JSO is supported by an NIH Ruth L. Kirschstein NRSA (F32 DK091040), AK is supported by a Postdoctoral Fellowship from the American Diabetes Association (7-10-MI-05), and AHH is supported by an American Heart Association Established Investigator Award (12EIA8270000). Flow cytometry experiments were performed in the VMC Flow Cytometry Shared Resource. The VMC Flow Cytometry Shared Resource is supported by the Vanderbilt Digestive Disease Research Center (DK058404).

1. Reagents and Supplies
Prior to initiating this experimental protocol, prepare the following reagents:
<ol>
 <li>70% ethanol</li>
 <li>1X PBS</li>
 <li>1X DPBS (without Ca and Mg) supplemented with 0.5% BSA</li>
 <li>FACS buffer: 1X DPBS (without Ca and Mg), 2 mM EDTA, and 1% FCS</li>
 <li>ACK buffer: 150 mM NH4Cl, 10 mM KHCO3, and 0.1 mM Na2EDTA in water</li>
</ol>
2. Harvesting and Preparation of Adipose Tissue
<ol>
 <li>Euthanize mice according to IACUC-approved procedures specific to each institution.</li>
 <li>Thoroughly wet the fur with 70% ethanol.</li>
 <li>Make an incision at the level of the xiphoid process (lower part of the sternum) and open the thoracic cavity to expose the heart, taking care not to sever any major blood vessels.</li>
</ol>
Note: At this point, it is also helpful to leave the diaphragm intact as much as possible and to cut a notch out of the right side of the rib cage to allow blood and perfusate to flow out of the thoracic cavity.
<ol start="4">
 <li>Clip the right atrium to allow blood and perfusate to escape the circulatory system.</li>
</ol>
Note: When dealing with obese mice, excess pericardial AT may need to be removed to permit access to the heart.
<ol start="5">
 <li>Grasp the heart with forceps and gently insert a needle into the left ventricle through the apex. Slowly perfuse the mouse with 15 ml sterile PBS.</li>
</ol>
Note: Reduce the rate of perfusion if the lungs begin to fill and expand.
<ol start="6">
 <li>Open the peritoneal cavity and remove the perigonadal fat pads using care to avoid any gonadal tissues.</li>
 <li>Place fat pads in a weigh boat on ice containing 2 ml 1X DPBS (without Mg or Ca) supplemented with 0.5% BSA, and mince the AT into fine pieces.</li>
</ol>
Note: Limit the amount of AT per weigh boat to 1.2 g. If the amount of AT exceeds 1.2 g, divide it evenly between two weigh boats.
<ol start="8">
 <li>Keep AT samples on ice and prepare 3 ml collagenase digest solution per AT sample consisting of 1X DPBS supplemented with 0.5% BSA, 10 mM CaCl2, and 4 mg/ml collagenase, type II.</li>
</ol>
3. Collagenase Digestion
<ol>
 <li>Transfer AT to 50 ml conical tubes by pouring the homogenate and rinsing the weigh boat with 1 ml DPBS (0.5% BSA) and 3 ml collagenase II digest solution.</li>
 <li>Incubate AT homogenate in a rotational shaker (200 rpm) at 37 &deg;C for 20 min.</li>
 <li>Add 10 ml DPBS (0.5% BSA) to conical tubes and place on ice.</li>
 <li>Triturate homogenate numerous times using a 10 ml serological pipette, and pass cell suspensions through 100 &mu;m filter in to a new 50 ml conical tube.</li>
 <li>Centrifuge cell suspension at 500 x g for 10 min at 4 &deg;C.</li>
 <li>Decant supernatant and resuspend SVF cell pellet in 3 ml ACK buffer to lyse contaminating erythrocytes.</li>
 <li>Add 12 ml FACS buffer and centrifuge cell suspension at 500 x g for 10 min at 4 &deg;C.</li>
 <li>Decant supernatant and resuspend SVF cell pellet in FACS buffer.</li>
</ol>
Note: Use the size of the cell pellet as a guide for how much FACS buffer to resuspend in. If the cell pellet covers the bottom of the 50 ml conical tube, use 0.5-1 ml FACS buffer; otherwise, resuspend in 0.25-0.5 ml.
<ol start="9">
 <li>Place samples on ice and prepare 1:10 dilution aliquots of each sample for cell counting by mixing 40 &mu;l FACS buffer, 50 &mu;l trypan blue solution (0.2%), and 10 &mu;l cell suspension.</li>
 <li>Count viable cells based on trypan blue exclusion and dilute cell suspensions to a final concentration of 5-10 x 106 cells/ml.</li>
</ol>
4. Staining of Cell Surface Antigens
<ol>
 <li>Add anti-mouse CD16/CD32 antibody (Fc block) to a final concentration of 0.5-1 &mu;g/106 cells, and incubate on ice for 10 min.</li>
 <li>Transfer samples (&ge;106 cells) to 12 x 75 mm polystyrene round bottom tubes. Prepare separate tubes if analyzing ATMs and T cells, and combine extra cells to prepare an adequate number of tubes to accommodate the required compensation and modified fluorescence minus one (FMO) controls.</li>
</ol>
Note: As an example, when quantifying the proportion of ATMs based on F4/80 and CD11b, the following compensation and FMO controls will need to be prepared:
- Unstained (cells)
- DAPI or propidium iodide (PI) single stain (cells; do not add viability dye until step 5.1)
- F4/80 APC single stain (cells or compensation beads)
- CD11b FITC single stain (cells or compensation beads)
- FMO 1 (cells): Rat IgG2a &kappa; isotype control APC + CD11b FITC + viability dye (added at step 5.1)
- FMO 2 (cells): F4/80 APC + Rat IgG2b &kappa; isotype control FITC + viability dye (added at step 5.1)
<ol start="3">
 <li>Add fluorophore-conjugated primary antibodies and/or isotype controls at the appropriate concentration (see Table of reagents and materials).</li>
 <li>Protect samples from light and incubate at 4 &deg;C for 30 min.</li>
 <li>Add 2 ml FACS buffer and centrifuge cell suspension at 500 x g for 5 min at 4 &deg;C.</li>
 <li>Decant supernatant and resuspend SVF cell pellet in 2 ml FACS buffer.</li>
 <li>Centrifuge cell suspension 500 x g for 5 min at 4 &deg;C, and resuspend SVF cell pellet in &ge;400 &mu;l FACS buffer.</li>
 <li>Transfer samples to 12 x 75 mm polystyrene round bottom tubes equipped with a 35 &mu;m cell strainer tube tops.</li>
 <li>Protect from light, and store samples at 4 &deg;C until FACS analysis.</li>
</ol>
Note: For optimum results cells should be analyzed immeadiately; however, FACS analysis with this protocol has been successfully conducted on labeled cells stored in FACS buffer for 1-2 hr at 4 &deg;C. If cells need to be fixed to increase storage time, labeled cells can be fixed with 2% paraformaldehyde at 4 &deg;C for 24 hr prior to FACS analysis. Antibody companies suggest that labeled cells can be stored for up to one week; however, this has not been tested within the context of this procedure.
5. FACS Analysis
<ol>
 <li>Prior to FACS analysis, add viability dye to samples and appropriate controls to allow for live/dead cell discrimination.</li>
</ol>
Note: Numerous viability dyes are commercially available, but DAPI and PI are recommended depending on the excitation/emission profiles of the fluorophore-conjugated antibodies being used. DAPI and Propidium iodide are added to each sample at a final concentration of 0.2 mg/ml.
<ol start="2">
 <li>Use an unstained negative control sample, preferably cells isolated from the same tissue as the experimental samples, to adjust side scatter (SSC) and forward scatter (FSC) so that the cell population(s) of interest are on scale.</li>
</ol>
Note: For the analysis of AT SVF cells, it is recommended that SSC be displayed in a log scale versus FSC in a linear scale. The use of a log scale for SSC is especially important when analyzing ATMs, which are often very large and granular.
<ol start="3">
 <li>Draw an initial light scatter gate based on the type of cell(s) being analyzed, and adjust the photomultiplier tube (PMT) gain so that the unstained cells are on the far left of a single-parameter histogram (approximately centered on 102) for the appropriate channels.</li>
</ol>
Note: It is recommended that lymphocytes and macrophages (or other myeloid cells) be analyzed separately due to differences in autofluorescence.
<ol start="4">
 <li>Use single stained controls or antibody capture compensation beads to perform multi-color compensation.</li>
</ol>
Note: The use of compensation beads is recommended; however, compatibility of each antibody must be ensured. For example, compensation beads may not cross-react with rabbit antibodies, in which case isolated cells are required to obtain an appropriate single stain control.
<ol start="5">
 <li>Set experimental gates based on modified FMO controls.</li>
 <li>Set the flow cytometer to collect the appropriate number of events based on the prevalence of the population of interest and record experimental data.</li>
 <li>Export FCS data files for offline analysis. There are numerous programs available for flow cytometry data analysis. We recommend Cytobank, a web-based platform that allows investigatores to store and analyze data and generate figures from any computer with internet access 18. Additionally, Cytobank offers the ability to make data public accessible or restrict access to collaborators.</li>
</ol>

<ol>
	<li>Rodbell, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metabolism+of+Isolated+Fat+Cells.+I.+Effects+of+Hormones+on+Glucose+Metabolism+and+Lipolysis.">Metabolism of Isolated Fat Cells. I. Effects of Hormones on Glucose Metabolism and Lipolysis.</a> The Journal of Biological Chemistry. 239, 375-380 (1964).</li><li>Danse, L. H., Verschuren, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fish+oil-induced+yellow+fat+disease+in+rats.+III.+Lipolysis+in+affected+adipose+tissue.">Fish oil-induced yellow fat disease in rats. III. Lipolysis in affected adipose tissue.</a> Veterinary Pathology. 15, 544-548 (1978).</li><li>Weisberg, S. P., 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=Obesity+is+associated+with+macrophage+accumulation+in+adipose+tissue.">Obesity is associated with macrophage accumulation in adipose tissue.</a> The Journal of Clinical Investigation. 112, 1796-1808 (2003).</li><li>Xu, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+inflammation+in+fat+plays+a+crucial+role+in+the+development+of+obesity-related+insulin+resistance.">Chronic inflammation in fat plays a crucial role in the development of obesity-related insulin resistance.</a> The Journal of Clinical Investigation. 112, 1821-1830 (2003).</li><li>Mathis, D., Shoelson, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunometabolism:+an+emerging+frontier.">Immunometabolism: an emerging frontier.</a> Nature Reviews. Immunology. 11, 81 (2011).</li><li>Stefanovic-Racic, 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=Dendritic+cells+promote+macrophage+infiltration+and+comprise+a+substantial+proportion+of+obesity-associated+increases+in+CD11c++cells+in+adipose+tissue+and+liver.">Dendritic cells promote macrophage infiltration and comprise a substantial proportion of obesity-associated increases in CD11c+ cells in adipose tissue and liver.</a> Diabetes. 61, 2330-2339 (2012).</li><li>Liu, 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=Genetic+deficiency+and+pharmacological+stabilization+of+mast+cells+reduce+diet-induced+obesity+and+diabetes+in+mice.">Genetic deficiency and pharmacological stabilization of mast cells reduce diet-induced obesity and diabetes in mice.</a> Nature Medicine. 15, 940-945 (2009).</li><li>Feuerer, 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=Lean,+but+not+obese,+fat+is+enriched+for+a+unique+population+of+regulatory+T+cells+that+affect+metabolic+parameters.">Lean, but not obese, fat is enriched for a unique population of regulatory T cells that affect metabolic parameters.</a> Nature Medicine. 15, 930-939 (2009).</li><li>Nishimura, S., 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=CD8++effector+T+cells+contribute+to+macrophage+recruitment+and+adipose+tissue+inflammation+in+obesity.">CD8+ effector T cells contribute to macrophage recruitment and adipose tissue inflammation in obesity.</a> Nature Medicine. 15, 914-920 (2009).</li><li>Winer, S., 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=Normalization+of+obesity-associated+insulin+resistance+through+immunotherapy.">Normalization of obesity-associated insulin resistance through immunotherapy.</a> Nature Medicine. 15, 921-929 (2009).</li><li>Winer, D. 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=B+cells+promote+insulin+resistance+through+modulation+of+T+cells+and+production+of+pathogenic+IgG+antibodies.">B cells promote insulin resistance through modulation of T cells and production of pathogenic IgG antibodies.</a> Nature Medicine. 17, 610-617 (2011).</li><li>Wu, 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=Activation+of+invariant+natural+killer+T+cells+by+lipid+excess+promotes+tissue+inflammation,+insulin+resistance,+and+hepatic+steatosis+in+obese+mice.">Activation of invariant natural killer T cells by lipid excess promotes tissue inflammation, insulin resistance, and hepatic steatosis in obese mice.</a> Proceedings of the National Academy of Sciences of the United States of America. 109. , 1143-1152 (2012).</li><li>Wu, D., 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=Eosinophils+sustain+adipose+alternatively+activated+macrophages+associated+with+glucose+homeostasis.">Eosinophils sustain adipose alternatively activated macrophages associated with glucose homeostasis.</a> Science. 332, 243-247 (2011).</li><li>Elgazar-Carmon, V., Rudich, A., Hadad, N., Levy, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophils+transiently+infiltrate+intra-abdominal+fat+early+in+the+course+of+high-fat+feeding.">Neutrophils transiently infiltrate intra-abdominal fat early in the course of high-fat feeding.</a> Journal of Lipid Research. 49, 1894-1903 (2008).</li><li>Talukdar, S., 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=Neutrophils+mediate+insulin+resistance+in+mice+fed+a+high-fat+diet+through+secreted+elastase.">Neutrophils mediate insulin resistance in mice fed a high-fat diet through secreted elastase.</a> Nature Medicine. 18, 1407-1412 (2012).</li><li>Hagman, D. K., 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=Characterizing+and+quantifying+leukocyte+populations+in+human+adipose+tissue:+impact+of+enzymatic+tissue+processing.">Characterizing and quantifying leukocyte populations in human adipose tissue: impact of enzymatic tissue processing.</a> Journal of Immunological Methods. 386, 50-59 (2012).</li><li>Basu, S., Campbell, H. M., Dittel, B. N., Ray, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Purification+of+specific+cell+population+by+fluorescence+activated+cell+sorting+(FACS).">Purification of specific cell population by fluorescence activated cell sorting (FACS).</a> J. Vis. Exp. (41), e1546 (2010).</li><li>Kotecha, N., Krutzik, P. O., Irish, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Web-based+analysis+and+publication+of+flow+cytometry+experiments.">Web-based analysis and publication of flow cytometry experiments.</a> Current Protocols in Cytometry. Chapter 10, Unit 10 (2010).</li><li>Cho, C. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Angiogenic+role+of+LYVE-1-positive+macrophages+in+adipose+tissue.">Angiogenic role of LYVE-1-positive macrophages in adipose tissue.</a> Circulation Research. 100, e47-e57 (2007).</li><li>Roederer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spectral+compensation+for+flow+cytometry:+visualization+artifacts,+limitations,+and+caveats.">Spectral compensation for flow cytometry: visualization artifacts, limitations, and caveats.</a> Cytometry. 45, 194-205 (2001).</li></ol>

We have nothing to disclose.

Increasing interest in the role of the immune system in the metabolic consequences of obesity has led to the widespread use of flow cytometry to characterize immune cells of the AT. Although the exact protocol will vary between laboratories based on their own experience and available equipment, the critical steps include collagenase digestion, differential centrifugation, and cell surface antigen labeling. The goal of the present article is to provide a detailed protocol and practical guide for the isolation of the AT SV...

Historically, the adipose tissue (AT) has been viewed as an inert organ of lipid storage, which expands and contracts in response to energy balance. We now understand that AT represents a dynamic endocrine organ that actively secretes a number of hormones, which directly influence feeding behavior and systemic glucose homeostasis. In addition, over the past decade there has been an increasing appreciation for the numerous populations of immune cells residing in the AT stromal vascular fraction (SVF), as well as their contribution to AT homeostasis.	
 The ability to separate the AT adipocyte and SVF using a collagenase digest followed by differential centrifugation was first described by Rodbell in 1964 1. Collagenase II is most often used for adipocyte and SVF separation due to maintenance of adipocyte insulin receptors1. Early on, enzymatic fractionation of AT was primarily employed to study adipocyte metabolism and to isolate preadipocytes. More recently, this technique, combined with the widespread availability of flow cytometers and the ever-increasing number of commercially available fluorophore-conjugated antibodies, has facilitated the characterization of AT immune cells.	
 Although the presence of immune cells in inflamed AT had been described previously 2, the seminal papers by Weisberg et al. and Xu et al. published in 2003 were the first to document the accumulation of AT macrophages (ATMs) in obesity, which secrete inflammatory cytokines and correlate with AT-specific and systemic insulin resistance 3,4. These observations served as the basis of a new field of investigation recently coined, "immunometabolism," 5 and have been followed up by studies implicating various immune cell populations, including dendritic cells 6, mast cells 7, T cells 8-10, B cells 11, NKT cells 12, eosinophils 13, and neutrophils 14,15 in the development of obesity associated insulin resistance.	The goal of this article is to provide a detailed description of the collagenase digest technique used to isolate cells of the AT SVF and to characterize ATMs and AT T cells via flow cytometry. This protocol has been optimized for mouse AT; however, viewers may benefit from reading an excellent article providing extensive detail on optimization of this technique for human AT 16. The target audience of this article includes investigators with limited experience working with mouse AT and performing flow cytometry. Several practical considerations for balancing cellular yield and viability with time and resources are presented as well as optimal flow cytometry controls for characterizing AT immune cell populations. In addition to our protocol, readers are referred to a recent JoVE article by Basu et al. for an excellent discussion of some of the technical aspects of flow cytometry to include proper controls and compensations 17.

<table border="1">
 <tbody>
 <tr>
 <td>Name</td>
 <td>Company</td>
 <td>Catalog #</td>
 <td></td>
 </tr>
 <tr>
 <td>DPBS (no Ca or Mg) 10 x 500 ml</td>
 <td>Life Technologies</td>
 <td>14190-250</td>
 <td></td>
 </tr>
 <tr>
 <td>DAPI</td>
 <td>Life Technologies</td>
 <td>D3571</td>
 <td></td>
 </tr>
 <tr>
 <td>BSA</td>
 <td>Sigma</td>
 <td>A2153-100G</td>
 <td></td>
 </tr>
 <tr>
 <td>collagenase</td>
 <td>Sigma</td>
 <td>C6885-5G</td>
 <td></td>
 </tr>
 <tr>
 <td>propidium iodide solution</td>
 <td>Sigma</td>
 <td>P4864-10 ml</td>
 <td></td>
 </tr>
 <tr>
 <td>stable stack 20 microliter</td>
 <td>Rainin</td>
 <td>SS-L10</td>
 <td></td>
 </tr>
 <tr>
 <td>20 microliter filter tips</td>
 <td>Rainin</td>
 <td>SRL10F</td>
 <td></td>
 </tr>
 <tr>
 <td>stable stack 250 microliter tips</td>
 <td>Rainin</td>
 <td>SS-L250</td>
 <td></td>
 </tr>
 <tr>
 <td>1000 microliter tips</td>
 <td>Rainin</td>
 <td>GPS-L1000</td>
 <td></td>
 </tr>
 <tr>
 <td>1000 microliter filter tips</td>
 <td>Rainin</td>
 <td>GP-L1000F</td>
 <td></td>
 </tr>
 <tr>
 <td>250 microliter Filter Tips</td>
 <td>Rainin</td>
 <td>SR-L200F</td>
 <td></td>
 </tr>
 <tr>
 <td>FC Block</td>
 <td>BD Biosciences</td>
 <td>553142</td>
 <td></td>
 </tr>
 <tr>
 <td>fisher 100mn strainers</td>
 <td>Fisherbrand</td>
 <td>22-363-549</td>
 <td></td>
 </tr>
 <tr>
 <td>medium weigh dish</td>
 <td>Fisherbrand</td>
 <td>02-202B</td>
 <td></td>
 </tr>
 <tr>
 <td>aluminum foil</td>
 <td>Fisherbrand</td>
 <td>1213100</td>
 <td></td>
 </tr>
 <tr>
 <td>mincing scissors</td>
 <td>Fisherbrand</td>
 <td>089531B</td>
 <td></td>
 </tr>
 <tr>
 <td>Vortex</td>
 <td>Fisherbrand</td>
 <td>2215365</td>
 <td></td>
 </tr>
 <tr>
 <td>50 ml conical tubes</td>
 <td>BD Falcon</td>
 <td>14-959-49A</td>
 <td></td>
 </tr>
 <tr>
 <td>filter top FACS tubes</td>
 <td>BD Falcon</td>
 <td>352235</td>
 <td></td>
 </tr>
 <tr>
 <td>10 ml pipette case 200</td>
 <td>BD Falcon</td>
 <td>1367520</td>
 <td></td>
 </tr>
 <tr>
 <td>round bottom tubes</td>
 <td>BD Falcon</td>
 <td>352058</td>
 <td></td>
 </tr>
 <tr>
 <td>5 ml syringe</td>
 <td>BD Falcon</td>
 <td>309646</td>
 <td></td>
 </tr>
 <tr>
 <td>V Bottom Plates</td>
 <td>Costar</td>
 <td>07-200-107</td>
 <td></td>
 </tr>
 <tr>
 <td>transfer bulb pipette</td>
 <td>Thermo Scientific</td>
 <td>13-711-22</td>
 <td></td>
 </tr>
 <tr>
 <td>Shaker</td>
 <td>Thermo Scientific</td>
 <td>11 676 071</td>
 <td></td>
 </tr>
 <tr>
 <td>Adhesive mat</td>
 <td>Thermo Scientific</td>
 <td>1368750</td>
 <td></td>
 </tr>
 <tr>
 <td>Cell Culture Centrifuge</td>
 <td>Sorvall</td>
 <td>75253839</td>
 <td></td>
 </tr>
 <tr>
 <td>Adapters</td>
 <td>Sorvall</td>
 <td>75003723</td>
 <td></td>
 </tr>
 <tr>
 <td>Rat anti-mouse CD16/CD32</td>
 <td>BD Biosciences</td>
 <td>553142</td>
 <td>Concentration: 0.5 - 1 &mu;g/ 106 cells</td>
 </tr>
 <tr>
 <td>Rat anti-mouse F4/80</td>
 <td>eBioscience</td>
 <td>17-4801</td>
 <td>Fluorophore conjugate: APC 
 Concentration: 0.2 &mu;g/ 106 cells 
 Isotype control catalog number: 17-4321</td>
 </tr>
 <tr>
 <td>Rat anti-mouse CD11b</td>
 <td>eBioscience</td>
 <td>11-0112</td>
 <td>Fluorophore conjugate: FITC 
 Concentration: 0.5 &mu;g/ 106 cells 
 Isotype control catalog number: 11/1/4031</td>
 </tr>
 <tr>
 <td>Armenian Hamster anti-mouse CD11c</td>
 <td>eBioscience</td>
 <td>12-0114</td>
 <td>Fluorophore conjugate: PE 
 Concentration: 0.8 &mu;g/ 106 cells 
 Isotype control catalog number: Dec-88</td>
 </tr>
 <tr>
 <td>Goat anti-mouse MGL1/2</td>
 <td>R&amp;D Systems</td>
 <td>FAB4297P</td>
 <td>Fluorophore conjugate: PE 
 Concentration: 0.1 &mu;g/ 106 cells 
 Isotype control catalog number: IC108P</td>
 </tr>
 <tr>
 <td>Goat anti-mouse CD206</td>
 <td>R&amp;D Systems</td>
 <td>FAB2535P</td>
 <td>Fluorophore conjugate: PE 
 Concentration: 0.1 &mu;g/ 106 cells 
 Isotype control catalog number: IC108P</td>
 </tr>
 <tr>
 <td>Rat anti-mouse CCR2</td>
 <td>R&amp;D Systems</td>
 <td>FAB5538P</td>
 <td>Fluorophore conjugate: PE 
 Concentration: 0.1 &mu;g/ 106 cells 
 Isotype control catalog number: IC013P</td>
 </tr>
 <tr>
 <td>Armenian Hamster anti-mouse TCR&beta;</td>
 <td>BD Biosciences</td>
 <td>553174</td>
 <td>Fluorophore conjugate: APC 
 Concentration: 0.2 &mu;g/ 106 cells 
 Isotype control catalog number: 553956</td>
 </tr>
 <tr>
 <td>Rat anti-mouse CD8a</td>
 <td>BD Biosciences</td>
 <td>552877</td>
 <td>Fluorophore conjugate: PE-Cy7 
 Concentration: 0.8 &mu;g/ 106 cells 
 Isotype control catalog number: 552784</td>
 </tr>
 <tr>
 <td>Rat anti-mouse CD4</td>
 <td>BD Biosciences</td>
 <td>557956</td>
 <td>Fluorophore conjugate: Alexa Fluor 700 
 Concentration: 0.2 &mu;g/ 106 cells 
 Isotype control catalog number: 557963</td>
 </tr>
 </tbody>
</table>
Table 1. List of Materials and Reagents.

isolation of adipose tissue immune cells

The discovery of increased macrophage infiltration in the adipose tissue (AT) of obese rodents and humans has led to an intensification of interest in immune cell contribution to local and systemic insulin resistance. Isolation and quantification of different immune cell populations in lean and obese AT is now a commonly utilized technique in immunometabolism laboratories; yet extreme care must be taken both in stromal vascular cell isolation and in the flow cytometry analysis so that the data obtained is reliable and interpretable. In this video we demonstrate how to mince, digest, and isolate the immune cell-enriched stromal vascular fraction. Subsequently, we show how to antibody label macrophages and T lymphocytes and how to properly gate on them in flow cytometry experiments. Representative flow cytometry plots from low fat-fed lean and high fat-fed obese mice are provided. A critical element of this analysis is the use of antibodies that do not fluoresce in channels where AT macrophages are naturally autofluorescent, as well as the use of proper compensation controls.

Collagenase digestion of AT followed by differential centrifugation was used to isolate the SVF from epididymal fat pads of male C57BL/6J mice fed a low fat (10% kcal from fat) or high fat (60% kcal from fat) diet (LFD and HFD, respectively) for 16 weeks. Cells of the SVF were then labeled with fluorophore-conjugated primary antibodies to quantify the proportion of viable ATMs (Figure 1) and AT T cells (Figure 2) via FACS analysis. Initial gating, including light scatter, doublet ...

Watch this Scientific Journal Video about Isolation of Adipose Tissue Immune Cells at JoVE.com

Isolation of Adipose Tissue Immune Cells

Adipose tissue (AT) is a site of intense immune cell activation and interaction. Almost all cells of the immune system are present in AT and their ratios are altered by obesity. Proper isolation, quantification, and characterization of AT immune cell populations are critical for understanding their role in immunometabolic disease.

The discovery of increased macrophage infiltration in  the adipose tissue (AT) of obese rodents and humans has led to an  intensification of interest in ...

Research

JoVE Journal

Immunology and Infection

36.0K 次观看.  Vanderbilt University School of Medicine. 脂肪组织（AT）是一个强烈的免疫细胞的激活和相互作用的部位。几乎所有的免疫系统的细胞是存在于AT和它们的比率是改变了肥胖。适当的隔离，定量，定性的AT免疫细胞群，是了解自己的角色在immunometabolic疾病的关键。

视频: Isolation of Adipose Tissue Immune Cells

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

Quantitative Assessment of Immune Cells in the Injured Spinal Cord Tissue by Flow Cytometry:  a Novel Use for a Cell Purification Method

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true quantitative analysis of cellular inflammation. Flow cytometry is a quick alternative method to quantify immune cells in the injured brain or spinal cord tissue. Historically, flow cytometry has been used to quantify immune cells collected from blood or dissociated spleen or thymus, and only a few studies have attempted to quantify immune cells in the injured spinal cord by flow cytometry using fresh dissociated cord tissue. However, the dissociated spinal cord tissue is concentrated with myelin debris that can be mistaken for cells and reduce cell count reliability obtained by the flow cytometer. We have advanced a cell preparation method using the OptiPrep gradient system to effectively separate lipid/myelin debris from cells, providing sensitive and reliable quantifications of cellular inflammation in the injured spinal cord by flow cytometry. As described in our recent study (Beck &amp; Nguyen et al., Brain. 2010 Feb; 133 (Pt 2): 433-47), the OptiPrep cell preparation had increased sensitivity to detect cellular inflammation in the injured spinal cord, with counts of specific cell types correlating with injury severity. Critically, novel usage of this method provided the first characterization of acute and chronic cellular inflammation after SCI to include a complete time course for polymorphonuclear leukocytes (PMNs, neutrophils), macrophages/microglia, and T-cells over a period ranging from 2 hours to 180 days post-injury (dpi), identifying a surprising novel second phase of cellular inflammation. Thorough characterization of cellular inflammation using this method may provide a better understanding of neuroinflammation in the injured CNS, and reveal an important multiphasic component of neuroinflammation that may be critical for the design and implementation of rational therapeutic treatment strategies, including both cell-based and pharmacological interventions for SCI.

Quantification of cellular inflammation in the injured/pathological CNS by flow cytometry is complicated by lipid/myelin debris that can have similar size and granulation to cells, decreasing sensitivity/accuracy. We have advanced a cell preparation method to remove myelin debris and improve cell detection by flow cytometry in the injured spinal cord.

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true ...

Seven Steps to Stellate Cells

Hepatic stellate cells are liver-resident cells of star-like morphology and are located in the space of Disse between liver sinusoidal endothelial cells and hepatocytes1,2. Stellate cells are derived from bone marrow precursors and store up to 80% of the total body vitamin A1, 2. Upon activation, stellate cells differentiate into myofibroblasts to produce extracellular matrix, thus contributing to liver fibrosis3. Based on their ability to contract, myofibroblastic stellate cells can regulate the vascular tone associated with portal hypertension4. Recently, we demonstrated that hepatic stellate cells are potent antigen presenting cells and can activate NKT cells as well as conventional T lymphocytes5. 

 Here we present a method for the efficient preparation of hepatic stellate cells from mouse liver. Due to their perisinusoidal localization, the isolation of hepatic stellate cells is a multi-step process. In order to render stellate cells accessible to isolation from the space of Disse, mouse livers are perfused in situ with the digestive enzymes Pronase E and Collagenase P. Following perfusion, the liver tissue is subjected to additional enzymatic treatment with Pronase E and Collagenase P in vitro. Subsequently, the method takes advantage of the massive amount of vitamin A-storing lipid droplets in hepatic stellate cells. This feature allows the separation of stellate cells from other hepatic cell types by centrifugation on an 8% Nycodenz gradient. The protocol described here yields a highly pure and homogenous population of stellate cells. Purity of preparations can be assessed by staining for the marker molecule glial fibrillary acidic protein (GFAP), prior to analysis by fluorescence microscopy or flow cytometry. Further, light microscopy reveals the unique appearance of star-shaped hepatic stellate cells that harbor high amounts of lipid droplets. 


 Taken together, we present a detailed protocol for the efficient isolation of hepatic stellate cells, including representative images of their morphological appearance and GFAP expression that help to define the stellate cell entity.

Here we describe a method for the isolation of hepatic stellate cells from mouse liver. For stellate cell purification, mouse livers are digested in situ and in vitro by pronase-collagenase treatment prior to density gradient centrifugation. This technique yields highly pure hepatic stellate cells.

Hepatic stellate cells are liver-resident  cells of star-like morphology and are located in the space of Disse between  liver sinusoidal endothelial cells ...

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam cells, immune cells, vascular endothelial and smooth muscle cells, platelets, extracellular matrix, and a lipid-rich core with extensive necrosis and fibrosis of surrounding tissues.1 The innate and adaptive arms of the immune response are involved in the initiation, development and persistence of atherosclerosis.2, 3 There is a significant body of evidence that different subsets of the immune cells, such as macrophages, dendritic cells, T and B lymphocytes, are present within the aortas of healthy and atherosclerosis-prone mice4. Additionally, immune cells are found in the surrounding aortic adventitia which suggests an important role of this tissue in atherogenesis.2
For some time, the quantitative detection of different types of immune cells, their activation status, and the cellular composition within the aortic wall was limited by RT-PCR and immunohistochemical methods for the study of atherosclerosis. Few attempts were made to perform flow cytometry using human aortas, and a number of problems, such as a high autofluorescence, have been reported5,6. Human atherosclerotic plaques were digested with collagenase 1, and free cells were collected and stained for CD14+/CD11c+ to highlight macrophage-derived foam cells. In this study, a "mock" channel was used to avoid false-positive staining.6 Necrotic materials accumulating during the digestion process give rise in a large amount of debris that generates a high autofluorescence in aortic samples. To resolve this problem, a panel of negative and positive controls has been proposed, but only double staining could be applied in these samples. We have developed a new flow cytometry-based method7 to analyze the immune cell composition and characterize the activation, proliferation, differentiation of immune cells in healthy and atherosclerosis-prone aorta. This method allows the investigation of the immune cell composition of the aortic wall and opens possibilities to use a broad spectrum of immunological methods for investigations of immune aspects of this disease.

This paper presents a flow cytometry-based method to investigate the immune composition of aortas. The paper also illustrates an additional technique that allows examining surrounding adventitia and vessel wall separately. This method opens possibilities to perform phenotypical analyses of aortic leukocytes and apply several immunological assays for atherosclerosis studies.

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam ...

Isolation of Functional Cardiac Immune Cells

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5 These immune cells, which include mast cells, T-cells and macrophages; store and release a variety of biologically active mediators including cytokines and proteases such as tryptase.6-8 These mediators have been shown to be key players in extracellular matrix metabolism by activating matrix metalloproteinases or causing collagen accumulation by modulating the cardiac fibroblasts' function.9-11 However, available techniques for isolating cardiac immune cells have been problematic because they use bacterial collagenase to digest the myocardial tissue. This technique causes activation of the immune cells and thus a loss of function. For example, cardiac mast cells become significantly less responsive to compounds that cause degranulation.12 Therefore, we developed a technique that allows for the isolation of functional cardiac immune cells which would lead to a better understanding of the role of these cells in cardiac disease.13, 14
This method requires a familiarity with the anatomical location of the rat's xiphoid process, axilla and falciform ligament, and pericardium of the heart. These landmarks are important to increase success of the procedure and to ensure a higher yield of cardiac immune cells. These isolated cardiac immune cells can then be used for characterization of functionality, phenotype, maturity, and co-culture experiments with other cardiac cells to gain a better understanding of their interactions.

This method for isolating functional immune cells from the heart provides an alternative to the conventional methods of collagenase digestion, which causes unwanted immune cell activation, resulting in a decreased responsiveness of these cells. Our method of isolation yields functional cardiac immune cells by avoiding problems associated with enzymatic digestion.

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5  These immune cells, which ...

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Throughout the last years, the contribution of alveolar type II epithelial cells (AECII) to various aspects of immune regulation in the lung has been increasingly recognized. AECII have been shown to participate in cytokine production in inflamed airways and to even act as antigen-presenting cells in both infection and T-cell mediated autoimmunity 1-8. Therefore, they are especially interesting also in clinical contexts such as airway hyper-reactivity to foreign and self-antigens as well as infections that directly or indirectly target AECII. However, our understanding of the detailed immunologic functions served by alveolar type II epithelial cells in the healthy lung as well as in inflammation remains fragmentary. Many studies regarding AECII function are performed using mouse or human alveolar epithelial cell lines 9-12. Working with cell lines certainly offers a range of benefits, such as the availability of large numbers of cells for extensive analyses. However, we believe the use of primary murine AECII allows a better understanding of the role of this cell type in complex processes like infection or autoimmune inflammation. Primary murine AECII can be isolated directly from animals suffering from such respiratory conditions, meaning they have been subject to all additional extrinsic factors playing a role in the analyzed setting. As an example, viable AECII can be isolated from mice intranasally infected with influenza A virus, which primarily targets these cells for replication 13. Importantly, through ex vivo infection of AECII isolated from healthy mice, studies of the cellular responses mounted upon infection can be further extended.
Our protocol for the isolation of primary murine AECII is based on enzymatic digestion of the mouse lung followed by labeling of the resulting cell suspension with antibodies specific for CD11c, CD11b, F4/80, CD19, CD45 and CD16/CD32. Granular AECII are then identified as the unlabeled and sideward scatter high (SSChigh) cell population and are separated by fluorescence activated cell sorting 3.
In comparison to alternative methods of isolating primary epithelial cells from mouse lungs, our protocol for flow cytometric isolation of AECII by negative selection yields untouched, highly viable and pure AECII in relatively short time. Additionally, and in contrast to conventional methods of isolation by panning and depletion of lymphocytes via binding of antibody-coupled magnetic beads 14, 15, flow cytometric cell-sorting allows discrimination by means of cell size and granularity. Given that instrumentation for flow cytometric cell sorting is available, the described procedure can be applied at relatively low costs. Next to standard antibodies and enzymes for lung disintegration, no additional reagents such as magnetic beads are required. The isolated cells are suitable for a wide range of functional and molecular studies, which include in vitro culture and T-cell stimulation assays as well as transcriptome, proteome or secretome analyses 3, 4.

We describe the rapid isolation of primary murine type II alveolar epithelial cells (AECII) by flow cytometric negative selection. These AECII show high viability and purity and are suitable for a wide range of functional and molecular studies regarding their role in respiratory conditions such as autoimmune or infectious diseases.

Throughout  the last years, the contribution of alveolar type II epithelial cells (AECII)  to various aspects of immune regulation in the lung has been ...

Isolation and Flow Cytometric Analysis of Immune Cells from the Ischemic Mouse Brain

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident cells. A key mechanism of this inflammatory response is the migration of circulating immune cells to the ischemic brain facilitated by chemokine release and increased endothelial adhesion molecule expression. Brain-invading leukocytes are well-known contributing to early-stage secondary ischemic injury, but their significance for the termination of inflammation and later brain repair has only recently been noticed.
Here, a simple protocol for the efficient isolation of immune cells from the ischemic mouse brain is provided. After transcardial perfusion, brain hemispheres are dissected and mechanically dissociated. Enzymatic digestion with Liberase&#160;is followed by density gradient (such as Percoll) centrifugation to remove myelin and cell debris. One major advantage of this protocol is the single-layer density gradient procedure which does not require time-consuming preparation of gradients and can be reliably performed. The approach yields highly reproducible cell counts per brain hemisphere and allows for measuring several flow cytometry panels in one biological replicate. Phenotypic characterization and quantification of brain-invading leukocytes after experimental stroke may contribute to a better understanding of their multifaceted roles in ischemic injury and repair.

Inflammation plays a central role in the pathogenesis of ischemic stroke. Increasing evidence suggests that it acts as a double-edged sword which exacerbates early brain injury, but also contributes to later repair. This protocol describes the isolation of immune cells from the ischemic brain and their subsequent flow cytometric phenotyping.

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident ...

Isolation, Characterization and Functional Examination of the Gingival Immune Cell Network

Immune cell networks in tissues play a vital role in mediating local immunity and maintaining tissue homeostasis, yet little is known of the resident immune cell populations in the oral mucosa and gingiva. We have established a technique for the isolation and study of immune cells from murine gingival tissues, an area of constant microbial exposure and a vulnerable site to a common inflammatory disease, periodontitis. Our protocol allows for a detailed phenotypic characterization of the immune cell populations resident in the gingiva, even at steady state. Our procedure also yields sufficient cells with high viability for use in functional studies, such as the assessment of cytokine secretion ex vivo. This combination of phenotypic and functional characterization of the gingival immune cell network should aid towards investigating the mechanisms involved in oral immunity and periodontal homeostasis, but will also advance our understanding of the mechanisms involved in local immunopathology.

We have established a technique for the isolation, phenotypic characterization and functional analysis of immune cells from murine gingiva.

Immune cell networks in tissues play a vital role in mediating local immunity and maintaining tissue homeostasis, yet little is known of the resident ...

Reliable and High Efficiency Extraction of Kidney Immune Cells

Immune system activation occurs in multiple kidney diseases and pathophysiological processes. The immune system consists of both adaptive and innate components and multiple cell types. Sometimes, the cell type of interest is present in very low numbers among the large numbers of total cells isolated from the kidney. Hence, reliable and efficient isolation of kidney mononuclear cell populations is important in order to study the immunological problems associated with kidney diseases. Traditionally, tissue isolation of kidney mononuclear cells have been performed via enzymatic digestions using different varieties and strengths of collagenases/DNAses yielding varying numbers of viable immune cells. Recently, with the development of the mechanical tissue disruptors for single cell isolation, the collagenase digestion step is avoided and replaced by a simple mechanical disruption of the kidneys after extraction from the mouse. Herein, we demonstrate a simple yet efficient method for the isolation of kidney mononuclear cells for every day immune cell extractions. We further demonstrate an example of subset analysis of immune cells in the kidney. Importantly, this technique can be adapted to other soft and non-fibrous tissues such as the liver and brain.

Techniques that are reliable and efficient for the isolation of kidney immune cells are needed for downstream applications. This requires surface antibody labeling of a small number of kidney immune cells. Herein, we describe a concise method for isolation of kidney immune cells that seemingly achieves this goal.

Immune system activation occurs in multiple kidney diseases and pathophysiological processes.  The immune system consists of both adaptive and innate ...

Isolation, Expansion, and Adipogenic Induction of CD34+CD31+ Endothelial Cells from Human Omental and Subcutaneous Adipose Tissue

Obesity is accompanied by an extensive remodeling of adipose tissue primarily via adipocyte hypertrophy. Extreme adipocyte growth results in a poor response to insulin, local hypoxia, and inflammation. By stimulating the differentiation of functional white adipocytes from progenitors, radical hypertrophy of the adipocyte population can be prevented and, consequently, the metabolic health of adipose tissue can be improved along with a reduction of inflammation. Also, by stimulating a differentiation of beige/brown adipocytes, the total body energy expenditure can be increased, resulting in weight loss. This approach could prevent the development of obesity co-morbidities such as type 2 diabetes and cardiovascular disease.
This paper describes the isolation, expansion, and differentiation of white and beige adipocytes from a subset of human adipose tissue endothelial cells that co-express the CD31 and CD34 markers. The method is relatively cheap and is not labor-intensive. It requires access to human adipose tissue and the subcutaneous depot is suitable for sampling. For this protocol, fresh adipose tissue samples from morbidly obese subjects [body mass index (BMI) &#62;35] are collected during bariatric surgery procedures. Using a sequential immunoseparation from the stromal vascular fraction, enough cells are produced from as little as 2&#8211;3 g of fat. These cells can be expanded in culture over 10&#8211;14 days, can be cryopreserved, and retain their adipogenic properties with passaging up to passage 5&#8211;6. The cells are treated for 14 days with an adipogenic cocktail using a combination of human insulin and the PPAR&#947; agonist-rosiglitazone.
This methodology can be used for obtaining proof of concept experiments on molecular mechanisms that drive adipogenic responses in adipose endothelial cells, or for screening new drugs that can enhance the adipogenic response directed either towards white or beige/brown adipocyte differentiation. Using small subcutaneous biopsies, this methodology can be used to screen out non-responder subjects for clinical trials aimed to stimulate beige/brown and white adipocytes for the treatment of obesity and co-morbidities.

The differentiation of white and beige adipocytes from adipose tissue vascular progenitors bears potential for metabolic improvement in obesity. We describe protocols for a CD34+CD31+ endothelial cell isolation from human fat and for a subsequent in vitro expansion and differentiation into white and beige adipocytes. Several downstream applications are discussed.

Obesity is accompanied by an extensive remodeling of adipose tissue primarily via adipocyte hypertrophy. Extreme adipocyte growth results in a poor ...