O.A.M. is supported by NIH grants DK062876 and DK092759; D.P.B. is supported by the University of Michigan Medical Scientist Training Program (T32GM007863), University of Michigan Training Program in Organogenesis (T32HD007605), University of Michigan Rackham Merit Fellowship, and Tylenol Future Care Fellowship.

All animal procedures are performed with the approval of the Institutional Animal Care and Use Committee (IACUC) of the University of Michigan.
1. Euthanization
NOTE: For the purpose of this video protocol, 4 to 6 month-old C57BL/6J mice are used.
<ol>
	<li>Place the mouse in an isoflurane vaporizer chamber and adjust the isoflurane flow rate to 5% or greater. Continue isoflurane exposure until one minute after breathing stops. Then remove the mouse from the vaporizer chamber and confirm euthanasia using an approved secondary measure. 
	NOTE: Approved secondary measures will vary by institution and animal protocol and may include cervical dislocation or decapitation.</li>
	<li>Place the euthanized mouse on the dissection pan. Sterilize the dorsal and ventral external surfaces of the mouse using 70% ethanol. Ensure that the external surface of the mouse is sufficiently wet to minimize contamination from fur during dissection.</li>
</ol>
2. Identification and Isolation of Major Subcutaneous Adipose Ddepots (Anterior Subcutaneous, Dorsolumbar, Inguinal, Gluteal) and Interscapular Brown Aadipose Tissue
<ol>
	<li>Identifying and isolating anterior subcutaneous WAT 
	NOTE: Anterior subcutaneous WAT is located between the scapulae, descending from the nape of the neck to the axillae of the mouse7. This depot has alternatively been described as suprascapular8 or interscapular20 WAT and lies directly on top of the interscapular BAT depot.
	<ol>
		<li>To isolate the anterior subcutaneous depot, lay the mouse on its stomach in a prone position. Secure the upper and lower limbs to the dissection pan with dissection pins.</li>
		<li>Use forceps to lift the dorsal skin at the nape of the neck. Use iris scissors to make a small (1 mm) cut in the skin.</li>
		<li>Insert one blade of the iris scissors into the initial cut and make a midline vertical incision (2&#8211;3 cm) through the skin, beginning at the nape of the neck and descending along the spine to mid-back.</li>
		<li>Make two horizontal incisions (1 cm each) using the iris scissors, extending laterally from midline, at the top and bottom of the initial vertical incision.</li>
		<li>Use forceps to carefully peel back the skin and expose the anterior subcutaneous depot.</li>
		<li>Use iris scissors to remove the depot following the natural borders of the tissue. 
		NOTE: This method isolates both anterior subcutaneous WAT and interscapular BAT.</li>
		<li>Place the dissected depot on the dissection pan and carefully remove the contaminating BAT using iris scissors.</li>
	</ol>
	</li>
	<li>Identifying and isolating classical BAT 
	NOTE: Classical BAT is located beneath the anterior subcutaneous WAT depot.
	<ol>
		<li>To isolate BAT, use iris scissors to cut horizontally along the bottom edge of the anterior subcutaneous tissue, following the natural border of the depot.</li>
		<li>Then, use iris scissors to make two vertical incisions along the lateral edges of the depot, following the natural borders of the tissue.</li>
		<li>Use forceps to carefully flip the depot up and reveal the butterfly-shaped interscapular BAT embedded within the WAT. Carefully dissect the BAT out from the surrounding WAT.</li>
	</ol>
	</li>
	<li>Identifying and isolating posterior subcutaneous WAT,
	<ol>
		<li>Lay the mouse on its back in a supine position.</li>
		<li>After securing the upper and lower limbs with dissection pins, use forceps to lift up the skin at the base of the sternum and make a small (1 mm) cut in the skin.</li>
		<li>Insert one blade of the iris scissors into the initial cut and make a midline incision (4&#173;&#8211;5 cm) through the skin, beginning at the base of the sternum and descending to the base of the tail. Exercise caution when making this incision because the ventral skin is very thin and is closely associated with the underlying peritoneal cavity wall.</li>
		<li>Make two horizontal incisions (1 cm each) using iris scissors, extending laterally from midline, at the top of the initial vertical incision.</li>
		<li>Use forceps to carefully peel back the skin from the peritoneal cavity and the leg to find posterior subcutaneous WAT, which should remain associated with the skin. Secure the outstretched skin with dissection pins to ease complete excision of the WAT. 
		NOTE: Although the posterior subcutaneous WAT appears to be continuous, it is actually comprised of three discrete depots: dorsolumbar, inguinal, and gluteal1. The dorsolumbar depot extends from the lumbar spine to the base of the hindlimb. The triangular inguinal depot extends from the base of the hindlimb ventrally across the groin and contains a prominent lymph node. The gluteal depot extends inferiorly from the base of the groin and wraps around the leg to the base of the tail.</li>
	</ol>
	</li>
</ol>
3. Identification and Isolation of Visceral Adipose Depots (Gonadal, Perirenal, Retroperitoneal, Omental, Pericardial)
<ol>
	<li>Identifying and isolating visceral WAT depots 
	NOTE: WAT depots, which are largely contained within the thoracic and peritoneal cavities, keep the mouse in a supine position. Secure the upper and lower limbs to the dissection pan with dissection pins.
	<ol>
		<li>Use forceps to lift up the thin peritoneal cavity wall at the base of the sternum and make a small cut (1 mm) using iris scissors.</li>
		<li>Insert one blade of the iris scissors into the cut and make a descending vertical incision (4&#8211;5 cm) from the top of the peritoneal cavity (base of the sternum) to the rectum.</li>
		<li>Make two horizontal incisions (1 cm each) with iris scissors, extending laterally from midline, at the top and bottom of the vertical incision.</li>
		<li>Use forceps to peel back the peritoneum and expose the abdominal cavity contents. Pin the outstretched peritoneum to the dissection pan.</li>
	</ol>
	</li>
	<li>Identifying and isolating gonadal WAT 
	NOTE: Gonadal WAT surrounds the uterus and ovaries in females (ovarian or parametrial) and the epididymis and testes in males (epididymal). Ovarian WAT surrounds the ovaries, uterus, and bladder. In obese animals, gonadal and perirenal WAT can appear to be continuous &#8212; in this case, separate the depots at the uterine horn and ovaries. Epididymal WAT is found bound to the epididymis, testes, and the prominent epididymal blood vessel.
	<ol>
		<li>Locate the gonads (testes or ovaries) and use forceps to lift up the associated gonadal WAT.</li>
		<li>Use iris scissors to carefully excise the WAT from the gonads.</li>
	</ol>
	</li>
	<li>Identifying and isolating perirenal WAT 
	NOTE: Perirenal WAT surrounds the kidneys bilaterally. In obese animals, this depot can appear to extend inferiorly to the top of the uterine horn and ovaries. Although the perirenal WAT has traditionally been classified as a visceral depot21, several groups have identified it as a brown depot based on lineage tracing and radiolabeled glucose uptake studies8,9,20. Histologically it is comprised of a mixture of white and brown adipocytes.
	<ol>
		<li>To excise the perirenal depot, locate the kidney and use forceps to lift it up and pull it midline to see a clear division between the perirenal and retroperitoneal depots.</li>
		<li>Excise the WAT directly associated with the kidneys. Ensure that the adrenal glands, located above the kidneys, are removed from the WAT.</li>
	</ol>
	</li>
	<li>Identifying and isolating retroperitoneal WAT 
	NOTE: Retroperitoneal WAT is located in a paravertebral position, along the border between the posterior abdominal wall and the spinal cord.
	<ol>
		<li>To excise this depot, use forceps to lift the kidney up and towards midline to clearly see the border between the perirenal and retroperitoneal depots. 
		NOTE: In obese animals, identifying this border can be challenging.</li>
		<li>Then, use iris scissors to carefully dissect the retroperitoneal WAT from the posterior peritoneal wall.</li>
	</ol>
	</li>
	<li>Identifying and isolating omental WAT 
	NOTE: Omental WAT is located along the greater curvature of the stomach. Although omental adipose is an important depot in humans, it is generally present only in morbidly obese mice.
	<ol>
		<li>To identify visceral omental WAT, use forceps to lift the stomach up. Use iris scissors to remove the associated adipose tissue along the inferior border of the stomach. Do not confuse omental WAT with the pancreas.</li>
	</ol>
	</li>
	<li>Identifying and isolating mesenteric WAT 
	NOTE: Mesenteric WAT is a web-like structure surrounding and associated with the small and large intestines.
	<ol>
		<li>To excise this depot, remove the intestines from the rest of the digestive tract by cutting at the base of the stomach and the rectum. Use forceps to lift the intestines out of the visceral cavity and unravel them.</li>
		<li>Use iris scissors to carefully dissect the mesenteric WAT away from the intestines, beginning at the duodenum and continuing to the end of the colon. Carefully remove lymph nodes that are closely associated with the mesenteric depot.</li>
	</ol>
	</li>
	<li>Identifying and isolating pericardial WAT 
	NOTE: Pericardial WAT is located outside the visceral pericardium and on the external surface of the parietal pericardium, often along the inferior aspect of the heart14.
	<ol>
		<li>To gain access to the thoracic cavity, keep the mouse in a supine position. Secure the upper and lower limbs to the dissection pan with dissection pins.</li>
		<li>Use forceps to lift the xyphoid process (cartilage at the base of the sternum) and make a small (1 mm) cut in the thoracic cavity wall.</li>
		<li>Insert one blade of the iris scissors into the cut and make two horizontal incisions (1 cm each) extending laterally from the base of the sternum. This will expose the diaphragm and inferior border of the thoracic cavity.</li>
		<li>Use dissecting scissors to make two ascending vertical incisions (3&#8211;4 cm) through the ribcage, extending superiorly from the lateral borders of the thoracic cavity to the clavicle.</li>
		<li>Use forceps to lift up the ventral half of the ribcage. Use dissecting scissors to make a final cut (2 cm) along the inferior length of the clavicle to remove the ribcage and expose the thoracic cavity contents.</li>
		<li>If present, carefully dissect the pericardial adipose from the outer surface of the pericardium.</li>
	</ol>
	</li>
</ol>
4. Identification and Isolation of Other Adipose Depots
<ol>
	<li>Identifying and isolating epicardial WAT 
	NOTE: Epicardial WAT is contained within the visceral pericardium and is directly associated with the surface of the myocardium.
	<ol>
		<li>If present, epicardial adipocytes can be observed histologically after isolation and fixation of the perfused heart in 10% neutral buffered formalin for 24 h.</li>
	</ol>
	</li>
	<li>Identifying and isolating popliteal WAT 
	NOTE: Popliteal WAT is located in the popliteal fossa in the posterior knee and contains a large lymph node. This depot is not typically visible in young animals.
	<ol>
		<li>To isolate popliteal WAT, use iris scissors to carefully remove the skin from the base of the hind limb to the foot.</li>
		<li>Place the patella against the dissection pan and secure the outstretched leg with a pin in the foot. Ensure that the popliteal fossa at the back of the knee is facing upward.</li>
		<li>Use iris scissors to make a cut at the inferior border of the medial and lateral heads of the gastrocnemius muscle. Use forceps to lift up the muscle and reveal the triangular popliteal depot.</li>
		<li>Use iris scissors to excise the depot along the natural tissue borders.</li>
	</ol>
	</li>
	<li>Identifying and isolating dermal WAT 
	NOTE: Dermal WAT is a thin layer of adipocytes located between the reticular dermis and the panniculus carnosus muscle layer.
	<ol>
		<li>To identify dermal adipocytes by histological methods, place the mouse on its stomach in a prone position. After securing the upper and lower limbs with dissection pins, spray the external surface of the mouse with 70% ethanol to wet the skin.</li>
		<li>Use a scalpel to remove the wetted fur from a square portion of skin on the back of the mouse.</li>
		<li>Use iris scissors to carefully excise the shaved portion of skin, which will include the reticular dermis, dermal WAT, panniculus carnosus, and some subcutaneous WAT.</li>
		<li>Use a scalpel to cut the excised skin into thin vertical strips.</li>
		<li>Position the thin vertical strips with the sticky subcutaneous WAT layer facing down.</li>
		<li>Starting at one end, roll the strip onto itself to form a spiral. The sticky subcutaneous WAT layer on the outside of the spiral will allow the roll to maintain its shape during fixation.</li>
		<li>Place the spiral in a well of a 24 well plate containing 10% neutral buffered formalin for 24 h prior to histological processing22.</li>
	</ol>
	</li>
	<li>Identifying and isolating intermuscular WAT 
	NOTE: Intermuscular WAT is broadly defined as adipocytes located beneath the deep fascia of muscles. This term includes adipocytes interspersed between muscle fibers of skeletal muscle, also known as intramuscular WAT, and adipocytes located within muscle bundles themselves. Wildtype mice generally do not have large numbers of intermuscular adipocytes. However, it is possible under certain conditions to identify intermuscular WAT by histological methods. Under some conditions, intermuscular adipocytes can also be found in smooth muscles, such as the diaphragm.
	<ol>
		<li>To isolate the tibialis anterior (TA) muscle, for example, place the mouse on its back in a supine position and secure the lower limbs to the dissection pan using dissection pins. Wet the skin with 70% ethanol to minimize contamination with fur.</li>
		<li>Use iris scissors to carefully remove the skin from the leg and expose the quadriceps muscle group, located above the knee on the femur shaft, and the TA located below the knee on the ventral surface of the tibia. 
		NOTE: The TA is thick near the proximal end of the tibia and more tendinous near the distal end.</li>
		<li>Use iris scissors to excise a piece of the muscle and fix in 10% neutral buffered formalin for 24 h prior to histological processing22.</li>
	</ol>
	</li>
	<li>Identifying and isolating intra-articular WAT depots 
	NOTE: Intra-articular WAT depots are located within synovial joints.
	<ol>
		<li>To identify infrapatellar WAT, for example, by histological methods, harvest leg bones as detailed in the BMAT section.</li>
		<li>After the removing the femur-tibial complex, use iris scissors and gauze pads to remove as much muscle and connective tissue as possible. Do not break the tibiofemoral joint.</li>
		<li>Fix the femur-tibial complex in 10% neutral buffered formalin for 24 h prior to decalcification and histological processing (described below, step 4.6.8).</li>
	</ol>
	</li>
	<li>Identifying and isolating bone marrow adipose tissue 
	NOTE: Bone marrow adipose tissue (BMAT) is contained within bones, interspersed with hematopoietic cells. Anatomically, BMAT can be classified as constitutive (distal tibia and caudal vertebrae) or regulated (mid-to-proximal tibia, femur, and lumbar vertebrae)10,11. Clean isolation of bone marrow adipocytes for RNA and protein analyses is challenging in mice. However, mouse bones can readily be harvested, fixed, decalcified and paraffin-embedded for histological analyses.
	<ol>
		<li>To harvest leg bones, for example, first remove the legs from the mouse. Use dissection scissors to cut the acetabulofemoral joint, keeping the femoral head intact.</li>
		<li>Use iris scissors to carefully remove the skin from the leg, revealing the leg muscles.</li>
		<li>Carefully dissect away the main muscles from the femur and tibia using iris scissors and gauze pads.</li>
		<li>When the femur is exposed, follow the edge of the bone to the articulation of the pelvis and femur and carefully release the femoral head from the acetabulofemoral joint. Use gauze to clean any remaining tissue from the femur.</li>
		<li>Follow the border of the tibia to the ankle joint. Carefully release the medial malleolus, located at the tip of the tibia, from the ankle joint.</li>
		<li>Once the femur-tibial complex has been isolated, remove as much muscle and connective tissue as possible using iris scissors and/or gauze.</li>
		<li>Separate the tibia from the femur by inserting one blade of the iris scissors into the tibiofemoral joint and gently cut through the medial and lateral collateral ligaments and the anterior and posterior cruciate ligaments. Do not remove the capsule of the knee joint to ensure that the tibia remains intact. Use gauze to clean any remaining tissue from the tibia.</li>
		<li>Fix bones in 10% neutral buffered formalin for 24 h at room temperature and then wash and store according to future needs.
		<ol>
			<li>To analyze bone parameters using microcomputed tomography (&#181;CT), store bones in Sorensen&#8217;s phosphate buffer, pH 7.423.</li>
			<li>For BMAT quantification and histological analyses, decalcify bones in 14% EDTA, pH 7.4, for 10-14 days.</li>
			<li>Following decalcification, use osmium tetroxide staining and &#181;CT analysis to quantify BMAT24. Otherwise, process and paraffin-embed bones for histology23.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Cinti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+adipose+organ+at+a+glance.">The adipose organ at a glance.</a> Disease Models &amp; Mechanisms. 5 (5), 588-594 (2012).</li><li>Rosen, E. D., Spiegelman, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+we+talk+about+when+we+talk+about+fat.">What we talk about when we talk about fat.</a> Cell. 156 (1-2), 20-44 (2014).</li><li>Sanchez-Gurmaches, J., Guertin, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipocyte+lineages:+tracing+back+the+origins+of+fat.">Adipocyte lineages: tracing back the origins of fat.</a> Biochimica et Biophysica Acta. 1842 (3), 340-351 (2014).</li><li>Bagchi, D. P., Forss, I., Mandrup, S., MacDougald, O. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SnapShot:+Niche+Determines+Adipocyte+Character+I.">SnapShot: Niche Determines Adipocyte Character I.</a> Cell Metabolism. 27 (1), 264-264 (2018).</li><li>Tchkonia, T., 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=Mechanisms+and+metabolic+implications+of+regional+differences+among+fat+depots.">Mechanisms and metabolic implications of regional differences among fat depots.</a> Cell Metabolism. 17 (5), 644-656 (2013).</li><li>Kajimura, S., Spiegelman, B. M., Seale, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brown+and+Beige+Fat:+Physiological+Roles+beyond+Heat+Generation.">Brown and Beige Fat: Physiological Roles beyond Heat Generation.</a> Cell Metabolism. 22 (4), 546-559 (2015).</li><li>Frontini, A., Cinti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distribution+and+development+of+brown+adipocytes+in+the+murine+and+human+adipose+organ.">Distribution and development of brown adipocytes in the murine and human adipose organ.</a> Cell Metabolism. 11 (4), 253-256 (2010).</li><li>Zhang, F., 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=An+Adipose+Tissue+Atlas:+An+Image-Guided+Identification+of+Human-like+BAT+and+Beige+Depots+in+Rodents.">An Adipose Tissue Atlas: An Image-Guided Identification of Human-like BAT and Beige Depots in Rodents.</a> Cell Metabolism. 27, 252-262 (2018).</li><li>Sanchez-Gurmaches, J., Guertin, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipocytes+arise+from+multiple+lineages+that+are+heterogeneously+and+dynamically+distributed.">Adipocytes arise from multiple lineages that are heterogeneously and dynamically distributed.</a> Nature Communications. 5, 4099 (2014).</li><li>Li, Z., Hardij, J., Bagchi, D. P., Scheller, E. L., MacDougald, O. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development,+regulation,+metabolism+and+function+of+bone+marrow+adipose+tissues.">Development, regulation, metabolism and function of bone marrow adipose tissues.</a> Bone. 110, 134-140 (2018).</li><li>Scheller, E. L., Cawthorn, W. P., Burr, A. A., Horowitz, M. C., MacDougald, O. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Marrow+Adipose+Tissue:+Trimming+the+Fat.">Marrow Adipose Tissue: Trimming the Fat.</a> Trends in Endocrinology &amp; Metabolism. 27 (6), 392-403 (2016).</li><li>Alexander, C. 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=Dermal+white+adipose+tissue:+a+new+component+of+the+thermogenic+response.">Dermal white adipose tissue: a new component of the thermogenic response.</a> The Journal of Lipid Research. 56 (11), 2061-2069 (2015).</li><li>Kruglikov, I. L., Scherer, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dermal+Adipocytes:+From+Irrelevance+to+Metabolic+Targets?.">Dermal Adipocytes: From Irrelevance to Metabolic Targets?.</a> Trends in Endocrinology &amp; Metabolism. 27 (1), 1-10 (2016).</li><li>Iacobellis, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+and+systemic+effects+of+the+multifaceted+epicardial+adipose+tissue+depot.">Local and systemic effects of the multifaceted epicardial adipose tissue depot.</a> Nature Reviews Endocrinology. 11 (6), 363-371 (2015).</li><li>Addison, O., Marcus, R. L., LaStayo, P. C., Ryan, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intermuscular+Fat:+A+Review+of+the+Consequences+and+Causes.">Intermuscular Fat: A Review of the Consequences and Causes.</a> International Journal of Endocrinology. 2014, 1-11 (2014).</li><li>Pond, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose+tissue+and+the+immune+system.">Adipose tissue and the immune system.</a> Prostaglandins, Leukotrienes, and Essential Fatty Acids. 73 (1), 17-30 (2005).</li><li>Kloppenburg, A. I. -. F. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+emerging+player+in+knee+osteoarthritis:+the+infrapatellar+fat+pad.">An emerging player in knee osteoarthritis: the infrapatellar fat pad.</a> Arthritis Research &amp; Therapy. 15 (225), 1-9 (2013).</li><li>Mann, A., Thompson, A., Robbins, N., Blomkalns, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localization,+Identification,+and+Excision+of+Murine+Adipose+Depots.">Localization, Identification, and Excision of Murine Adipose Depots.</a> Journal of Visualized Experiments. (94), e52174 (2014).</li><li>Casteilla, L., Cousin, B., Calise, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Choosing+an+adipose+tissue+depot+for+sampling:+factors+in+selection+and+depot+specificity.">Choosing an adipose tissue depot for sampling: factors in selection and depot specificity.</a> Methods in Molecular Biology. 155, 1-22 (2008).</li><li>de Jong, J. M., Larsson, O., Cannon, B., Nedergaard, 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+stringent+validation+of+mouse+adipose+tissue+identity+markers.">A stringent validation of mouse adipose tissue identity markers.</a> American Journal of Physiology-Endocrinology and Metabolism. 308 (12), E1085-E1105 (2015).</li><li>Cinti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+adipose+organ.">The adipose organ.</a> Prostaglandins, Leukotrienes, and Essential Fatty Acids. 73 (1), 9-15 (2005).</li><li>Parlee, S. D., Lentz, S. I., Mori, H., MacDougald, O. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantifying+size+and+number+of+adipocytes+in+adipose+tissue.">Quantifying size and number of adipocytes in adipose tissue.</a> Methods in Enzymology. 537, 93-122 (2014).</li><li>Scheller, E. L., 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=Region-specific+variation+in+the+properties+of+skeletal+adipocytes+reveals+regulated+and+constitutive+marrow+adipose+tissues.">Region-specific variation in the properties of skeletal adipocytes reveals regulated and constitutive marrow adipose tissues.</a> Nature Communications. 6, 7808 (2015).</li><li>Scheller, E. L., 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=Use+of+osmium+tetroxide+staining+with+microcomputerized+tomography+to+visualize+and+quantify+bone+marrow+adipose+tissue+in+vivo.">Use of osmium tetroxide staining with microcomputerized tomography to visualize and quantify bone marrow adipose tissue in vivo.</a> Methods in Enzymology. 537, 123-139 (2014).</li><li>Lukjanenko, L., Brachat, S., Pierrel, E., Lach-Trifilieff, E., Feige, J. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genomic+profiling+reveals+that+transient+adipogenic+activation+is+a+hallmark+of+mouse+models+of+skeletal+muscle+regeneration.">Genomic profiling reveals that transient adipogenic activation is a hallmark of mouse models of skeletal muscle regeneration.</a> PLoS One. 8 (8), e71084 (2013).</li><li>Pagano, A. F., 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=Muscle+Regeneration+with+Intermuscular+Adipose+Tissue+(IMAT)+Accumulation+Is+Modulated+by+Mechanical+Constraints.">Muscle Regeneration with Intermuscular Adipose Tissue (IMAT) Accumulation Is Modulated by Mechanical Constraints.</a> PLoS One. 10 (12), e0144230 (2015).</li><li>Khan, I. 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=Intermuscular+and+perimuscular+fat+expansion+in+obesity+correlates+with+skeletal+muscle+T+cell+and+macrophage+infiltration+and+insulin+resistance.">Intermuscular and perimuscular fat expansion in obesity correlates with skeletal muscle T cell and macrophage infiltration and insulin resistance.</a> International Journal of Obesity. 39 (11), 1607-1618 (2015).</li><li>Sulston, R. 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=Increased+Circulating+Adiponectin+in+Response+to+Thiazolidinediones:+Investigating+the+Role+of+Bone+Marrow+Adipose+Tissue.">Increased Circulating Adiponectin in Response to Thiazolidinediones: Investigating the Role of Bone Marrow Adipose Tissue.</a> Frontiers in Endocrinology. 7, 128 (2016).</li></ol>

The authors have nothing to disclose.

As the importance of the diverse molecular and functional characteristics of discrete adipocyte clusters is increasingly recognized, it is crucial that investigators within the field uniformly identify and excise adipose depots for further analyses. To date, few protocols exist for standardized localization and isolation of the wide range of mouse adipose depots. Previously published methods are focused primarily on one or two depots and lack the details necessary for uniform identification and excision by different inve...

Adipose tissues play critical roles in whole-body homeostasis, including storage and release of energy in response to local and global needs, thermoregulation, and secretion of adipokines to regulate energy balance, metabolism and immune responses1,2. Adipocytes are distributed throughout the body in discrete depots, and in some cases serve specialized roles within their microenvironments3,4,5. Historically, the study of adipose tissue has centered on white adipose tissue (WAT), and its role in maintaining energy homeostasis. Most adipocytes are distributed throughout the body in subcutaneous and visceral WAT depots. The characteristics of these depots are important for differential susceptibility to metabolic diseases. Subcutaneous adipocytes, located beneath the skin, have been associated with protective metabolic effects5. Visceral adipocytes, which surround the vital organs and are contained within the gonadal, perirenal, retroperitoneal, omental and pericardial depots, are commonly linked to metabolic disorders, including type 2 diabetes and cardiovascular disease2. Brown adipose tissues (BAT) have also been studied extensively. Brown and brown-like adipocytes express uncoupling protein 1 (UCP1) and play important roles in adaptive thermogenesis and glucose homeostasis6,7. Classical brown adipocytes are contained in the interscapular BAT depot8. Clusters of brown adipocytes are also found in other locations, including supraclavicular, infra/subscapular, cervical, paravertebral and periaortic depots8,9.
In addition to their location in major WAT and BAT depots, adipocytes exist in discrete niches throughout the body4, where they may perform specialized functions within their respective microenvironments. For example, bone marrow adipose tissue (BMAT) serves as a lipid reservoir, is a major source of circulating adiponectin, and closely interacts with osteoblasts, osteoclasts, and hematopoietic cells10,11. Dermal adipocytes contribute to widespread processes, including wound healing, immune response, thermoregulation, and hair follicle growth12,13. Further, epicardial adipocytes may produce several adipokines and chemokines that exert local and systemic effects on the development and progression of coronary artery disease14. Expansion of inter/intramuscular WAT has been positively correlated with increased adiposity, systemic insulin resistance, and decreased muscular strength and mobility15. In addition, popliteal adipocytes serve as a lipid reservoir for lymphatic expansion during infection16. While the specific roles of different articular depots are generally unknown, the Hoffa depot (infrapatellar) within the knee is now thought to contribute to pathologies, including anterior knee pain and osteoarthritis17.
Whereas regional differences in adipocyte character and function are under intense study, the field is currently limited by the lack of a standardized protocol for the identification and dissection of diverse mouse depots. Previously published methods have typically described the isolation of one or two specific depots and lack the level of detail required for uniform excision18,19. The protocol described in this manuscript provides a comprehensive guide for the specific anatomic locations and isolation steps of many different mouse adipose depots. Although WAT depots are the primary focus of this manuscript, the excision of interscapular BAT is also described in detail. Adipose tissues excised using this protocol can be used for a wide variety of experimental endpoints, including&#160;explant studies, histology, and gene expression analyses.
The goal of this manuscript is to provide investigators with a detailed protocol to clearly and precisely identify and isolate both prominent and less-studied mouse adipose depots (Figure 1). This resource will facilitate a more complete investigation of the developmental, molecular, and functional characteristics of adipocytes within diverse niches.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59499/59499fig1.jpg" /> 
Figure 1: Schematic depiction of mouse adipose depots dissected in this protocol. This image has been adapted from Bagchi et al., 20184. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59499/59499fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table>
	<tbody>
		<tr>
			<td>10% neutral buffered formalin</td>
			<td>Fisher Scientific</td>
			<td>22-110-869</td>
			<td></td>
		</tr>
		<tr>
			<td>24-well plates, untreated</td>
			<td>Sigma-Aldrich</td>
			<td>CLS3738</td>
			<td></td>
		</tr>
		<tr>
			<td>70% ethanol (dilute from 95%)</td>
			<td>Fisher Scientific</td>
			<td>04-355-226</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting forceps with curved tips</td>
			<td>VWR</td>
			<td>89259-946</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting pan</td>
			<td>Carolina Biological Supply Company</td>
			<td>629004</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting scissors (sharp/blunt tip)</td>
			<td>VWR</td>
			<td>82027-588</td>
			<td></td>
		</tr>
		<tr>
			<td>Gauze sponges</td>
			<td>Vitality Medical</td>
			<td>2634</td>
			<td>Curity 4 x 4 inch gauze sponge, 12 ply</td>
		</tr>
		<tr>
			<td>Handi-Pins for dissection</td>
			<td>Carolina Biological Supply Company</td>
			<td>629132</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris scissors (straight)</td>
			<td>VWR</td>
			<td>470018-890</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>VetOne</td>
			<td>501017</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>VWR</td>
			<td>100499-578</td>
			<td>Feather scalpel handle with blade, disposable</td>
		</tr>
	</tbody>
</table>

identification and dissection of diverse mouse adipose depots

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

Successful identification and isolation of various mouse adipose depots can be achieved using the protocol described above. The gross anatomical locations of subcutaneous (A, E-F), brown (B), visceral (C, D, G-J), and popliteal (K) depots are shown in Figure 2.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59499/59499fig2.jpg" /> 
Figure 2: Gross anatomical locations of mou...

Watch this Scientific Journal Video about Identification and Dissection of Diverse Mouse Adipose Depots at JoVE.com

Identification and Dissection of Diverse Mouse Adipose Depots

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

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

identification-and-dissection-of-diverse-mouse-adipose-depots

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

39.7K Views.  University of Michigan Medical School. Adipocytes exist in discrete depots and have diverse roles within their unique microenvironments. As regional differences in adipocyte character and function are uncovered, standardized identification and isolation of depots is crucial for advancement of the field. Herein, we present a detailed protocol for the excision of various mouse adipose depots.

Video: Identification and Dissection of Diverse Mouse Adipose Depots

Dissection of 6.5 dpc Mouse Embryos

Analysis of gene expression patterns during early stages of mammalian embryonic development can provide important clues about gene function, cell-cell interaction and signaling mechanisms that guide embryonic patterning. However, dissection of the mouse embryo from the decidua shortly after implantation can be a challenging procedure, and detailed step-by-step documentation of this process is lacking.

Here we demonstrate how post-implantation (6.5 dpc) embryos are isolated by first dissecting the uterus of a pregnant mouse (detection of the vaginal plug was designated day 0.5 poist coitum) and subsequently dissecting the embryo from maternal decidua. The dissection of Reichert's membrane is described as well as the removal of the ectoplacental cone.

Isolation of postimplantation-stage embryos allows one to study gene patterning and analyze cell-lineage decision making processes during embryonic development, but proper dissection of the early embryo can be challenging. This protocol describes a method for isolating early primitive-streak-stage embryos (~6.5 days post coitum [dpc]).

Analysis of gene expression patterns during early stages of mammalian embryonic development can provide important clues about gene function, cell-cell ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its remarkable neuronal cell plasticity, and its involvement in epilepsy, neurodegenerative diseases, and psychiatric disorders. The hippocampus is composed of distinct regions; the dentate gyrus, which comprises mainly granule neurons, and Ammon's horn, which comprises mainly pyramidal neurons, and the two regions are connected by both anatomic and functional circuits. Many different mRNAs and proteins are selectively expressed in the dentate gyrus, and the dentate gyrus is a site of adult neurogenesis; that is, new neurons are continually generated in the adult dentate gyrus. To investigate mRNA and protein expression specific to the dentate gyrus, laser capture microdissection is often used. This method has some limitations, however, such as the need for special apparatuses and complicated handling procedures. In this video-recorded protocol, we demonstrate a dissection technique for removing the dentate gyrus from adult mouse under a stereomicroscope. Dentate gyrus samples prepared using this technique are suitable for any assay, including transcriptomic, proteomic, and cell biology analyses. We confirmed that the dissected tissue is dentate gyrus by conducting real-time PCR of dentate gyrus-specific genes, tryptophan 2,3-dioxygenase (TDO2) and desmoplakin (Dsp), and Ammon's horn enriched genes, Meis-related gene 1b (Mrg1b) and TYRO3 protein tyrosine kinase 3 (Tyro3). The mRNA expressions of TDO2 and Dsp in the dentate gyrus samples were detected at obviously higher levels, whereas Mrg1b and Tyro3 were lower levels, than those in the Ammon's horn samples. To demonstrate the advantage of this method, we performed DNA microarray analysis using samples of whole hippocampus and dentate gyrus. The mRNA expression of TDO2 and Dsp, which are expressed selectively in the dentate gyrus, in the whole hippocampus of alpha-CaMKII+/- mice, exhibited 0.037 and 0.10-fold changes compared to that of wild-type mice, respectively. In the isolated dentate gyrus, however, these expressions exhibited 0.011 and 0.021-fold changes compared to that of wild-type mice, demonstrating that gene expression changes in dentate gyrus can be detected with greater sensitivity. Taken together, this convenient and accurate dissection technique can be reliably used for studies focused on the dentate gyrus.

A dissection technique for removal of the dentate gyrus from adult mouse under a stereomicroscope was demonstrated in this video-recorded protocol.

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Identification and Analysis of Mouse Erythroid Progenitors using the CD71/TER119 Flow-cytometric Assay

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of red cell formation is regulated by the hormone erythropoietin (Epo), whose synthesis is triggered by tissue hypoxia. A threat to adequate tissue oxygenation results in a rapid increase in Epo, driving an increase in erythropoietic rate, a process known as the erythropoietic stress response. The resulting increase in the number of circulating red cells improves tissue oxygen delivery. An efficient erythropoietic stress response is therefore critical to the survival and recovery from physiological and pathological conditions such as high altitude, anemia, hemorrhage, chemotherapy or stem cell transplantation. 
The mouse is a key model for the study of erythropoiesis and its stress response. Mouse definitive (adult-type) erythropoiesis takes place in the fetal liver between embryonic days 12.5 and 15.5, in the neonatal spleen, and in adult spleen and bone marrow. Classical methods of identifying erythroid progenitors in tissue rely on the ability of these cells to give rise to red cell colonies when plated in Epo-containing semi-solid media. Their erythroid precursor progeny are identified based on morphological criteria. Neither of these classical methods allow access to large numbers of differentiation-stage-specific erythroid cells for molecular study. Here we present a flow-cytometric method of identifying and studying differentiation-stage-specific erythroid progenitors and precursors, directly in the context of freshly isolated mouse tissue. The assay relies on the cell-surface markers CD71, Ter119, and on the flow-cytometric 'forward-scatter' parameter, which is a function of cell size. The CD71/Ter119 assay can be used to study erythroid progenitors during their response to erythropoietic stress in vivo, for example, in anemic mice or mice housed in low oxygen conditions. It may also be used to study erythroid progenitors directly in the tissues of genetically modified adult mice or embryos, in order to assess the specific role of the modified molecular pathway in erythropoiesis.

A flow-cytometric method for identification and molecular analysis of differentiation-stage-specific murine erythroid progenitors and precursors, directly in freshly &#x2013;harvested mouse bone marrow, spleen or fetal liver. The assay relies on cell-surface markers CD71, Ter119, and cell size.

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of ...

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current blastocyst injection protocol. Here we report that a simple rotation of the embryo, and injection through Trans-Inner cell mass (TICM) increased the percentage of chimeric mice from 31% to 50%, with no additional equipment or further specialized training. 26 different inbred clones, and 35 total clones were injected over a period of 9 months. There was no significant difference in either pregnancy rate or recovery rate of embryos between traditional injection techniques and TICM. Therefore, without any major alteration in the injection process and a simple positioning of the blastocyst and injecting through the ICM, releasing the ES cells into the blastocoel cavity can potentially improve the quantity of chimeric production and subsequent germline transmission.

Here we present a protocol to increase chimera production without the use of new equipment. A simple orientation change of the embryo for injection can increase the number of embryos produced, and potentially reduce the timeline to germline transmission.

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current ...

Preparation of Adipose Progenitor Cells from Mouse Epididymal Adipose Tissues

Obesity and metabolic disorders such as diabetes, heart disease, and cancer, are all associated with dramatic adipose tissue remodeling. Tissue-resident adipose progenitor cells (APCs) play a key role in adipose tissue homeostasis and can contribute to the tissue pathology. The growing use of single cell analysis technologies &#8211; including single-cell RNA-sequencing and single-cell proteomics &#8211; is transforming the stem/progenitor cell field by permitting unprecedented resolution of individual cell expression changes within the context of population- or tissue-wide changes. In this article, we provide detailed protocols to dissect mouse epididymal adipose tissue, isolate single adipose tissue-derived cells, and perform fluorescence activated cell sorting (FACS) to enrich for viable Sca1+/CD31-/CD45-/Ter119- APCs. These protocols will allow investigators to prepare high quality APCs suitable for downstream analyses such as single cell RNA sequencing.

We present a simple method to isolate highly viable adipose progenitor cells from mouse epididymal fat pads using fluorescence activated cell sorting.

Obesity and metabolic disorders such as diabetes, heart disease, and cancer, are all associated with dramatic adipose tissue remodeling. Tissue-resident ...

Efficient Dissection and Culture of Primary Mouse Retinal Pigment Epithelial Cells

Eye disorders affect millions of people worldwide, but the limited availability of human tissues hinders their study. Mouse models are powerful tools to understand the pathophysiology of ocular diseases because of their similarities with human anatomy and physiology. Alterations in the retinal pigment epithelium (RPE), including changes in morphology and function, are common features shared by many ocular disorders. However, successful isolation and culture of primary mouse RPE cells is very challenging. This paper is an updated audiovisual version of the protocol previously published by Fernandez-Godino et al. in 2016 to efficiently isolate and culture primary mouse RPE cells. This method is highly reproducible and results in robust cultures of highly polarized and pigmented RPE monolayers that can be maintained for several weeks on Transwells. This model opens new avenues for the study of the molecular and cellular mechanisms underlying eye diseases. Moreover, it provides a platform to test therapeutic approaches that can be used to treat important eye diseases with unmet medical needs, including inherited retinal disorders and macular degenerations.

This protocol, which was originally reported by Fernandez-Godino et al. in 20161, describes a method to efficiently isolate and culture mouse RPE cells, which form a functional and polarized RPE monolayer within one week on Transwell plates. The procedure takes approximately 3 hours.

Eye disorders affect millions of people worldwide, but the limited availability of human tissues hinders their study. Mouse models are powerful tools to ...

Induction and Diverse Assessment Indicators of Experimental Autoimmune Encephalomyelitis

Multiple sclerosis (MS) is a typical autoimmune disease of the central nervous system (CNS) characterized by inflammatory infiltration, demyelination, and axonal damage. Currently, there are no measures to cure MS completely, but multiple disease-modifying therapies (DMT) are available to control and mitigate disease progression. There are significant similarities between the CNS pathological features of experimental autoimmune encephalomyelitis (EAE) and MS patients. EAE has been widely used as a representative model to determine MS drugs' efficacy and explore the development of new therapies for MS disease. Active induction of EAE in mice has a stable and reproducible effect and is particularly suitable for studying the effects of drugs or genes on autoimmune neuroinflammation. The method of immunizing C57BL/6J mice with myelin oligodendrocyte glycoprotein (MOG35-55) and the daily assessment of disease symptoms using a clinical scoring system is mainly shared. Given the complex etiology of MS with diverse clinical manifestations, the existing clinical scoring system can't satisfy the assessment of disease treatment. To avoid the shortcomings of a single intervention, new indicators to assess EAE based on clinical manifestations of anxiety-like moods and osteoporosis in MS patients are created to provide a more comprehensive assessment of MS treatment.

The present protocol describes the induction of experimental autoimmune encephalomyelitis in a mouse model using myelin oligodendrocyte glycoprotein and monitoring the disease process using a clinical scoring system. Experimental autoimmune encephalomyelitis-related symptoms are analyzed using mouse femur micro-computed tomography analysis and open field test to assess the disease process comprehensively.

Multiple sclerosis (MS) is a typical autoimmune disease of the central nervous system (CNS) characterized by inflammatory infiltration, demyelination, and ...

Isolation and Identification of Vascular Endothelial Cells from Distinct Adipose Depots for Downstream Applications

Vascular endothelial cells lining the wall of the vascular system play important roles in a variety of physiological processes, including vascular tone regulation, barrier functions, and angiogenesis. Endothelial cell dysfunction is a hallmark predictor and major driver for the progression of severe cardiovascular diseases, yet the underlying mechanisms remain poorly understood. The ability to isolate and perform analyses on endothelial cells from various vascular beds in their native form will give insight into the processes of cardiovascular disease. This protocol presents the procedure for the dissection of mouse subcutaneous and mesenteric adipose tissues, followed by isolation of their respective arterial vasculature. The isolated arteries are then digested using a specific cocktail of digestive enzymes focused on liberating functionally viable endothelial cells. The digested tissue is assessed by flow cytometry analysis using CD31+/CD45&#8722; cells as markers for positive endothelial cell identification. Cells can be sorted for immediate downstream functional assays or used to generate primary cell lines. The technique of isolating and digesting arteries from different vascular beds will provide options for researchers to evaluate freshly isolated vascular cells&#160;from arteries of interest and allow them to perform a wide range of functional tests on specific cell types.

This protocol details a method for the dissection of mouse adipose depots and the isolation and digestion of respective arteries to liberate and then identify the endothelial cell population. Freshly isolated cells used in downstream applications will advance the understanding of vascular cell biology and the mechanisms of vascular dysfunction.

Vascular endothelial cells lining the wall of the vascular system play important roles in a variety of physiological processes, including vascular tone ...

Identification and Dissection of Diverse Mouse Adipose Depots

Summary

Explore More Videos

Chapters in this video

Dissection of 6.5 dpc Mouse Embryos

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Dissection and Staining of Drosophila Larval Ovaries

Identification and Analysis of Mouse Erythroid Progenitors using the CD71/TER119 Flow-cytometric Assay

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

Preparation of Adipose Progenitor Cells from Mouse Epididymal Adipose Tissues

Efficient Dissection and Culture of Primary Mouse Retinal Pigment Epithelial Cells

Induction and Diverse Assessment Indicators of Experimental Autoimmune Encephalomyelitis

Isolation and Identification of Vascular Endothelial Cells from Distinct Adipose Depots for Downstream Applications