This work is supported by R15 DK097644 and R15 DK108668. We thank Dr. Chaitanya K Gavini and Dr. Megan Rich for prior contributions and Dr. Stanley Dannemiller for ensuring our&#160;compliance with institutional animal use guidelines. A special thank you to Dr. Tim Bartness for providing the fundamental research necessary to build this method and its associated studies. Figures 1A,&#160;C,&#160;D&#160;and Figure 2A were created using Biorender.com.

These methods can be applied to both rat and mouse models and were performed with institutional approval (Kent State University, IACUC Approval #359 and #340 CN 12-04).&#160;Prior to the implementation of the protocol, animals should be housed in conformance with the Guide for the Care and Use of Laboratory Animals.
1. Preparing the transponder reader
NOTE: Prior to use, the transponder reader must be set; the following steps only include the setting changes necessary for this study. This portion of the protocol is directly associated with the DAS-8027-IUS portable readers; other reader models should follow instructions provided by the manual to achieve programming results.
<ol>
	<li>Set Audio Beep to OFF.
	<ol>
		<li>Turn on the device by pressing the SCAN button and wait for the lighting to appear on the OLED display. Press and hold the BACK/MENU button to get to the menu screen.</li>
		<li>Using the NEXT/ENTER button, scroll through the options until OPERATIONAL SETUP. Here, toggle the up or down arrows to turn YES and open the operational submenu.</li>
		<li>Using the NEXT/ENTER button, scroll to AUDIO BEEP. As the default setting is ON, toggle the up or down arrows and change the setting to OFF.</li>
		<li>Press the NEXT/ENTER button to save this setting change.</li>
	</ol>
	</li>
	<li>Set Vibrate Upon Read to ON.
	<ol>
		<li>Follow step 1.1&#160;through step 1.2 or complete the next step directly after step 1.4.</li>
		<li>Using the NEXT/ENTER button, scroll to VIBRATE UPON READ. As the default setting is OFF, toggle the up and down arrows and change the setting to ON to feel, via vibration, when the reading has been completed regardless of being able to view the screen.</li>
	</ol>
	</li>
</ol>
2. Program transponders
NOTE: Each implanted transponder should first be programmed with an animal identification (animal ID or transponder ID). This nomenclature can be used as secondary identification for the test subject (e.g., four digits for mouse strain abbreviation, location of transponder, and an additional three to four digits to indicate animal number). Programming can be completed days in advance of surgery while keeping the transponders sterile prior to surgery.
<ol>
	<li>Enter the ID Code on the transponder.
	<ol>
		<li>Apply a booster coil to the reader head&#8212;a specific accessory for model DAS 8027-IUS, which helps in the programming procedure.</li>
		<li>Using a gloved hand, place the transponder (within the applicator) into the booster coil.</li>
		<li>Turn on the device by pressing the SCAN button and wait for the OLED display to light up. Press and hold the BACK/MENU button to get to the menu screen.</li>
		<li>Using the NEXT/ENTER button, scroll through the options until WRITE TRANSPONDER ID. Here, toggle the up or down arrows to turn YES.</li>
		<li>Using the NEXT/ENTER button, toggle to ENTER ID CODE.</li>
		<li>Use the up and down arrow keys to scroll through numbers and letters. Press NEXT/ENTER after each character selection to move to the following character.</li>
		<li>When the ID code is complete, press SCAN to write the transponder.</li>
		<li>Remove the transponder from the booster coil and repeat as necessary. Check that the transponder reads temperature changes by warming the enclosed transponders between gloved hands and measuring using the temperature scanner. 
		&#8203;NOTE: AUTO MULTI WRITE and SEQUENTIAL COUNT settings can be set to ON to allow for multiple or sequential transponder programming during a session. Each transponder should be tested during programming.</li>
	</ol>
	</li>
</ol>
3. Prepare &#34;home cage balls&#34;
<ol>
	<li>Place 5 cm x 5 cm odorless/control towel into a tea ball.</li>
	<li>Place these home cage balls in new home cages after surgery to begin habituating the animal to the method in which the contextual stimuli will be presented during testing. Replace these home cage balls every 2 weeks.</li>
</ol>
4. Surgery and postoperative care
<ol>
	<li>Weigh and record the subjects&#39; pre-surgery body weight. Using an induction chamber, provide anesthesia (e.g., 2-5% isoflurane) to the animal.</li>
	<li>Using electric clippers, completely shave the hind limb. Administer analgesia (e.g., 5 mg/kg of ketoprofen, s.c.) in compliance with institutional guidelines. 
	NOTE: Additional analgesia may be required if this procedure is combined with other surgical methods.</li>
	<li>Clean the area with 70% alcohol (or commercially available sterile alcohol wipe) and povidone-iodine wash (or commercially available sterile, individually wrapped betadine swabs) alternating at least three times, ending with povidone-iodine.</li>
	<li>Return the animal to the induction chamber, and anesthetize the animal to surgical levels. Then, set up the mouse in a face mask for continued exposure to anesthesia. Apply neomycin ophthalmic ointment to the eyes of the animal to prevent dryness while under anesthesia. 
	NOTE: The procedure should not start until the mouse shows no evidence of pain reception (i.e., corneal reflex, tail pinch response, toe pinch reflex).</li>
	<li>Using only surgical scissors, make a shallow cut through the skin on the right hind limb.</li>
	<li>Moving parallel to the gastrocnemius, place the sharp edge of a preprogrammed and uncapped sterile transponder into the incision. Ensure that the green plunger faces up and is visible. Continue pushing the transponder applicator into the incision until the opening of the transponder applicator is no longer visible. 
	NOTE: Do not accidentally press the green plunger on the transponder applicator during step 4.6. Premature discharge of the transponder will lead to improper placement.</li>
	<li>Turn the applicator 180&#176;, resulting in the green plunger facing down toward the mouse&#39;s limb, no longer visible to the experimenter. Push the transponder applicator into the final location. Once in ideal placement, adjacent to or partially enclosed in the gastrocnemius, push the green plunger, allowing the applicator&#39;s pressure to guide the investigator&#39;s hand back away from the mouse.</li>
	<li>Using forceps, hold together the opened skin and place a wound clip with a sterile autoclip or sterile suture. If needed, use absorbable sutures prior to the sterile autoclip to close the fascia layer. Using the transponder-reader, check the temperature of the mouse muscle.</li>
	<li>Remove the mouse from anesthesia and place it in a clean home cage placed atop a water-circulating heating pad set to low for recovery. Ensure that the home cage includes a tea ball with an odorless towel to begin habituation. 
	NOTE: The mouse should awaken from surgery within 15 min. Food can be placed at the bottom of the cage for easy access during recovery days.</li>
	<li>Postoperative care
	<ol>
		<li>Record mouse weights and temperatures daily using a transponder-reader for at least 2 days after surgery or until mice regain or stabilize body weight.</li>
		<li>Administer non-narcotic analgesia (e.g., 5 mg/kg of ketoprofen, s.c.) once daily to the mice for at least 2 days post surgery, with additional doses provided as needed. 
		NOTE: Mice and rats should fully recover within 5-8 days of surgery and can undergo habituation and testing procedures.</li>
	</ol>
	</li>
</ol>
5. Testing preparation - home cage
<ol>
	<li>Constructing risers 
	NOTE: The below step is based on 194 mm x 181 mm x 398 mm mouse filter-topped cages. To fit larger cages (e.g., a rat home cage), the width will need to be adjusted.
	<ol>
		<li>Cut the PVC pipe with a ratcheting PVC cutter into eight sections and assemble following Figure 1C. This will give an open tabletop structure that can hold approximately four cages. Make the desired number of risers.</li>
	</ol>
	</li>
	<li>Room setup
	<ol>
		<li>Assign a location to each riser within the testing room. Separate the risers set to receive different contextual stimuli (i.e., odors) by a minimum of 2 m to avoid confounding variables. 
		NOTE: Each mouse should have an assigned testing location within the testing room and on the physical risers as much as is feasible to avoid developing associations between different locations and thermogenic stimuli.</li>
		<li>Using magnetic strips, attach surgical sheets or gowns across the risers, creating a visual barrier between the researcher and the test subjects. Set this barrier to minimize temperature changes resulting from mouse activity when viewing experimenters moving toward the cage or around the testing room.</li>
		<li>(Optional) Place mirrors on the surface below the risers to ease viewing of the cage bottom during testing. 
		NOTE: Risers can be sanitized through a cage wash system. Cloth or surgical sheets should be laundered prior to habituation and testing.</li>
	</ol>
	</li>
	<li>Tea ball preparation
	<ol>
		<li>Prepare tea balls with control and PO towels (approximately 5 cm x 5 cm). To avoid cross-contamination, prepare control-odor tea balls first. 
		NOTE: Predator-odor towels should be pathogen-tested prior to use. These towels should also be contained, and materials that interact with them should be immediately sanitized (i.e., cage wash), preventing exposure of the odor to other animals.</li>
	</ol>
	</li>
</ol>
6. Temperature testing - home cage
NOTE: Animals need to be habituated to the entire testing procedure, excluding experimental contextual, or pharmacological stimuli. This should be completed a minimum of 4x before testing.
<ol>
	<li>Transfer the animals to the prepared testing&#160;room. Place the animals in a preassigned location on the riser. This location should be the same throughout all habituation and testing procedures.</li>
	<li>Remove the &#34;home cage ball&#34; from the mouse home cage and re-cover the cages with a cloth or surgical sheet. Allow the mice to acclimate to the testing space for 1-2 h.</li>
	<li>After acclimation is complete, use the scanner to measure and record the baseline temperature of each subject. Avoid manipulating the cloth coverings during measurements. 
	NOTE: Pharmacological agents can be applied here. Wait time post injection or application can be added as needed before testing. Recording a secondary baseline directly before testing is recommended after the addition of a pharmacological agent to monitor the response to pharmacological stimuli. If odor response is not being tested, temperature measurements of the mice can begin directly after injection. Randomization should be employed when providing any stimuli.</li>
	<li>Uncover the cage and place the tea ball (control or PO) onto the floor of the home cage. Replace the cage lid and cloth covering.</li>
	<li>Begin the stopwatch. Measure the temperatures of the test subjects in the same order of tea ball placement. Record temperatures and clock time of measurements following the desired time points.</li>
	<li>When the experiment is complete, remove the treatment ball. Place the mice that received PO in a new home cage with the original &#34;home cage ball&#34;. Return the &#34;home cage ball&#34; to the cage of the mice that received control odor. Transfer the mice to the housing location. 
	&#8203;NOTE: The above procedure can be translated to rat models within appropriately sized cages. Adjustments to the measurements suggested in Figure 1C may be required to allow for better access to the bottom of the home cage.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64264/64264fig01.jpg" /> 
Figure 1: Transponders and home cage temperature testing. (A) Diagram of unilateral transponder placement for testing temperature in a mouse gastrocnemius. Once programmed and placed, the transponder-reader (DAS-8027-IUS, shown) can be used to measure temperature. (B) Left, photo of an open mesh stainless steel tea ball and a 5 cm x 5 cm towel. Right, enclosed tea ball, used to hold habituation and odor towels in home cage testing. (C) Schematic of risers constructed with PVC piping for home cage testing. (D) Workflow of home cage testing protocol. (E) Facility images of home cage testing area. Left, four mouse cages atop a riser. Magnetic strips are located on the adjacent wall, and magnets and surgical cloth are on the table. Right, covered mouse cages on risers. (A), (C), and (D) were created with Biorender.com.&#160;<a href="https://www.jove.com/files/ftp_upload/64264/64264fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
7. Temperature testing - treadmill walking
<ol>
	<li>Assign each animal a treadmill as their assigned location for habituation and testing procedures.</li>
	<li>Prepare the treadmills for testing, ensuring that the shockers are functional. 
	NOTE: For treadmill walking, treadmills should be set at the lowest available pace that promotes continuous movement but not running for both habituation and testing. For the 1012M-2 Modular Enclosed Metabolic Treadmill, this is 5.2 m/min for mice and 7 m/min for rats. This pace may need to be adjusted based on the obesity of the subject.&#160;Shockers should be set to an intensity and repetition rate of 5.0.</li>
	<li>Habituation
	<ol>
		<li>Move the mice to the testing room. Allow mice 1-2 h to acclimate to the room transfer in their home cages.</li>
		<li>After acclimation, guide the mice to the opening of their assigned treadmill and close the treadmill. Start the belt, shocker, and stopwatch.</li>
		<li>Allow the mice to walk on the treadmills for 15 min, using shock stimulus as motivation for movement. Stop the test immediately if an animal remains on an active shocker for an extended period.</li>
		<li>After the test, remove the mice and return them to the home cages.</li>
		<li>Clean the treadmills using liquid detergent and water.</li>
	</ol>
	</li>
	<li>Testing
	<ol>
		<li>Move the mice to the testing room. Allow the mice 1-2 h to acclimate to the room transfer in their home cages.</li>
		<li>Measure and record baseline temperature prior to moving the mouse to the treadmill. 
		NOTE: For tests including pharmacological agents, apply or inject them here, following the schematic shown in Figure 2A. Wait time after injection can be added as needed before the mice are placed on the treadmill. Randomization should be employed when providing any stimuli.</li>
		<li>Place 5 cm x 5 cm squares of control or PO towels within the treadmill closest to the front of the treadmill. Adhere the towels to the ceiling of the treadmill or underneath for easy placement and removal.</li>
		<li>Guide the mice into the assigned treadmill. Turn on the treadmill belt and shocker.</li>
		<li>Start the stopwatch. Take measurements of the test subjects in the same order in which the mice were set up in the treadmills. Record the temperatures and clock time of the measurements following the desired time points. 
		NOTE: Temperature can reliably be measured from outside the treadmill while a mouse is inside an enclosed treadmill during walking activity. For rats, the treadmill size and transponder-reader&#160;distance limitations may require an experimenter to keep the back of the treadmill open to insert the reader inside the treadmill, closer to the subject.</li>
		<li>When the test is complete, turn off the shockers and treadmills; return the mice to their home cages. Transfer the mice to the housing location.</li>
		<li>Clean the treadmills using liquid detergent and water, paying specific attention to remove any residual PO.</li>
		<li>When experiments are complete, euthanize the animals (e.g., using CO2 inhalation), and visually confirm the transponder location.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64264/64264fig02.jpg" /> 
Figure 2: Activity-controlled temperature testing. (A) Workflow of activity-controlled temperature testing with a pharmacological agent using treadmill walking. (B) Facility images of treadmills. Left, an image of full equipment setup. Right, a closer image of individual treadmills and shockers. (A) was created with Biorender.com.&#160;<a href="https://www.jove.com/files/ftp_upload/64264/64264fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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K., Bartness, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+activation+of+the+sympathetic+innervation+of+adipose+tissues+by+melanocortin+receptor+stimulation.">Differential activation of the sympathetic innervation of adipose tissues by melanocortin receptor stimulation.</a> Endocrinology. 148 (11), 5339-5347 (2007).</li><li>Vaughan, C. H., Shrestha, Y. B., Bartness, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+a+novel+melanocortin+receptor-containing+node+in+the+SNS+outflow+circuitry+to+brown+adipose+tissue+involved+in+thermogenesis.">Characterization of a novel melanocortin receptor-containing node in the SNS outflow circuitry to brown adipose tissue involved in thermogenesis.</a> Brain Research. 1411, 17-27 (2011).</li><li>Kort, W. J., Hekking-Weijma, J. M., TenKate, M. T., Sorm, V., VanStrik, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microchip+implant+system+as+a+method+to+determine+body+temperature+of+terminally+ill+rats+and+mice.">A microchip implant system as a method to determine body temperature of terminally ill rats and mice.</a> Laboratory Animals. 32 (3), 260-269 (1998).</li><li>Mei, J., 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=Body+temperature+measurement+in+mice+during+acute+illness:+Implantable+temperature+transponder+versus+surface+infrared+thermometry.">Body temperature measurement in mice during acute illness: Implantable temperature transponder versus surface infrared thermometry.</a> Scientific Reports. 8, 3526 (2018).</li><li>Warn, P. 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=Infrared+body+temperature+measurement+of+mice+as+an+early+predictor+of+death+in+experimental+fungal+infections.">Infrared body temperature measurement of mice as an early predictor of death in experimental fungal infections.</a> Laboratory Animals. 37 (2), 126-131 (2003).</li><li>Hargreaves, K., Dubner, R., Brown, F., Flores, C., Joris, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+and+sensitive+method+for+measuring+thermal+nociception+in+cutaneous+hyperalgesia.">A new and sensitive method for measuring thermal nociception in cutaneous hyperalgesia.</a> Pain. 32 (1), 77-88 (1988).</li><li>Fiebig, K., Jourdan, T., Kock, M. H., Merle, R., Thone-Reineke, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+infrared+thermography+for+temperature+measurement+in+adult+male+NMRI+nude+mice.">Evaluation of infrared thermography for temperature measurement in adult male NMRI nude mice.</a> Journal of the American Association for Laboratory Animal Science. 57 (6), 715-724 (2018).</li><li>Franco, N. H., Geros, A., Oliveira, L., Olsson, I. A. S., Aguiar, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ThermoLabAnimal+-+A+high-throughput+analysis+software+for+non-invasive+thermal+assessment+of+laboratory+mice.">ThermoLabAnimal - A high-throughput analysis software for non-invasive thermal assessment of laboratory mice.</a> Physiology &amp; Behavior. 207, 113-121 (2019).</li><li>Koganti, S. R., 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=Disruption+of+KATP+channel+expression+in+skeletal+muscle+by+targeted+oligonucleotide+delivery+promotes+activity-linked+thermogenesis.">Disruption of KATP channel expression in skeletal muscle by targeted oligonucleotide delivery promotes activity-linked thermogenesis.</a> Molecular Therapy. 23 (4), 707-716 (2015).</li><li>Bal, N. C., Periasamy, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uncoupling+of+sarcoendoplasmic+reticulum+calcium+ATPase+pump+activity+by+sarcolipin+as+the+basis+for+muscle+non-shivering+thermogenesis.">Uncoupling of sarcoendoplasmic reticulum calcium ATPase pump activity by sarcolipin as the basis for muscle non-shivering thermogenesis.</a> Philosophical Transactions of the Royal Society B. 375 (1793), 20190135 (2020).</li><li>Hicks, C., 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=Body+temperature+and+cardiac+changes+induced+by+peripherally+administered+oxytocin,+vasopressin+and+the+non-peptide+oxytocin+receptor+agonist+WAY+267,464:+a+biotelemetry+study+in+rats.">Body temperature and cardiac changes induced by peripherally administered oxytocin, vasopressin and the non-peptide oxytocin receptor agonist WAY 267,464: a biotelemetry study in rats.</a> British Journal of Pharmacology. 171 (11), 2868-2887 (2014).</li><li>Kasahara, Y., 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=Oxytocin+receptor+in+the+hypothalamus+is+sufficient+to+rescue+normal+thermoregulatory+function+in+male+oxytocin+receptor+knockout+mice.">Oxytocin receptor in the hypothalamus is sufficient to rescue normal thermoregulatory function in male oxytocin receptor knockout mice.</a> Endocrinology. 154 (11), 4305-4315 (2013).</li><li>Kasahara, Y., 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=Role+of+the+oxytocin+receptor+expressed+in+the+rostral+medullary+raphe+in+thermoregulation+during+cold+conditions.">Role of the oxytocin receptor expressed in the rostral medullary raphe in thermoregulation during cold conditions.</a> Frontiers in Endocrinology. 6, 180 (2015).</li><li>Yuan, J., Zhang, R., Wu, R., Gu, Y., Lu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+oxytocin+to+rectify+metabolic+dysfunction+in+obese+mice+are+associated+with+increased+thermogenesis.">The effects of oxytocin to rectify metabolic dysfunction in obese mice are associated with increased thermogenesis.</a> Molecular and Cellular Endocrinology. 514, 110903 (2020).</li><li>Scholl, J. L., Afzal, A., Fox, L. C., Watt, M. J., Forster, G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sex+differences+in+anxiety-like+behaviors+in+rats.">Sex differences in anxiety-like behaviors in rats.</a> Physiology &amp; Behavior. 211, 112670 (2019).</li></ol>

The authors declare that they have no conflicts of interest.

This temperature testing protocol provides the field with an avenue to measure skeletal muscle thermogenesis directly. This is critical as research delves into identifying the mechanisms underlying muscle thermogenesis33. The method provides two cost-effective protocols for measuring skeletal muscle thermogenesis under contextual and pharmacological conditions. This protocol emphasizes the importance of both habituation and acclimation within these procedures. Habituation is used to repeatedly int...

Within metabolic research, the examination of skeletal muscle thermogenesis is a promising new avenue for probing body weight homeostasis. The published literature supports the idea that the thermogenic responses of one of the body&#39;s largest organ systems&#8212;the skeletal muscle&#8212;provide an avenue for increasing energy expenditure and other metabolic effects, thereby effectively rebalancing systems within diseases such as obesity1,2,3. If the muscle can be considered a thermogenic organ, studies must utilize a practical methodology to study thermogenic changes within this organ. The desire to understand the endothermic impact of skeletal muscles and the utility of this methodology for studying non-shivering muscle thermogenesis are not specific to metabolic studies. Disciplines including evolution4, comparative physiology5, and ecophysiology6,7 have shown a vested interest in understanding the ways in which muscle thermogenesis may contribute to endothermy and how this mechanism adapts to the environment. The presented protocol provides the critical methods necessary to address these questions.
The provided method can be utilized in the assessment of both contextual and pharmacological stimuli modulation of muscle temperature, including the unique technique of providing predator odor (PO) to shift the context to replicate predator threat. Prior reports have demonstrated the ability of PO to rapidly induce a sizable increase in muscle thermogenesis8. Moreover, pharmacological stimuli can also alter muscle temperature. This has been demonstrated in the context of PO-induced muscle thermogenesis, where pharmacological blockade of peripheral &#946;-adrenergic receptors, using nadolol, inhibited the ability of PO to induce muscle thermogenesis without significantly affecting contractile thermogenesis during treadmill walking8. Central administration of melanocortin receptor agonists in rats has also been used to discern brain mechanisms altering thermogenesis9,10.
Provided here is a preliminary investigation of the ability of the neurohormone oxytocin (Oxt) to alter muscle thermogenesis in mice. Similar to predator threat, social encounters with a same-sex conspecific increase body temperature, a phenomenon referred to as social hyperthermia11. Given the relevance of Oxt to social behavior12, it has been speculated that Oxt is a mediator of social hyperthermia in mice. Indeed, an oxytocin receptor antagonist decreases social hyperthermia in mice11, and mouse pups lacking Oxt show deficits in behavioral and physiological aspects of thermoregulation, including thermogenesis13. Given that Harshaw et al. (2021) did not find evidence supporting &#946;3 adrenergic receptor-dependent brown adipose tissue (BAT) thermogenesis with social hyperthermia11, it has been posited that social hyperthermia may be driven by Oxt&#39;s induction of muscle thermogenesis.
To measure skeletal muscle thermogenesis, the following protocol uses the implantation of preprogrammed IPTT-300 transponders adjacent to the muscle of interest within a mouse or rat8,10,14,15. These transponders are glass-encapsulated microchips that are read using corresponding transponder readers. Little to no research has utilized this technology in this capacity, though studies have suggested a need for the specificity provided by this method16,17. Previous investigations have shown the reliability of this method and a variety of ways in which temperature transponders can be used in comparison with other temperature-testing methods18 or in conjunction with surgical methods (e.g., cannulation19). However, studies of this nature rely on different strategic placements to measure overall body temperature20,21,22 or specified tissues such as BAT23,24,25.
Rather than measuring temperature from these locations or while using ear or rectal thermometers26, the method described here provides specificity for the muscle of interest. The ability to target a site by directly implanting transponders adjacent to the muscles of interest is more effective for probing muscle thermogenesis specifically. It provides a new avenue in addition to those provided by surface infrared thermometry27,28 or cutaneous temperature measurements via thermocouple29. Furthermore, the data provided through this method offer a range of avenues of research, avoiding the need for large, expensive, high-tech equipment and software such as infrared thermography30,31,32.
This method has been successfully used to measure temperature in the quadriceps and gastrocnemius, either unilaterally or bilaterally. This method has also been effective in conjunction with stereotaxic surgery14,15. Within ~7-10 cm&#160;of the transponder limb, portable transponder readers (DAS-8027/DAS-7007R) are used to scan, measure, and display the temperature. This distance has been critical and valuable to prior investigations8,9,10 because it minimizes potential stressors and temperature-altering variables such as animal handling during the testing procedures. Using timers, measurements can then be recorded and collected over a period of time without direct interaction with the animals.
To further minimize the disturbance of mice during testing, this method describes the assembly and use of risers made of PVC piping to give the experimenter access to the bottom of the home cages during testing. Using the risers in tandem with the digital reader, temperature measurements of the transponder limb can be made without any animal interaction after the stimulus is placed. At a minimal cost, this method can be used in conjunction with pharmacological and contextual stimuli, making it quite accessible for researchers. Additionally, this method can be employed with a substantial number of subjects (~16 mice or ~12 rats) at a time, saving time in increasing the overall throughput for any research project.
Introduced in this method is a&#160;crafted mechanism for presenting odors to mice using stainless steel mesh tea infuser balls, from now on referred to as &#34;tea balls&#34;. Though these tea balls are ideal for containing any odor material, in these studies, towels that served as in-cage bedding over 2-3 weeks for ferrets, a natural predator of mice and rats, are placed within each treatment tea ball. Each towel is cut into 5 cm x 5 cm squares. This aliquoting is also repeated with otherwise identical odorless control towels. Presenting these odors without a barrier (i.e., tea ball) led to mice shredding the fibers within their cages, increasing physical activity. This behavior was not as salient in rats. Tea balls provide a ventilated casing to the towel, giving full access to the odor while staying protected for the entirety of the experimental trial. These tea balls can be sanitized in accordance with animal use protocols, prepared, and introduced directly after surgery to begin habituating the animals to the structure along with the control stimulus. Mice can then live with the additional enrichment, decreasing the salience of the acute stimulus presentation.
Habituation to the presence of the tea ball is only one aspect of habituation that is critical to this method. The described habituation protocol also consists of repeated exposure to the testing procedure to normalize the testing environment (i.e., personnel, transportation and movement to the testing location, exposure to stimulus). This extended habituation minimizes nuanced responses from the animals and focuses measurements on the desired dependent variables (e.g., pharmacological or contextual stimuli). Previous assessment of this protocol has identified four trials as the minimum number of habituations necessary before temperature testing within home cages in rats8. If testing is separated by long periods (more than 2-3 weeks), the animals must be habituated again. For repeated habituation, a minimum of one to two trials are sufficient. However, if temperature tests are separated by more prolonged bouts of time, repeating more trials may be necessary.
In the continued effort to accustom mice and rats to the testing procedure, an acclimation period before stimulus presentation should be included in every experimental trial. This acclimation time is critical to rebalance temperature and activity after being shifted to the testing location. Rodents tend to have sharp temperature increases due to translocation. Acclimation should consist of a minimum of 1 h without interaction from the experimenter on the day of testing before any addition of a pharmacological agent or contextual stimuli. This is necessary each day of testing.
In the outlined home-cage temperature tests, mice have the free range of their home cage to roam in response to the tested stimulus. This can cause variable shifts in activity, impacting the accuracy of temperature readings and, therefore, the analysis of the thermogenic effects of the independent variable (e.g., pharmacological or contextual stimulus). In recognition of the potential changes in temperature due to activity level, a protocol is included below describing the use of temperature during treadmill walking. The published literature describes the successful use of this procedure in rats, and it is currently being employed with mice8,10,14,15. Treadmill walking maintains a constant speed of activity for the testing subject. For this study, treadmills are strictly used to control activity level and, therefore, are set to the lowest available speed on the treadmill to promote walking for mice and a similarly low setting for rats.
The following procedure is outlined for the temperature measurement of unilateral gastrocnemius in mice and predator odor presentation. The design can be used in conjunction with pharmacological agents and is transferrable to rats and other skeletal muscle groups (i.e., quadriceps) in mice. For rats, transponders can be placed in the gastrocnemius bilaterally and in brown adipose tissue. Due to size and distance limitations, only one transponder can be used per mouse. Minor modifications (e.g., the removal of contextual stimuli) can be made to assess thermogenic responses to pharmacological agents.

<table><tbody><tr><td>1012M-2 Modular Enclosed Metabolic Treadmill for Mice, 2 Lanes w/ Shock</td><td>Columbus Instruments</td><td></td><td></td></tr><tr><td>1012R-2 Modular Enclosed Metabolic Treadmill for Rats, 2 Lanes w/ Shock</td><td>Columbus Instruments</td><td></td><td></td></tr><tr><td>1-1/4 in. Ratcheting PVC Cutter</td><td>BrassCraft</td><td></td><td></td></tr><tr><td>1 mL Syringes</td><td>Fisher Scientific</td><td>BD 309659</td><td></td></tr><tr><td>Betadine Swabs</td><td>Fisher Scientific</td><td>19-898-945</td><td></td></tr><tr><td>Booster Coil</td><td>BioMedic Data Systems</td><td></td><td>Transponder Accessory</td></tr><tr><td>Electric Clippers</td><td>Andis</td><td></td><td>40 Ultraedge Clipper Blade</td></tr><tr><td>Flexible Mirror Sheets</td><td>Amazon</td><td></td><td>Self Adhesive Non Glass Mirror Tiles</td></tr><tr><td>Forceps</td><td>Fisher Scientific</td><td>89259-940</td><td></td></tr><tr><td>Heating Pad</td><td></td><td></td><td></td></tr><tr><td>Induction Chamber (isoflurane)</td><td>Kent Scientific</td><td>VetFlo-0730</td><td>3.0 L Low Cost Chambers for Traditional Vaporizers</td></tr><tr><td>Ketoprophen</td><td>Med-Vet Intl.</td><td>RXKETO-50</td><td></td></tr><tr><td>Magnetic Strips</td><td>Amazon</td><td></td><td></td></tr><tr><td>Magnets</td><td>Amazon</td><td></td><td>DIYMAG Magnetic Hooks 40lbs</td></tr><tr><td>Needles</td><td>Med-Vet Intl.</td><td>26400</td><td></td></tr><tr><td>Neomycin/Polymixin/Bacitracin with Hydrocortisone Ophthalmic Ointment, 3.5 g</td><td>Med-Vet Intl.</td><td>RXNPB-HC</td><td></td></tr><tr><td>Oasis Absorbable Suture</td><td>Med-Vet Intl.</td><td>MV-H821-V</td><td></td></tr><tr><td>Predator (Ferret) Odor Towels</td><td>Marshall BioResources</td><td></td><td></td></tr><tr><td>PVC pipe</td><td></td><td></td><td></td></tr><tr><td>Reflex Wound Clip Remover</td><td>CellPoint Scientific</td><td></td><td></td></tr><tr><td>Reflex Wound Clip, 7 mm (mouse)</td><td>CellPoint Scientific</td><td></td><td></td></tr><tr><td>Reflex Wound Clip, 9 mm (rat)</td><td>CellPoint Scientific</td><td></td><td></td></tr><tr><td>Srerile Autoclip, 7 mm (mouse)</td><td>CellPoint Scientific</td><td></td><td>Wound Clip Applier (mouse)</td></tr><tr><td>Stainless Strainers Interval Seasonings Tea Infuser</td><td>Amazon</td><td></td><td></td></tr><tr><td>Sterile Autoclip, 9 mm (rat)</td><td>CellPoint Scientific</td><td></td><td>Wound Clip Applier (rat)</td></tr><tr><td>Sterile Saline</td><td>Med-Vet Intl.</td><td>RX0.9NACL-10</td><td></td></tr><tr><td>Surgical Scissors</td><td>Fisher Scientific</td><td>08-951-5</td><td></td></tr><tr><td>Surgical Sheets</td><td></td><td></td><td></td></tr><tr><td>Towels (Control/Habituation)</td><td>Amazon</td><td></td><td>100% Cotton Towels, white</td></tr><tr><td>Transponders</td><td>BioMedic Data Systems</td><td></td><td>Model: IPTT-300</td></tr><tr><td>Transponders Reader</td><td>BioMedic Data Systems</td><td></td><td>Model: DAS-8027-IUS/ DAS-7007R</td></tr><tr><td>Versaclean</td><td>Fisher Scientific</td><td>18-200-700</td><td>liquid detergent</td></tr><tr><td>Webcol Alcohol Preps</td><td>Covidien</td><td>22-246-073&amp;nbsp;</td><td></td></tr><tr><td>Wedge pieces for PVC pipe</td><td></td><td></td><td></td></tr></tbody></table>

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.

Transponders were unilaterally implanted into the right gastrocnemius of ten 4-6-month-old, wild-type (WT) mice bred from the SF1-Cre strain (Tg(Nr5a1-cre)7Lowl/J, Strain #012462, C57BL/6J and FVB backgrounds; female N = 5; male N = 5). After recovery, the mice were habituated to a home cage temperature-testing procedure that did not include a contextual stimulus (e.g., PO). Temperature measurements using a transponder wand were recorded within their housing room and after transfer to the testing location. Mice were give...

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Measuring Skeletal Muscle Thermogenesis in Mice and Rats

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. ...

measuring-skeletal-muscle-thermogenesis-in-mice-and-rats

Research

JoVE Journal

Behavior

2.8K Views.  Kent State University. Dit protocol is belangrijk, omdat het een directe en kosteneffectieve methode biedt voor het meten van de temperatuur van de skeletspier. Bovendien kan het in verschillende contexten en onder stimulatie van farmacologische middelen worden gebruikt. Het grote voordeel van deze techniek is het aantal dieren dat tegelijkertijd getest kan worden.De minimale verstoring van dieren die hier wordt beschreven, vermijdt de noodzaak van hightech methoden zoals infraroodthermografie. Individuen ku...

Video: Measuring Skeletal Muscle Thermogenesis in Mice and Rats

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

Mouse Body Temperature Measurement Using Infrared Thermometer During Passive Systemic Anaphylaxis and Food Allergy Evaluation

Mouse body temperature measurement is of paramount importance for investigating allergies and anaphylactic symptoms. Rectal probes for temperature readings is common, and they have been proven to be accurate and invaluable in this regard. However, this method of temperature measurement requires the mice to be anesthetized in order to insert the probe without injury to the animal. This limits the ability to observe other phenotypes of the mouse simultaneously. In order to investigate other phenotypes while measuring temperatures, rectal probes are not ideal, and another method is desired. Here, we introduce a noninvasive method of temperature measurement that foregoes the requirement for mouse anesthesia while maintaining equal reliability to rectal probes in measuring body temperature. We use an infrared thermometer that detects body surface temperatures at ranges between 2 and 150 mm. This method of body temperature measurement is successful in reliably replicating temperature change trends during passive system anaphylaxis experiments in mice. We show that body surface temperatures are about 2.0 &#176;C lower than rectal probe measurements, but the degree of temperature drop follows the same trend. Furthermore, we use the same technique to observe mice in a food allergy model to evaluate temperature and activity levels simultaneously.

Here we present a new method to accurately measure body temperature differences in passive systemic anaphylaxis (PSA) and food allergy mouse models using an infrared thermometer. This procedure has been accurately duplicated in previous PSA results.

Mouse body temperature measurement is of paramount importance for investigating allergies and anaphylactic symptoms. Rectal probes for temperature ...

Immunology and Infection

Functional Imaging of Brown Fat in Mice with 18F-FDG micro-PET/CT

Brown adipose tissue (BAT) differs from white adipose tissue (WAT) by its discrete location and a brown-red color due to rich vascularization and high density of mitochondria. BAT plays a major role in energy expenditure and non-shivering thermogenesis in newborn mammals as well as the adults 1. BAT-mediated thermogenesis is highly regulated by the sympathetic nervous system, predominantly via &beta; adrenergic receptor 2, 3. Recent studies have shown that BAT activities in human adults are negatively correlated with body mass index (BMI) and other diabetic parameters 4-6. BAT has thus been proposed as a potential target for anti-obesity/anti-diabetes therapy focusing on modulation of energy balance 6-8. While several cold challenge-based positron emission tomography (PET) methods are established for detecting human BAT 9-13, there is essentially no standardized protocol for imaging and quantification of BAT in small animal models such as mice. Here we describe a robust PET/CT imaging method for functional assessment of BAT in mice. Briefly, adult C57BL/6J mice were cold treated under fasting conditions for a duration of 4 hours before they received one dose of 18F-Fluorodeoxyglucose (FDG). The mice were remained in the cold for one additional hour post FDG injection, and then scanned with a small animal-dedicated micro-PET/CT system. The acquired PET images were co-registered with the CT images for anatomical references and analyzed for FDG uptake in the interscapular BAT area to present BAT activity. This standardized cold-treatment and imaging protocol has been validated through testing BAT activities during pharmacological interventions, for example, the suppressed BAT activation by the treatment of &beta;-adrenoceptor antagonist propranolol 14, 15, or the enhanced BAT activation by &beta;3 agonist BRL37344 16. The method described here can be applied to screen for drugs/compounds that modulate BAT activity, or to identify genes/pathways that are involved in BAT development and regulation in various preclinical and basic studies.

Functional Imaging of Brown Fat in Mice with 18F-FDG micro-PET/CT

A method of functional imaging of mouse brown adipose tissue (BAT) is described in which cold-stimulated uptake of 18F-Fluorodeoxyglucose (FDG) in BAT is non-invasively assessed with a standardized micro-PET/CT protocol. This method is robust and sensitive to detect differences in BAT activities in mouse models.

Brown adipose tissue (BAT) differs from white adipose  tissue (WAT) by its discrete location and a brown-red color due to rich  vascularization and high ...

Biology

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 ...

Medicine

An Intra-Tissue Radiometry Microprobe for Measuring Radiance In Situ in Living Tissue

Organisms appear opaque largely because their outer tissue layers are strongly scattering to incident light; strongly absorbing pigments, such as blood, typically have narrow absorbances, such that the mean free path of light outside the absorbance peaks can be quite long. As people cannot see through tissue, they generally imagine that tissues like the brain, fat, and bone contain little or no light. However, photoresponsive opsin proteins are expressed within many of these tissues, and their functions are poorly understood. Radiance internal to tissue is also important for understanding photosynthesis. For example, giant clams are strongly absorbing yet maintain a dense population of algae deep in the tissue. Light propagation through systems like sediments and biofilms can be complex, and these communities can be major contributors to ecosystem productivity. Therefore, a method for constructing optical micro-probes for measuring scalar irradiance (photon flux intersecting a point) and downwelling irradiance (photon flux crossing a plane perpendicularly) to better understand these phenomena inside living tissue has been developed. This technique is also tractable in field laboratories. These micro-probes are made from heat-pulled optical fibers that are then secured in pulled glass pipettes. To change the angular acceptance of the probe, a 10-100 &#181;m sized sphere of UV-curable epoxy mixed with titanium dioxide is then secured to the end of a pulled, trimmed fiber. The probe is inserted into living tissue, and its position is controlled using a micromanipulator. These probes are capable of measuring in situ tissue radiance at spatial resolutions of 10-100 &#181;m or on the scale of single cells. These probes were used to characterize the light reaching the adipose and brain cells 4 mm below the skin of a living mouse and to characterize the light reaching similar depths within living algae-rich giant clam tissue.

An Intra-Tissue Radiometry Microprobe for Measuring Radiance In Situ in Living Tissue

In this paper, a method for measuring radiance in situ in living tissue is described. This work includes details of the construction of micro-scale probes for different measurements of radiance and irradiance, provides guidance for mounting tissue for the characterization of radiance, and outlines computational methods for analyzing the resulting data.

Organisms appear opaque largely because their outer tissue layers are strongly scattering to incident light; strongly absorbing pigments, such as blood, ...

Bioengineering

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 ...

Measurement of Insulin- and Contraction-Stimulated Glucose Uptake in Isolated and Incubated Mature Skeletal Muscle from Mice

Skeletal muscle is an insulin-responsive tissue and typically takes up most of the glucose that enters the blood after a meal. Moreover, it has been reported that skeletal muscle may increase the extraction of glucose from the blood by up to 50-fold during exercise compared to resting conditions. The increase in muscle glucose uptake during exercise and insulin stimulation is dependent on the translocation of glucose transporter 4 (GLUT4) from intracellular compartments to the muscle cell surface membrane, as well as phosphorylation of glucose to glucose-6-phosphate by hexokinase II. Isolation and incubation of mouse muscles such as m. soleus and m. extensor digitorum longus (EDL) is an appropriate ex vivo model to study the effects of insulin and electrically-induced contraction (a model for exercise) on glucose uptake in mature skeletal muscle. Thus, the ex vivo model permits evaluation of muscle insulin sensitivity and makes it possible to match muscle force production during contraction ensuring uniform recruitment of muscle fibers during measurements of muscle glucose uptake. Moreover, the described model is suitable for pharmacological compound testing that may have an impact on muscle insulin sensitivity or may be of help when trying to delineate the regulatory complexity of skeletal muscle glucose uptake.
Here we describe and provide a detailed protocol on how to measure insulin- and contraction-stimulated glucose uptake in isolated and incubated soleus and EDL muscle preparations from mice using radiolabeled [3H]2-deoxy-D-glucose and [14C]mannitol as an extracellular marker. This allows accurate assessment of glucose uptake in mature skeletal muscle in the absence of confounding factors that may interfere in the intact animal model. In addition, we provide information on metabolic viability of incubated mouse skeletal muscle suggesting that the method&#160;applied possesses some caveats under certain conditions when studying muscle energy metabolism.

Intact regulation of muscle glucose uptake is important for maintaining whole body glucose homeostasis. This protocol presents assessment of insulin- and contraction-stimulated glucose uptake in isolated and incubated mature skeletal muscle when delineating the impact of various physiological interventions on whole body glucose metabolism.

Skeletal muscle is an insulin-responsive tissue and typically takes up most of the glucose that enters the blood after a meal. Moreover, it has been ...

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 ...

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 ...

Measuring Skeletal Muscle Thermogenesis in Mice and Rats

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