo. Thus, a second protocol describes how to selectively activate BA in vivo using pharmacology in anesthetized mice at thermoneutrality (30 &#176;C), with anesthesia and thermoneutrality limiting other non-BA thermogenic processes (i.e., physical activity). The pharmacological strategy to activate BA is treating mice with the &#946;3-adrenergic receptor agonist CL-316,246. The reason is that cold exposure promotes a sympathetic response releasing norepinephrine to activate &#946;-adrenergic receptors in BA, which activates UCP1 and fat oxidation. Furthermore, &#946;3-adrenergic receptor expression is highly enriched in adipose tissue in mice.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>CLAMS-Oxymax System</td>
			<td>Columbus Instruments</td>
			<td>CLAMS-center feeder-ENC</td>
			<td>Including enviromental enclosure and Zirconia oxygen sensor</td>
		</tr>
		<tr>
			<td>Desktop PC with Oxymax Software</td>
			<td>HP/Columbus</td>
			<td>N/A</td>
			<td>PC needed to be purchased separately</td>
		</tr>
		<tr>
			<td>Drierite jug (Calcium Sulfate with Cobalt Chloride Indicator)</td>
			<td>Fisher Scientific</td>
			<td>23-116681</td>
			<td>Needed to dry the gas entering the oxygen sensor, humidity can damage the sensor</td>
		</tr>
		<tr>
			<td>NMR for body composition</td>
			<td>Echo-MRI</td>
			<td>Echo-MRI 100</td>
			<td>Measure lean and fat mass in alive mice. It is necessary for ANCOVA analyses.</td>
		</tr>
		<tr>
			<td>CL-316-243</td>
			<td>Sigma</td>
			<td>C5976</td>
			<td>Injected to the mice subcutaneously to activate thermogenesis</td>
		</tr>
		<tr>
			<td>High fat diet</td>
			<td>Research Diets</td>
			<td>D12266B</td>
			<td>Provided to the mice prior and during measurements</td>
		</tr>
		<tr>
			<td>Pentobarbital/Nembutal</td>
			<td>Pharmacy at DLAM</td>
			<td>N/A</td>
			<td>Anesthesia for the mice</td>
		</tr>
		<tr>
			<td>Primary standard grade gas (tank and regulator)</td>
			<td>Praxair</td>
			<td>NI CD5000O6P-K/PRS 2012-2331-590</td>
			<td>20.50% Oxygen, 0.50% CO2 balanced with nitrogen used for calibration</td>
		</tr>
	</tbody>
</table>

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.

Figure 4 shows VO2, VCO2, Heat production/Energy expenditure (EE), Respiratory Exchange Ratio (RER), and X, Y, Z physical activity values obtained using the metabolic cages of the CLAMS system. The VO2 and VCO2 provided by the CLAMS system is the volume of gas (mL) per minute and can be already divided by the body weight or the lean mass values by entering these weight values in the CLAMS software before starting the measurements. However, body wei...

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

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 ...

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4.9K Views.  Universite Laval. This manuscript describes a protocol to measure the basal metabolic rate and the oxidative capacity of thermogenic adipocytes in obese mice.

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

Measuring Skeletal Muscle Thermogenesis in Mice and Rats

Skeletal muscle thermogenesis provides a potential avenue for better understanding metabolic homeostasis and the mechanisms underlying energy expenditure. Surprisingly little evidence is available to link the neural, myocellular, and molecular mechanisms of thermogenesis directly to measurable changes in muscle temperature. This paper describes a method in which temperature transponders are utilized to retrieve direct measurements of mouse and rat skeletal muscle temperature.
Remote transponders are surgically implanted within the muscle of mice and rats, and the animals are given time to recover. Mice and rats must then be repeatedly habituated to the testing environment and procedure. Changes in muscle temperature are measured in response to pharmacological or contextual stimuli in the home cage. Muscle temperature can also be measured during prescribed physical activity (i.e., treadmill walking at a constant speed) to factor out changes in activity as contributors to the changes in muscle temperature induced by these stimuli.
This method has been successfully used to elucidate mechanisms underlying muscle thermogenic control at the level of the brain, sympathetic nervous system, and skeletal muscle. Provided are demonstrations of this success using predator odor (PO; ferret odor) as a contextual stimulus and injections of oxytocin (Oxt) as a pharmacological stimulus, where predator odor induces muscle thermogenesis, and Oxt suppresses muscle temperature. Thus, these datasets display the efficacy of this method in detecting rapid changes in muscle temperature.

Mice and rats are surgically implanted with remote temperature transponders and then habituated to the testing environment and procedure. Changes in muscle temperature are measured in response to pharmacological or contextual stimuli in the home cage or during prescribed physical activity (i.e., treadmill walking at a constant speed).

Skeletal muscle thermogenesis provides a potential avenue for better understanding metabolic homeostasis and the mechanisms underlying energy expenditure. ...

Human Brown Adipose Tissue Depots Automatically Segmented by Positron Emission Tomography/Computed Tomography and Registered Magnetic Resonance Images

Reliably differentiating brown adipose tissue (BAT) from other tissues using a non-invasive imaging method is an important step toward studying BAT in humans. Detecting BAT is typically confirmed by the uptake of the injected radioactive tracer 18F-Fluorodeoxyglucose (18F-FDG) into adipose tissue depots, as measured by positron emission tomography/computed tomography (PET-CT) scans after exposing the subject to cold stimulus. Fat-water separated magnetic resonance imaging (MRI) has the ability to distinguish BAT without the use of a radioactive tracer. To date, MRI of BAT in adult humans has not been co-registered with cold-activated PET-CT. Therefore, this protocol uses 18F-FDG PET-CT scans to automatically generate a BAT mask, which is then applied to co-registered MRI scans of the same subject. This approach enables measurement of quantitative MRI properties of BAT without manual segmentation. BAT masks are created from two PET-CT scans: after exposure for 2 hr to either thermoneutral (TN) (24 &#176;C) or cold-activated (CA) (17 &#176;C) conditions. The TN and CA PET-CT scans are registered, and the PET standardized uptake and CT Hounsfield values are used to create a mask containing only BAT. CA and TN MRI scans are also acquired on the same subject and registered to the PET-CT scans in order to establish quantitative MRI properties within the automatically defined BAT mask. An advantage of this approach is that the segmentation is completely automated and is based on widely accepted methods for identification of activated BAT (PET-CT). The quantitative MRI properties of BAT established using this protocol can serve as the basis for an MRI-only BAT examination that avoids the radiation associated with PET-CT.

The method presented here uses 18F-Fluorodeoxyglucose (18F-FDG) positron emission tomography/computed tomography (PET-CT) and fat-water separated magnetic resonance imaging (MRI), each scanned following 2 hr exposure to thermoneutral (24 &#176;C) and cold conditions (17 &#176;C) in order to map brown adipose tissue (BAT) in adult human subjects.

Reliably differentiating brown adipose tissue (BAT) from other tissues using a non-invasive imaging method is an important step toward studying BAT in ...

Medicine

Exploring Metabolite Exchange in Brown Adipose Tissue

Arteriovenous Metabolomics to Measure In Vivo Metabolite Exchange in Brown Adipose Tissue

Brown adipose tissue (BAT) plays a crucial role in regulating metabolic homeostasis through a unique energy expenditure process known as non-shivering thermogenesis. To achieve this, BAT utilizes a diverse menu of circulating nutrients to support its high metabolic demand. Additionally, BAT secretes metabolite-derived bioactive factors that can serve as either metabolic fuels or signaling molecules, facilitating BAT-mediated intratissue and/or intertissue communication. This suggests that BAT actively participates in systemic metabolite exchange, an interesting feature that is beginning to be explored. Here, we introduce a protocol for in vivo mouse-level optimized BAT arteriovenous metabolomics. The protocol focuses on relevant methods for thermogenic stimulations and an arteriovenous blood sampling technique using Sulzer's vein, which selectively drains interscapular BAT-derived venous blood and systemic arterial blood. Next, a gas chromatography-based metabolomics protocol using those blood samples is demonstrated. The use of this technique should expand the understanding of BAT-regulated metabolite exchange at the inter-organ level by measuring the net uptake and release of metabolites by BAT.

Author Spotlight: Illuminating New Avenues for Adipose Tissue Metabolism and Disease Prevention 

In this protocol, methods relevant for BAT-optimized arteriovenous metabolomics using GC-MS in a mouse model are outlined. These methods allow for the acquisition of valuable insights into BAT-mediated metabolite exchange at the organismal level.

Brown adipose tissue (BAT) plays a crucial role in regulating metabolic homeostasis through a unique energy expenditure process known as non-shivering ...

Biochemistry

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.

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 ...

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 ...

Infrared Thermography for the Detection of Changes in Brown Adipose Tissue Activity

Measuring brown adipose tissue (BAT) activity by positron emission tomography computed tomography (PET-CT) via the accumulation of 18F-fluorodeoxyglucose (FDG) after a meal or in obese or diabetic patients fails as the method of choice. The main reason is that 18F-FDG competes with the postprandial high glucose plasma concentration for the same glucose transporter on the membrane of BAT cells. In addition, BAT uses fatty acids as a source of energy as well, which is not visible with PET-CT and could be changed along with glucose concentration in obese and diabetic patients. Therefore, to estimate the physiological importance of BAT in animals and humans, a new infrared thermography method used in recent publications is applied.
After overnight fasting, BAT activity was measured by infrared thermography before and after a meal in human volunteers and&#160;female wild-type mice. The camera software calculates the object&#39;s temperature using distance from the object, skin emissivity, reflected room temperature, air temperature, and relative humidity. In mice, the shaved area above the BAT was a region of interest for which average and maximal temperatures were measured. The phase of the estrous cycle in female mice was determined after an experiment by vaginal smears stained with cresyl violet (0.1%) stain solution. In healthy volunteers, two skin areas of the neck were selected: the supraclavicular area (above the collarbone, where BAT cells are present) and the interclavicular area (between the collarbones, where there is no BAT tissue detected). BAT activity is determined by the subtraction of those two values. Also, the average and maximal temperatures of skin areas could be determined in animals and human participants.
Changes in BAT activity after a meal measured by infrared thermography, a non-invasive and more sensitive method, were shown to be sex, age, and phase of the estrous cycle dependent in laboratory animals. As part of diet-induced thermogenesis, BAT activation in humans was also proven to be sex, age, and body mass index dependent. Further determining the pathophysiological changes in BAT activity after a meal will be of great importance for participants with high glucose plasma concentrations (obesity and diabetes mellitus type 2), as well as in different laboratory animals (knock-out mice). This method is also a variable tool for determining possible activating drugs that could rejuvenate BAT activity.

Here, we present a protocol for measuring brown adipose tissue activity after a meal in humans and laboratory animals.

Measuring brown adipose tissue (BAT) activity by positron emission tomography computed tomography (PET-CT) via the accumulation of 18F-fluorodeoxyglucose ...

Using a Combination of Indirect Calorimetry, Infrared Thermography, and Blood Glucose Levels to Measure Brown Adipose Tissue Thermogenesis in Humans

In mammals, brown adipose tissue (BAT) is activated rapidly in response to cold in order to maintain body temperature. Although BAT has been studied greatly in small animals, it is difficult to measure the activity of BAT in humans. Therefore, little is known about the heat-generating capacity and physiological significance of BAT in humans, including the degree to which components of the diet can activate BAT. This is due to the limitations in the currently most used method to assess the activation of BAT-radiolabeled glucose (fluorodeoxyglucose or 18FDG) measured by positron emission tomography-computerized tomography (PET-CT).
This method is usually performed in fasted subjects, as feeding induces glucose uptake by the muscles, which can mask the glucose uptake into the BAT. This paper describes a detailed protocol for quantifying total-body human energy expenditure and substrate utilization from BAT thermogenesis by combining indirect calorimetry, infrared thermography, and blood glucose monitoring in carbohydrate-loaded adult males. To characterize the physiological significance of BAT, measures of the impact of BAT activity on human health are critical. We demonstrate a protocol to achieve this by combining carbohydrate loading and indirect calorimetry with measurements of supraclavicular changes in temperature. This novel approach will help to understand the physiology and pharmacology of BAT thermogenesis in humans.

Here, we present a protocol to quantify the physiological significance of the impact of brown adipose tissue (BAT) activity on human metabolism. This is achieved by combining carbohydrate loading and indirect calorimetry with measurements of supraclavicular changes in temperature. This novel approach can help develop a pharmacological target for BAT thermogenesis in humans.

In mammals, brown adipose tissue (BAT) is activated rapidly in response to cold in order to maintain body temperature. Although BAT has been studied ...

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 ...

Visualization and Quantification of Brown and Beige Adipose Tissues in Mice using [18F]FDG Micro-PET/MR Imaging

Brown and beige adipocytes are now recognized as potential therapeutic targets for obesity and metabolic syndromes. Non-invasive molecular imaging methods are essential to provide critical insights into these thermogenic adipose depots. Here, the protocol presents a PET/MR imaging-based method to evaluate the activity of brown and beige adipocytes in mouse interscapular brown adipose tissue (iBAT) and inguinal subcutaneous white adipose tissue (iWAT). Visualization and quantification of the thermogenic adipose depots were achieved using [18F]FDG, the non-metabolizable glucose analog, as the radiotracer, when combined with the precise anatomical information provided by MR imaging. The PET/MR imaging was conducted 7 days after cold acclimation and quantitation of [18F]FDG signal in different adipose depots was conducted to assess the relative mobilization of thermogenic adipose tissues. Removal of iBAT substantially increased cold-evoked [18F]FDG uptake in iWAT of the mice.

Visualization and Quantification of Brown and Beige Adipose Tissues in Mice using [18F]FDG Micro-PET/MR Imaging

Functional imaging and quantitation of thermogenic adipose depots in mice using a micro-PET/MR imaging-based approach.

Brown and beige adipocytes are now recognized as potential therapeutic targets for obesity and metabolic syndromes. Non-invasive molecular imaging methods ...