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Identification, isolation and characterization of murine adult adipogenic progenitor cells Joe, Aaron Wai Bun 2009

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IDENTIFICATION, ISOLATION AND CHARACTERIZATION OF MURINE ADULT ADIPOGENIC PROGENITOR CELLS  by   Aaron Wai Bun Joe  B.Sc. (Hons), University of British Columbia, 2003    A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF   DOCTOR OF PHILOSOPHY   in   The Faculty of Graduate Studies   (Experimental Medicine)     THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)   July 2009    © Aaron Wai Bun Joe, 2009  ii ABSTRACT  White adipose tissue, or fat, is a complex endocrine tissue important for energy storage and metabolism, and has significant effects on various physiological phenomena, including growth, behaviour, reproduction and immune-modulation. It has been proposed that fat cells, or adipocytes, arise from connective tissue cells that fill with lipid; however, mounting evidence suggests that adipocytes represent a distinct lineage with its own cellular origins. Yet very little is known about the cells that give rise to new adipocytes.  Here, my colleagues and I developed a strategy to isolate purified populations of adipogenic progenitor (AP) cells from subcutaneous fat, visceral fat and skeletal muscle, using fluorescence-activated cell sorting. These cells are capable of robust adipogenic differentiation, even at the single cell level. We confirmed their commitment to the adipogenic lineage using a variety of assays, and reveal that they are lineage-restricted cells, incapable of osteogenic, chondrogenic or myogenic differentiation. Thus, we have developed an enabling technology to allow interrogation of the adipocyte lineage among different tissues and fat depots, during different physiological, pathological or developmental stages.  Recent evidence suggests that fat depots with a greater ability to generate new adipocytes are associated with lower metabolic risk. Using our isolation strategy, we confirmed that metabolically healthier depots are associated with greater AP  iii abundance and activity, uncovering a link between stem cell biology and metabolic disease. However, adipocyte production in non-adipose tissues, such as skeletal muscle and bone marrow, is associated with chronic disease and aging. To explore possible reasons for this dichotomy, we examined the role of APs in a model of skeletal muscle injury. Our results suggest that APs expand after damage to assist in muscle regeneration by establishing a pro-myogenic niche, ascribing to them a novel function that is independent of adipogenesis.  Together, our strategy to interrogate the adipogenic lineage has allowed us to formulate new hypotheses to explain adipose and skeletal muscle physiology. This technology forms the basis for future work that will to allow us to understand how new adipocytes are formed, and perhaps permit the manipulation of adipogenic progenitors for therapeutic benefit.   iv TABLE OF CONTENTS  ABSTRACT ............................................................................................................... ii TABLE OF CONTENTS ........................................................................................... iv LIST OF TABLES................................................................................................... viii LIST OF FIGURES ................................................................................................... ix LIST OF ABBREVIATIONS..................................................................................... xii ACKNOWLEDGEMENTS........................................................................................ xv DEDICATION.......................................................................................................... xvi CO-AUTHORSHIP STATEMENT.......................................................................... xvii 1. INTRODUCTION.................................................................................................... 1 1.1. Adipose tissue........................................................................................ 1 1.1.1. Adipose tissue anatomical and cellular organization .................. 1 1.1.2. Physiological roles of adipose tissue .......................................... 3 1.1.3. Origins of adipose tissue............................................................. 4 1.1.4. Adipocytes and adipose tissue cellularity ................................... 7 1.1.5. Differences between depots ....................................................... 8 1.2. Post-natal regulation of adipose tissue mass ................................... 11 1.2.1. Adipocyte hypertrophy .............................................................. 12 1.2.2. Adipocyte hyperplasia............................................................... 13 1.2.3. Proposed regulators of adipocyte hyperplasia.......................... 15 1.3. Pathogenesis of metabolic disease associated with abnormal adipose tissue mass ................................................................................... 16 1.3.1. Obesity...................................................................................... 16 1.3.2. Lipodystrophy ........................................................................... 17  v 1.3.3. The adipose tissue expandability hypothesis............................ 18 1.4. Adipogenesis in non-adipose tissues ................................................ 21 1.5. Objectives of this work ........................................................................ 23 1.6. Tables.................................................................................................... 27 1.7. References............................................................................................ 28 2. Depot-specific differences in adipogenic progenitor abundance and proliferative response to high fat diet .................................................................. 40 2.1. Introduction .......................................................................................... 40 2.2. Results and discussion ....................................................................... 43 2.2.1. Different mechanisms underlie SAT and VAT expansion ......... 43 2.2.2. Prospective identification of adipogenic progenitors................. 44 2.2.3. APs are more abundant in subcutaneous than visceral fat....... 46 2.2.4. AP proliferation in vivo is highly correlated with SAT, but not VAT expansion ................................................................................... 47 2.2.5. Subcutaneous depot APs proliferate in response to high fat diet...................................................................................................... 48 2.2.6. Concluding remarks.................................................................. 48 2.3. Experimental procedures .................................................................... 51 2.4. Tables.................................................................................................... 57 2.5. Figures .................................................................................................. 59 2.6. References............................................................................................ 68 3. Muscle injury activates resident fibro/adipogenic progenitors that Facilitate the terminal differentiation of myogenic cells..................................... 71 3.1. Introduction .......................................................................................... 71 3.2. Results .................................................................................................. 74  vi 3.2.1. Purification of mesenchymal progenitors with distinct developmental potential from skeletal muscle .................................... 74 3.2.2. Single lin- α7- Sca-1+ cells produce both fibroblasts and adipocytes in vitro ............................................................................... 77 3.2.3. FAP engraftment is dictated by the environment ...................... 78 3.2.4. FAPs and MPs have distinct developmental potentials and do not arise from a common progenitor ................................................... 79 3.2.5. Fibro/adipogenic progenitors are rapidly induced to proliferate upon muscle damage......................................................... 81 3.2.6. Fibro/adipogenic progenitors provide an environment favoring myogenic differentiation ........................................................ 84 3.3. Discussion ............................................................................................ 89 3.4. Experimental procedures .................................................................... 94 3.5. Tables.................................................................................................. 101 3.6. Figures ................................................................................................ 102 3.7. References.......................................................................................... 119 4. CONCLUSION................................................................................................... 124 4.1. Thesis summary ................................................................................. 124 4.2. Physiological relevance..................................................................... 128 4.3. Aging as a model to explore AP dysfunction .................................. 130 4.4. Cell-autonomous differences between APs harvested from different depots and tissues..................................................................... 132 4.5. APs from fat and skeletal muscle are closely associated with blood vessels............................................................................................. 135 4.6. Identification of APs in other tissues ............................................... 136 4.7. Developmental origins of AP cells.................................................... 137  vii 4.8. Adipogenic progenitors or adipogenic stem cells? ........................ 138 4.9. Final remarks ...................................................................................... 140 4.10. Figures .............................................................................................. 141 4.11. References........................................................................................ 146 5. APPENDICES.................................................................................................... 150 5.1. Appendix A: Supplementary information for Chapter 2.................. 150 5.2. Appendix B: Supplementary information for Chapter 3.................. 159 5.3. Appendix C. Preliminary data............................................................ 168 5.4. Appendix D. UBC Research Ethics Board certificates of approval174   viii LIST OF TABLES  Table 1.1. Factors secreted by adipocytes. ......................................................... 27 Table 2.1. Limiting dilution analysis of Sca-1/CD34 expressing populations from SAT and VAT. ......................................................................................... 57 Table 3.1. FAP cells do not contribute to skeletal muscle in vivo ................... 101 Table A1. Single lin- Sca-1+ CD34+ cell deposition ............................................ 151   ix LIST OF FIGURES Figure 2.1. Different mechanisms underlie subcutaneous and visceral fat expansion after exposure to HFD.................................................................. 59 Figure 2.2. Prospective isolation of adipogenic cells from SAT and VAT ........ 61 Figure 2.3. Quantification of adipogenic progenitors using limiting dilution analysis............................................................................................................ 63 Figure 2.4. Transplanted lin- Sca-1+ CD34+ cells produce new adipocytes in wildtype, syngeneic animals.......................................................................... 65 Figure 2.5. Greater adipogenic progenitor activity in subcutaneous compared to visceral fat................................................................................. 67 Figure 3.1. Prospective isolation of progenitor populations from skeletal muscle............................................................................................................ 102 Figure 3.2. Lin- Sca-1+ CD34+ cells generate both fibroblasts and adipocytes105 Figure 3.3. Developmental potential of sorted progenitor populations in vivo................................................................................................................. 106 Figure 3.4. Skeletal muscle-derived FAPs and MPs have distinct developmental potentials and do not arise from a common progenitor.. 108 Figure 3.5. FAPs proliferate in response to muscle damage ........................... 111 Figure 3.6. FAPs are in close proximity to myofibers and express pro- differentiation signals after damage............................................................ 114 Figure 3.7. FAPs enhance myogenic differentiation ......................................... 116 Figure 4.1. Proposed model for the effect of progenitor content and activity on fat depot expansion................................................................................. 141 Figure 4.2. Models of stem/progenitor cell contribution to fibro/adipogenic infiltration of skeletal muscle....................................................................... 142  x Figure 4.3. Proposed model for the role of fibro/adipogenic progenitors in normal and impaired skeletal muscle regeneration................................... 144 Figure A1. Detection of BrdU labeling in adipocyte nuclei by immuno- electron microscopy (IEM) ........................................................................... 152 Figure A2. Confirmation of Sca-1 and CD34 stain specificity using isotype control antibodies......................................................................................... 154 Figure A3. Detection of BrdU labeling of endogenous lin- Sca-1+ CD34+ cells by flow cytometry ......................................................................................... 155 Figure A4. Proliferation of non-adipogenic R3 (lin- Sca-1+ CD34+) cells is not correlated to fat depot size .......................................................................... 157 Figure B1. Confirmation of Sca-1 CD34 stain specificity using isotype- matched control antibodies ......................................................................... 160 Figure B2. Linear regression of colony forming data from freshly isolated skeletal muscle DN (Sca-1- CD34-) cells...................................................... 161 Figure B3. Skeletal muscle DP (Sca-1+ CD34+) cells do not form bone or cartilage. ........................................................................................................ 162 Figure B4. Quantitative gene expression in sorted cells.................................. 163 Figure B5. An alternative strategy using α7 integrin to identify adipogenic and myogenic cells from dissociated skeletal muscle.............................. 165 Figure B6. Co-cultivation control experiment.................................................... 166 Figure B7. Sca-1+ mononuclear cells are juxtaposed to skeletal muscle fibers after damage....................................................................................... 167 Figure C1. The effect of aging on adipogenic progenitor (AP) abundance in subcutaneous and visceral fat depots........................................................ 169 Figure C2. The effect of aging on adipogenic (AP) and myogenic (MP) progenitor abundance in skeletal muscle................................................... 170  xi Figure C3. The prospective isolation of adipogenic progenitor cells from tendon............................................................................................................ 172   xii LIST OF ABBREVIATIONS  AF Alexa fluor AP adipogenic progenitor APC allophycocyanin BD Becton Dickinson bFGF basic fibroblast growth factor BrdU 5-bromo-2-deoxyuridine BSA bovine serum albumin CD cluster of differentiation CFU-AP colony forming unit- adipocyte progenitor DMEM Dulbecco's modified eagle medium DN double negative DNA deoxyribonucleic acid DP double positive EDTA ethylenediaminetetraacetate FACS fluorescence-activated cell sorting FAP fibro-adipogenic progenitor FITC fluorescein isothiocyanate FMO fluorescence-minus one FSP fibroblast specific protein g gram GFP green fluorescent protein GH growth hormone H&E hematoxylin and eosin HDL high density lipoprotein HFD high fat diet IBMX isobutylmethylxanthine IEM immuno-electron microscopy Ig immunoglobulin  xiii IGF insulin-like growth factor IL interleukin IP intraperitoneal kg kilogram L litre LDA limiting dilution analysis LDL low density lipoprotein lin lineage markers mg milligram ml millilitre MP myogenic progenitor MSC mesenchymal stem cell MTT 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide MyHC myosin heavy chain ND no signal detected NTX notexin PBS phosphate-buffered saline PCR polymerase chain reaction PDGF platelet-derived growth factor PE phycoerythrin PECY7 phycoerythrin cy7 PFA paraformaldehyde PI propidium iodide qPCR quantitative polymerase chain reaction qRT-PCR quantitative reverse transcription polymerase chain reaction RCF relative centrifugal force RNA ribonucleic acid S-AP subcutaneous adipogenic progenitors SAT subcutaneous adipose tissue SD standard diet OR standard deviation SEM standard error of the mean  xiv SMA smooth muscle actin SV stromovascular TBP TATA-binding protein TGF transforming growth factor TNF tumor necrosis factor TPBS Tris phosphate buffered saline µg microgram µ l microlitre V-AP visceral adipogenic progenitors VAT visceral adipose tissue VLDL very low density lipoprotein YFP yellow fluorescent protein    xv ACKNOWLEDGEMENTS  I am grateful to all my past teachers, who drove my curiosity and nurtured my development. Particularly, I thank Dr. Fabio Rossi, who over the past 8 years has had the greatest impact on my development, and provided me with an environment in which I could both learn and succeed.  I have received much support from colleagues and professors while navigating the murky waters of Science and Academia. Particularly, I thank my supervisory committee for their inspiration, and the UBC MD/PhD Program for providing me with the opportunity to follow my passion and to present my research all over the world.  I thank the many past and present members of the Biomedical Research Centre for their camaraderie and expertise. Notably, I thank V. Canale, C-K. Chang, S. Corbel, J. Duenas, Y. Even, H. Frei, V. Gylfadottir, J. Haddon, A. Johnson, D. Lemos, K. McNagny, A. Natarajan, J. Qiao, A. Scott, L. So, J. Wang and C. Yang. Importantly, this work could not have been performed without the findings and technical expertise of L. Yi, for which I am especially grateful.  Finally, I thank my friends and family, who remain a great source of love and kindness. Above all, I wish to acknowledge my wife Jennifer, without whom none of this would have been possible.  xvi DEDICATION         To all my family: past, present and future.  xvii CO-AUTHORSHIP STATEMENT  This thesis contains manuscripts that have been submitted for publication, for all of which I am the lead author. I have been fortunate to have received assistance from co-authors. Our specific contributions are detailed here.  Chapter 2. I designed and performed experiments, analyzed data, interpreted results and wrote the manuscript. L. Yi designed and performed experiments and analyzed data. Y. Even and A.W. Vogl designed and performed experiments. F.M.V. Rossi designed experiments, interpreted results and co-wrote the manuscript.  Chapter 3. I designed and performed experiments, analyzed data, interpreted results and wrote the manuscript. L. Yi designed and performed experiments and analyzed data. A. Natarajan designed and performed experiments. L. So and J. Wang performed experiments. F. Le Grand and M. Rudnicki provided novel reagents and performed experiments. F.M.V. Rossi designed experiments, interpreted results and co-wrote the manuscript.   1 1. INTRODUCTION 1.1. ADIPOSE TISSUE The production and storage of energy in the form of fat is a highly conserved and ancient process, and is critical for survival in both invertebrates and vertebrates (McKay et al., 2003). In vertebrates, fat is stored in adipose tissue, which is primarily comprised of specialized cells for lipid storage and metabolism, called adipocytes. Two types of adipocytes exist in mammals, white and brown, which are typically located in distinct depots (Gesta et al., 2007). Both types of adipocytes are characterized by lipid-filled vacuoles; however, white adipocytes store energy whereas brown adipocytes dissipate energy as heat (Cypess et al., 2009; Enerback, 2009; Nechad, 1986; Seale et al., 2008). Several functional and ontological differences exist between both types of adipocytes, many of which are under intense investigation and are beyond the scope of this thesis. Here, I present research performed under the paradigm of white adipogenesis; therefore, references to fat, adipose tissue and adipocytes in this work refer to white adipose tissue and adipocytes unless otherwise specified.  1.1.1. Adipose tissue anatomical and cellular organization In mammals, fat is distributed all throughout the body. Although there are variations in the specific sites of fat accumulation between species, subcutaneous and intra- abdominal fat represent the two major types of depots common to all mammals  2 (Gesta et al., 2007). In humans, fat is typically found in subcutaneous depots surrounding the thighs, buttocks and abdomen, in intra-abdominal depots throughout the omentum and around the intestines and kidneys, and in other sites including the intermuscular, perivascular, intra-articular, retro-orbital and medullary (bone marrow) spaces (Gesta et al., 2007; Slavin, 1985). In rodents, large depots of visceral fat are also located adjacent the gonads, called epididymal fat pads in males, and parametrial fat pads in females (Slavin, 1985).  Histological analysis of adipose tissue reveals mostly a dense carpet of large, lipid- laden, spherical or oval shaped adipocytes, interspersed with occasional blood vessels, macrophages, granulocytes and loose connective tissue stroma (Hausman et al., 2001; Slavin, 1985). The adipose tissue vascular stroma contains blood vessels, nerves and stromal cells, as well as collagen and reticular fibers that provide mechanical support (Hausman and Hausman, 2006; Slavin, 1985). Adipose tissue is highly-vascularized, with each adipocyte in contact with at least one capillary (Slavin, 1985). Studies have shown that adipose tissue blood supply is equal to or greater than that found in skeletal muscle, and in obese patients can approach 30% of cardiac output (Gersh and Still, 1945; Hausberger and Widelitz, 1963; Hausman et al., 2001; Slavin, 1985). As a result, circulating factors, including glucocorticoids and sex hormones, can significantly alter adipose tissue physiology (Gesta et al., 2007). Neuronal inputs also affect adipose tissue function, as both classical and contemporary studies have confirmed that fat contains rich sympathetic innervation that plays an important role in regulating lipolysis, controlling  3 the mobilization of lipids (Hausman et al., 2001; Slavin, 1985; Youngstrom and Bartness, 1995). Thus, the cellular organization of adipose tissue reveals it to be a dynamic tissue capable of endocrine and neuronal modulation.  1.1.2. Physiological roles of adipose tissue Although traditionally viewed as an organ for energy storage, lipid handling and thermal insulation, developments over the past decade have shown that fat plays an emerging role as an endocrine organ capable of influencing physiological processes ranging from metabolism to behaviour and reproduction (Fliers et al., 2003; Trayhurn and Beattie, 2001). Nonetheless free fatty acids and cholesterol represent the vast majority of products released by adipocytes (Trayhurn and Beattie, 2001). Free fatty acids are quantitatively the greatest energy substrate in animals, are stored in adipocytes as triglycerides, and are mobilized upon sympathetic activation of hormone-sensitive lipase (Belfrage et al., 1983; Trayhurn and Beattie, 2001). Cholesterol is also stored and released by adipocytes, and is a key component of cell membranes, bile and steroid hormones (Angel and Fong, 1983; Trayhurn and Beattie, 2001). Thus, by regulating these key processes, fat tissue is critically important for survival; nonetheless, it may have an even larger physiological role as it also releases a number of secreted protein products, the various functions of which are only recently becoming elucidated.  The role of adipose tissue as an endocrine organ was highlighted by the discovery of leptin, which generated a sea change in the field, revealing for the first time that  4 adipocytes were capable of secreting proteins that could influence distant organs (Zhang et al., 1994). Although the most established view of leptin suggests that it leads to an inhibition of food intake, the mechanistic detail is not completely clear (Fliers et al., 2003; Trayhurn and Beattie, 2001). Leptin is also known to interact with a number of neuroendocrine systems, including the hypothalamic-pituitary-adrenal axis, neuropeptide Y and melanocortin, and therefore can affect growth, reproduction, stress response, circadian rhythm and emotion (Fliers et al., 2003; Kishi and Elmquist, 2005; Trayhurn and Beattie, 2001). Adipose tissue is known to secrete a number of other bioactive molecules, including adiponectin, adipsin, plasminogen activator inhibitor-1, resistin, angiotensinogen, tumor necrosis factor-α (TNF-α), interleukin-6 (IL-6), and transforming growth factor β (TGF-β) (Fliers et al., 2003; Kishi and Elmquist, 2005; Matsuzawa, 2006b; Trayhurn and Beattie, 2001). The effects of these molecules are diverse and under intense scrutiny, and include hemodynamic regulation, effects on energy expenditure and insulin sensitivity, and immune- and inflammatory modulation (a summary of the best-characterized adipocyte-secreted factors is shown in Table 1.1). Clearly, adipose tissue is a specialized tissue that plays an incredibly complex role in normal physiology and in the pathogenesis of metabolic disease.  1.1.3. Origins of adipose tissue Adipose tissue is generally thought to arise from mesoderm; however, like skeletal muscle and bone comprising the skull and face, adipose tissues around the head have been shown to arise from neural crest, which is of ectodermal origin (Billon et  5 al., 2007; Gesta et al., 2007). Thus, like bone, adipose is thought to arise from embryonic mesenchyme, regardless of germ layer. However, nothing is known about the cells that give rise to adipose tissue largely due to a lack of markers identifying them.  A number of classical, descriptive studies in animal fetuses have detailed the events leading to the appearance of adipocytes; however, the broad interpretation of their data is difficult due to great variation in these events among both species and depots (Hausman, 1985; Hausman et al., 1980). For example, little or no cellular organization is observed in the primordial adipose tissue of both subcutaneous and intra-abdominal depots of the dog, sheep, guinea pig, rabbit and calf (Bell, 1909; Flemming, 1871; Hammar, 1895); however development of the same depots in rodents is characterized by a low-level condensation of cells immediately prior to adipocyte formation (Hammar, 1895; Wasserman, 1965). On the other hand, a great deal of cellular organization and cell condensation is found in the adipose tissue primordia of the interscapular fat pad in rodents, and in all depots in the fetal sheep and goat (Hammar, 1895; Thompson and Jenkins, 1970; Wensvoort, 1967). These studies suggest depot-specific differences in the steps leading up to adipocyte formation, and potentially the precursor cells that become them. Yet, despite the disparities between depots, a spatial and temporal relationship between adipocyte and blood vessel formation has been consistently described, likely highlighting a significant relationship in their ontogeny (Hausman et al., 1980). However,  6 confirmation of this remains to be seen, as the cellular origin of adipose tissue is unknown.  Historically, it was put forward that adipose tissue is the result of lipid storage in connective tissue cells such as fibroblasts, and is not a specialized tissue with its own developmental lineage (Flemming, 1871; Hausman, 1985; Virchow, 1857). The concept that adipose tissue had a distinct origin from connective tissue began to take hold after the publication of a case report in 1915, which described the development of a large fat pad arising from an autologous abdominal skin graft onto the back of a patient’s hand (Cahill and Renold, 1983; Strandberg, 1915). This revealed differences in the adipogenic potential between different skin regions and suggested the existence of specialized cells in the abdominal skin that are destined to become adipocytes.  Still, the concept that adipose tissue can arise from fibroblasts and connective tissues in vivo remains controversial, and has been supported by the generation of fibroblast cell lines (including 3T3-L1 and 3T3-F422A) that are capable of efficient adipogenic differentiation (Green and Kehinde, 1974). However, that some tissues never generate fat, yet fibroblasts can be found throughout the body, suggests that only certain types of fibroblasts are adipogenic (Hausman, 1985). These discrepancies may be due to the relative ease with which cells are called ‘fibroblasts’. Fibroblasts are generally identified based on in vitro morphology and growth characteristics, and not molecular markers. Furthermore, the term is typically  7 reserved as a default classification for cells that lack the ability to differentiate into other cell types. Thus, although adipocytes may arise from cells that morphologically resemble fibroblasts, since the ability to undergo adipogenesis is only found in a rare subset of these cells, adipose tissue precursors likely represent a distinct population. Furthermore, recent developments revealing the role of adipose tissue as an endocrine organ strongly suggest that adipocytes are specialized cells, and not simply the result of fibroblasts filling with lipid. Molecular strategies allowing the positive identification of both fibroblasts and adipogenic precursors will allow the careful examination of their developmental relationship, and provide important insights in to the ontogeny of the adipose lineage.  1.1.4. Adipocytes and adipose tissue cellularity Adipose tissue is mainly comprised of adipocytes, which represent the smallest functional unit of fat tissue and are collectively responsible for the physiological effects of a depot. Mature adipocytes are spherical or oval shaped cells with a flattened nucleus and a thin rim of cytoplasm surrounding a large central lipid droplet (Slavin, 1987). Adipocyte diameter increases by about 10-fold during lipid accumulation, representing an increase in cell surface area and volume of 100-fold and 1000-fold, respectively (Bjorntorp and Sjostrom, 1985). Despite the scant cytoplasm, electron microscopy has revealed that adipocytes contain a variety of organelles similar to that found in most cells, including a typical plasma membrane, numerous mitochondria, Golgi apparatus, microtubules, microfilaments, ribonucleoproteins, rough and smooth endoplasmic reticulum and a number of  8 plasmalemmal invaginations which may represent sites involved with lipid transport (Slavin, 1985, 1987). Thus, analysis of adipocyte structure and function reveal them to be metabolically active cells that are uniquely well suited as a storage depot for lipids, and as a dynamic source of lipid and protein products.  Despite the fact that most adipose tissues are exposed to similar levels of circulating factors due to their high vascularity, there have been reports of differences in the size of adipocytes both within a depot, and between different depots (Hausman et al., 2001). Furthermore, characterization of small versus large adipocytes has revealed significant differences in their function, gene expression and secretory profile (Bluher et al., 2002; Bluher et al., 2004; Jernas et al., 2006; Khan et al., 2009; Rajala and Scherer, 2003). Together, these findings suggest heterogeneity in the adipocyte population. Although the implications of this are currently unclear, it highlights the need to consider adipose tissue physiology as a complex array of adipocytes and depots, each of which may carry a different function.  1.1.5. Differences between depots Clinical studies have consistently revealed that greater metabolic risks are specifically attributed to the expansion of the visceral fat depot, suggesting the existence of depot-specific differences in adipose tissue function (Cali and Caprio, 2009; Gesta et al., 2007; Wang et al., 2005). There are two non-exclusive theories that may explain this phenomenon: 1) anatomical proximity to the liver and drainage into the portal circulation grants visceral fat the ability to preferentially influence  9 metabolism; and 2) the existence of depot-specific, intrinsic differences in adipocyte function results in fat depots that have different physiological effects (Bjorntorp, 1990; Gesta et al., 2007; Lafontan and Berlan, 2003).  Recent evidence from studies in which different fat pads were transplanted subcutaneously or viscerally into wild-type mice, confirmed that intrinsic characteristics of fat depots can play a role in determining their effects on metabolism (Tran et al., 2008). In support of this phenomenon, depot-specific patterns in adipocyte size and number have emerged. For example, it has been shown that subcutaneous depots typically contain smaller adipocytes than visceral depots, and as mentioned above, adipocyte size can be a major determinant of adipocyte function and secretory profile. Moreover, cells isolated from the stromo- vascular fraction of depots containing smaller adipocytes tend to have greater proliferative capacity (Crandall and DiGirolamo, 1990; Crandall et al., 1984; Hausman et al., 2001). Although the causes of these differences are unclear, it is plausible that there may be disparities in their adipogenic progenitors (AP), which may thereby confer different depots with different types of adipocytes.  The examination of depot-specific differences in adipocytes is a relatively new development. Most of what is known about adipogenesis arose from studies performed on immortalized fibroblasts, such as the 3T3-L1 or 3T3-F422A, which faithfully recapitulate the biochemical steps resulting in lipid droplet formation but cannot reproduce depot-specific effects. Therefore, other experimental paradigms  10 are required to examine these disparities. In support of cell-autonomous differences in adipocytes and their precursors, patients with congenital generalized lipodystrophy, who carry a mutation in the seipin gene (encoding a protein of unknown function), lack adipose tissue in subcutaneous, visceral, intrathoracic and bone marrow depots, yet retain normal fat in the retro-orbital space, buccal region, and in the palms and soles (Agarwal and Garg, 2006; Gesta et al., 2007). Similarly, administration of a monoclonal antibody raised against adipocyte plasma membranes to chick embryos resulted in a reduction in visceral fat without any effect on the femoral or pectoral fat depots, suggesting that adipocytes from different depots do not share the same surface antigens (Wu et al., 2000). In an attempt to compare the adipocyte precursor populations, differences in adipogenicity have been described between primary stromo-vascular (SV) cells isolated from different depots (Tchkonia et al., 2006). Furthermore, microarray analysis revealed significant disparities in gene expression between SV cells isolated from mouse subcutaneous versus visceral fat (Gesta et al., 2006). Nonetheless, data generated from SV cells is confounded by the non-adipogenic cell subsets, which include numerous vessels, endothelial cells, stromal cells and fibroblasts. Thus, the field awaits the development of strategies to prospectively purify APs to allow for the direct investigation of adipocytes and their precursors isolated from different depots.    11 1.2. POST-NATAL REGULATION OF ADIPOSE TISSUE MASS Fat mass must increase in size to accommodate the growth of the organism and its ability to store nutritional excess. Classical literature suggested that no new adipocytes are generated after birth, and that throughout life adipocytes simply grow in size to meet caloric storage requirements (Rosen and Spiegelman, 2000). However, experiments using labeled nucleotides over the past several decades have shown that new adipocytes can indeed be generated. Thus, it is clear that fat mass expansion is the result of two non-exclusive growth modalities: increasing adipocyte size (hypertrophy) and number (hyperplasia). Although changes in adipocyte size is thought to be governed by nutrition levels and modulators of lipogenesis and lipolysis, little is known about the cellular source of new adipocytes, and whether it only arises in early life as a consequence of adipose tissue development, occurs throughout life as a consequence of normal cell turnover, or can be modulated by external factors (Greenwood and Hirsch, 1974; Hausman, 1985; Johnson et al., 1978; Miller et al., 1984; Rosen and Spiegelman, 2000; Spalding et al., 2008). Furthermore, the emerging importance of depot-specific adipose physiology implies that regulation of hypertrophy and hyperplasia may occur differently depending on the fat pad. Therefore, interrogation of APs isolated from different depots may provide the answer to many of these fundamental questions.   12 1.2.1. Adipocyte hypertrophy Adipocyte hypertrophy is the increase in adipocyte size that results from lipid accumulation into existing adipocytes due to a positive caloric balance. Comparative analyses of adipose cellularity in a number of species and depots revealed that total body adiposity is more tightly associated with adipocyte hypertrophy than hyperplasia, suggesting that it is the more robust modality used for fat tissue expansion (Hausman, 1985). Adipocyte size is considered to be the passive by- product of mechanisms controlling lipid storage and lipid mobilization, which are dependent on an organism’s energy balance. Hypertrophy results mostly from the uptake of circulating lipids arising from dietary fats by the low density lipoprotein (LDL) receptor family (Tacken et al., 2001). Although the new production of fatty acids can occur in adipocytes, it is regulated by hormones such as insulin and glucagon, and is a relatively minor contributor to total adipocyte lipid (Large et al., 2004).  Histological analysis of fat tissue taken from obese subjects suggests that hypertrophy precedes hyperplasia during tissue expansion; however, more recent work suggests that these characteristics may be depot-specific (Bjorntorp, 1974; Bjorntorp et al., 1979; Hausman et al., 2001). For example, intra-abdominal adipocytes, including adipocytes in the visceral fat pad, tend to grow by hypertrophy (DiGirolamo et al., 1998). Since adipocyte function and secretory profile are related to cell size, and visceral fat expansion is associated with the onset of metabolic disease, a link between visceral fat function and growth modality has been  13 proposed. Indeed, adipocyte hypertrophy has been associated with insulin insensitivity, adipose inflammation and adipocyte death (Khan et al., 2009; Livingston et al., 1972; Rajala and Scherer, 2003). Thus, although all fat cells must increase in size during lipid accumulation, excessive hypertrophy is associated with adipocyte dysfunction.  1.2.2. Adipocyte hyperplasia Adipose hyperplasia, also called de novo adipogenesis, is the production of new fat cells. This phenomenon is less robust than adipose hypertrophy, and has been detected in careful nucleotide incorporation studies, which revealed the existence of labeled DNA in mature adipocytes. Although this result may be interpreted as arising from either mitotic division of mature adipocytes or production of new fat cells from a progenitor population, the first hypothesis is usually ignored because other than rare reports of de-differentiation resulting in proliferation in vitro, the division of mature adipocytes has not been observed and is generally considered impossible (Hausman et al., 2001). Moreover, the existence of rare subsets of adipose SV cells capable of adipogenic differentiation suggests that a progenitor population exists in the adipose vascular stroma, supporting the second hypothesis. Yet, little is known about the regulation of these cells in vivo.  Depot-specific differences in adipose hyperplasia have been observed, with the subcutaneous depot growing more by hyperplasia than the intra-abdominal depots (DiGirolamo et al., 1998). This preference towards a hyperplastic response may be  14 attributed to depot-specific differences in adipogenic progenitor content or activity, as the greatest proliferation is observed in SV cells isolated from subcutaneous compared to intra-abdominal depots (Djian et al., 1983; Wang et al., 1989). Notably, fat depots that tend to grow by hypertrophy are also the ones with the lowest tendency to grow by hyperplasia, and have the lowest proliferative capacity in their SV cells (Hausman et al., 2001). Although, as noted above, these types of results are confounded by the fact that SV cells are a heterogeneous population also containing cells of the vasculature and stroma, they suggest an important link between progenitor activity and adipose growth modality.  Recent data suggests a link between adipose hyperplasia and adipose tissue function. Subcutaneous adipocyte hyperplasia commonly occurs in type-2 diabetics treated with thiazolidinediones and is associated with improved insulin sensitivity and metabolic profiles (Spiegelman, 1998). Likewise, in ob/ob mutant mice over- expressing adiponectin (tgAdipoQ-ob/ob, a model of severe obesity mainly due to adipocyte hyperplasia), the progression of metabolic disease was much diminished compared to typical ob/ob mice and metabolic parameters were comparable to that of wild type animals (Kim et al., 2007). Most strikingly, these results were observed despite that tgAdipoQ-ob/ob animals have twice the fat mass of normal ob/ob, suggesting that fat tissue quality has a greater effect on metabolic pathogenesis than quantity. Thus, expansion by adipocyte hyperplasia may allow a fat pad to maintain a pool of smaller, healthier, more functional adipocytes.   15 1.2.3. Proposed regulators of adipocyte hyperplasia Since markers for the purification of APs had not been identified, very little is known about the cells that give rise to new adipocytes in vivo. As a result, examination of the factors regulating adipocyte hyperplasia relied on indirect methods, such as histological enumeration of adipose cellularity, and on in vitro analyses of SV cells or immortalized fibroblasts. Based on these types of studies, over the past couple of decades a number of hormonal, neuronal and paracrine modulators de novo adipogenesis have been proposed, including glucocorticoids, growth hormone (GH), insulin-like growth factor-1 (IGF-1), thyroxine, adrenergics and the sympathetic nervous system, TGF-β, TNF-α, and basic fibroblast growth factor (bFGF) (Chen et al., 1996; Hausman et al., 2001; Hausman et al., 1990; Hausman et al., 1993; Kras et al., 1999; Kras et al., 2000; Richardson et al., 1992; Teichert-Kuliszewska et al., 1992; Wright and Hausman, 1993). Although these factors may well have effects on adipose biology, whether they specifically regulate APs, and the level at which act (e.g. survival, expansion, commitment or differentiation), remains unclear.   16 1.3. PATHOGENESIS OF METABOLIC DISEASE ASSOCIATED WITH ABNORMAL ADIPOSE TISSUE MASS Adipose tissue is an endocrine gland capable of regulating a number of physiological systems, and also plays critical roles in energy balance, lipid metabolism, and thermal regulation. Thus, changes in adipose tissue mass can result in serious, and potentially life-threatening, consequences. As mentioned, a change in fat mass requires changes in adipocyte size and/or number. Likewise, recent studies suggest that dysregulation of fat mass alone is insufficient to cause metabolic disease. To explore the relationship between adipose cellularity, adipocyte function and metabolic disease, here I briefly discuss two extreme conditions of abnormal adipose mass: obesity and lipodystrophy.  1.3.1. Obesity Obesity is characterized by excessive fat tissue expansion, significantly contributes to disease and mortality worldwide, and is approaching pandemic levels in the industrialized world (Belanger-Ducharme and Tremblay, 2005; Roth et al., 2004; Weiss et al., 2004). Despite that obese individuals are at an elevated risk for a number of non-metabolic complications, including osteoarthritis, cancer and sleep apnea, the vast majority of obesity-related mortality is the result of metabolic complications, including insulin-resistance, dyslipidemia, type-2 diabetes and cardio- and cerebro-vascular disease, which together comprise the ‘metabolic syndrome’ (Reaven, 1988; Virtue and Vidal-Puig, 2008; Weiss et al., 2004). Particularly, insulin  17 resistance in obese individuals plays a central role in the development of many of these metabolic complications and is well-established on a population level; however, its etiology remains unclear (DeFronzo and Ferrannini, 1991; Virtue and Vidal-Puig, 2008). Proposals that obesity-related adipose tissue inflammation and changes in the adipocyte secretory profile lead to insulin resistance are well- supported and remains active areas of research (Lazar, 2005; Matsuzawa, 2006a; Weisberg et al., 2003). Yet, some of the morbidly obese retain normal insulin resistance and metabolic profiles (Karelis et al., 2005; Kim et al., 2007; Virtue and Vidal-Puig, 2008). Thus, other changes in the activity or function of adipocytes or their progenitors are likely to be also required for metabolic disease progression.  1.3.2. Lipodystrophy Intriguingly, a failure in the development or function of adipose tissue, called lipodystrophy, also leads to the development of insulin resistance and the metabolic syndrome (Moitra et al., 1998; Savage et al., 2003). Although different types of lipodystrophy have been described in both humans and mouse models, they are all characterized by an inability to accumulate lipids in adipocytes. It is believed that this excess in circulating lipids is shunted towards ectopic storage in the liver or skeletal muscle, and the resulting ‘lipotoxicity’ leads to the development of insulin resistance, likely through modifications in insulin signaling (Garg, 2006; Gray and Vidal-Puig, 2007; Heilbronn et al., 2004; Kim et al., 2000; Moitra et al., 1998; Savage et al., 2003; Virtue and Vidal-Puig, 2008). Together, this causes impairment in glucose uptake, elevation of blood glucose levels, and the development of diabetes and  18 cardio- and cerebro-vascular disease. It is striking that despite being at opposite ends of the fat mass spectrum, obesity and lipodystrophy both present with identical metabolic ramifications. These findings suggest that the mechanisms underlying their pathogenesis may be shared.  1.3.3. The adipose tissue expandability hypothesis In lipodystrophy, the inability to appropriately generate or expand adipose tissue to store lipids leads to hyperlipidemia, which underlies the development of insulin resistance and the metabolic syndrome. Indeed, transplantation of functional fat into lipodystrophic A/ZIP mice results in a complete reversal of the associated diabetes (Gavrilova et al., 2000). Therefore, it appears that functioning adipose tissue can act as a buffer for circulating lipids, preventing fat accumulation in the liver or skeletal muscle. In the case of obesity, it is unclear whether the same inability to appropriately store lipids underlies the development of metabolic disease. In support of this theory, the presence of ectopic fat storage in obese patients is pathognomonic of insulin resistance, thus revealing a link between hyperlipidemia and the development of the metabolic syndrome (Heilbronn et al., 2004). Likewise, several lines of evidence suggest that increasing the cellularity of adipose tissue by de novo adipogenesis is associated with an amelioration of metabolic parameters associated with obesity. Firstly, thiazolidinediones, which stimulate new adipocyte production, are effective in the treatment of non-alcoholic fatty liver disease and type 2 diabetes, and improves dyslipidemia, insulin sensitivity and blood glucose levels (Spiegelman, 1998; Virtue and Vidal-Puig, 2008). Secondly, fat depots that are less  19 expandable are associated with increased metabolic risk. As mentioned above, visceral fat tends to grow by hypertrophy rather than hyperplasia, resulting in alterations in adipocyte function and secretory profile, and an increased association with increased metabolic disease. Thirdly, fat depots, such as the subcutaneous depot, that have a large capacity for de novo adipogenesis, are associated with reduced metabolic risk. Likewise, morbidly obese patients who do not develop metabolic disease appear to deposit fat in their subcutaneous depots (Karelis et al., 2005). Further, these observations are unlikely to be limited to subcutaneous depot, as the grossly obese yet metabolically healthy tgAdipoQ-ob/ob appears to expand all of its fat by hyperplasia (Kim et al., 2007). Together, it appears that increasing the ability to generate new adipocytes may be protective against the development of metabolic disease.  Therefore, a key similarity in the development of insulin resistance in both lipodystrophy and obesity is that adipose tissue loses its ability to buffer circulating lipids. The adipose tissue expandability hypothesis posits that adipose tissue has a limited capacity to increase in mass, which is determined both by genetic, environmental and possibly depot-specific factors influencing hypertrophy and hyperplasia (Gray and Vidal-Puig, 2007; Virtue and Vidal-Puig, 2008). As this capacity is reached, the majority of adipocytes become excessively hypertrophic, and adipose tissue gradually loses its ability to accommodate extra lipid, resulting in systemic lipotoxicity. Therefore, it appears that adipose expansion by hypertrophy is only effective up to a certain limit, after which adipocyte hyperplasia must take over  20 to prevent the changes in adipocyte function associated with excess size. Hence, the ability to generate new adipocytes increases the total lipid buffering capacity. Together, this hypothesis presents a theory that unifies the mechanisms underlying the development of metabolic disease and explains that depot-specific differences in metabolic risk may be related to differences in depot expandability. Although other mechanisms are likely to also contribute to metabolic pathogenesis, the expandability hypothesis importantly underscores the significance of identifying the cells responsible for de novo adipogenesis.   21 1.4. ADIPOGENESIS IN NON-ADIPOSE TISSUES Ectopic fat accumulation accompanying systemic lipotoxicity is the result of aberrant intracellular lipid deposition, primarily in hepatocytes, pancreatic β-cells, myocardiocytes, and skeletal myocytes (Higa et al., 1999; Lee et al., 1994; Lee et al., 2001; Zhou et al., 2000). This often results in insidious changes in parenchymal cell function, including alteration of normal signaling pathways and induction of apoptosis (Anderson and Borlak, 2008; Unger, 2005). It is important to make the distinction between aberrant intracellular lipid deposition and true adipogenesis, which is defined by the differentiation and maturation of adipocytes from a precursor population. Both have been observed in non-adipose tissues, but the former requires assimilation and changes in function of parenchymal cells, whereas the latter results in the production of new adipocytes from a progenitor population and is responsible for adipose hyperplasia.  De novo adipogenesis has been observed in skeletal muscle, tendon and bone marrow, suggesting that APs exist in these tissues; however, adipocyte infiltration into all three tissues is a robust event associated with chronic disease and aging, suggesting that AP activation is part of a pathological process (Hashimoto et al., 2003; Jarvinen et al., 1997; Rosen and Bouxsein, 2006; Wallace and McNally, 2008). In bone marrow, adipocytes are thought to arise from mesenchymal stem cells (MSC), a cell population derived from bone marrow stromal cell cultures that are capable of differentiation along the osteogenic, chondrogenic and adipogenic lineages (Pittenger et al., 1999). Yet, MSCs have never been prospectively isolated,  22 and their contribution to physiological tissue homeostasis is unclear; therefore, it is controversial whether they exist in vivo (Javazon et al., 2004). In skeletal muscle, there have been reports of adipocytes arising from cultures of single myofibers (Shefer et al., 2004). Although these results have been interpreted to mean that APs arise from skeletal muscle satellite cells, single myofiber cultures are well known to contain capillaries and associated perivascular cells. Thus, the generation of adipocytes from a non-myogenic cell in these experiments cannot be formally excluded. Furthermore, recent lineage tracing studies using Myf5 reporter mice revealed that white adipocytes do not share a common lineage with skeletal muscle (Seale et al., 2008). It is therefore highly unlikely that adipocytes arise from myogenic cells.  Regardless of their source, several lines of evidence suggest the existence of APs in non-adipose tissues. Yet, how these cells arise in non-adipose tissues, what regulates their activation and differentiation, and what their functions are, remains unclear. Likewise, whether they share developmental relationships with adipose tissue APs and whether adipocytes in non-adipose tissues can affect total body metabolism is unknown. The answer to these questions requires direct experimentation on APs, and will likely have important implications on the fields of aging, chronic tissue disease, and metabolism.   23 1.5. OBJECTIVES OF THIS WORK Identifying the cells responsible for generating adipocytes is the major objective of this work. Since mature adipocytes are post-mitotic, and since not all cells can become adipocytes (as different tissues and fat depots each have a specific capacity for de novo adipogenesis), we proposed that new adipocytes arise from resident adipogenic stem/progenitor cells. The stem/progenitor cell model for tissue homeostasis posits that a tissue’s complement of functioning mature cells is maintained by a population of stem/progenitors, which can undergo differentiation through discrete hierarchical steps to generate new mature cells, as required. The distinction between stem and progenitor cell is based on the ability to self-renew. Stem cells are capable of self-renewal and therefore can generate new stem cells to allow life-long tissue homeostasis, whereas progenitors can only generate new mature cells and thus have a finite capacity for tissue maintenance. Tissues that are well-characterized by the stem/progenitor model in vivo include blood, skeletal muscle and the epithelial tissues of the skin and gut (Cohn et al., 1991; Kuang et al., 2007; Sherwood et al., 2004; Spangrude et al., 1988; Szilvassy et al., 1989; Tumbar et al., 2004).  The confirmation of this model in vivo requires stringent experimentation on a relatively pure cell population, which typically involves cell purification using surface antigen-based methods such as magnetic beads or fluorescence-activated cell sorting (FACS). To date, very few attempts have been made to isolate adipogenic cells using these methods, as the vast majority of studies in the field rely on either  24 immortalized cell lines or primary SV cells isolated from adipose preparations, called pre-adipocytes (Rosen and Spiegelman, 2000). Although many studies rely on bulk culture and transplantation studies to characterize the lineage potential of their test population, clonal analysis is necessary to meet the requirements of the stem/progenitor cell hypothesis. However, clonal analysis in certain cell populations can be challenging, and the results may be obscured by the requirement for extensive cell propagation and cultivation. Thus, careful statistical quantification of the activities of purified, minimally manipulated cells, both in vitro and in vivo are expected. These include limiting dilution analysis and quantitative (ideally competitive) transplantation assays to determine the frequency of stem/progenitor activity, and serial transplantation studies to determine the extent of self-renewal capacity. Although these types of studies typically require many years of experimentation, they are the gold standard for confirmation of the stem/progenitor model for tissue homeostasis. Importantly, they require the identification and purification of a putative stem/progenitor cell population.  Thus, in this work, my colleagues and I sought to develop a strategy to identify and characterize adipogenic progenitors, and to reveal new insights into their function. In our first study, our main objective was to purify APs from visceral and subcutaneous adipose depots using FACS and carefully quantify their activities to reveal whether depot-specific differences in APs might underlie differences in a depot’s growth modality. Next, we asked whether we could detect AP contribution to de novo adipogenesis in vivo using DNA-labeling strategies, and whether their contribution  25 may be regulated by dietary factors. Given that nothing is known about depot specific differences in adipogenic progenitors, or the regulators modulating their contribution to adipocyte hyperplasia, we believe our work will provide valuable insight to the field, and potentially lead to the development of strategies to stimulate or suppress their activity for therapeutic and experimental purposes.  In our second study, we sought to identify the cells responsible for adipocyte infiltration into skeletal muscle, as a first step towards understanding the differences between adipose and non-adipose AP cells. Furthermore, since fibrosis and adipogenesis are invariably linked in diseased muscle, we asked whether both tissue types may arise from the same cell, and whether they may develop from myogenic cells. Finally, since adipocyte infiltration accompanies chronic muscle injury, it is proposed that adipogenic progenitors exist in skeletal muscle, yet the reasons are unclear. Thus, we sought to determine whether they might play a functional role during muscle injury. We hope that this work will provide a novel appreciation for the cells responsible for de novo adipogenesis in skeletal muscle, and assign to them a physiological function that is independent of adipogenic differentiation.  While adipocyte hyperplasia has been observed adult tissues, the existence of stem/progenitor cells responsible for adipocyte homeostasis and expansion has not been formally shown. The identification and isolation of adipocyte progenitor cells provides the technology to enable careful interrogation of the ontogeny, function and  26 maintenance of adipocytes, in a tissue- and depot-specific fashion. As adipocytes have been ascribed to a multitude of functions and are found all throughout the body, the implications of this work are far-reaching, allowing for future work to span the fields of metabolism, development, aging, musculo-skeletal disease, regenerative medicine and stem cell biology. The work described in this thesis presents a valuable step towards our complete understanding of the adipocyte lineage.    27 1.6. TABLES Table 1.1. Factors secreted by adipocytes.   Factor Effect of adipocyte hypertrophy on factor release  Functions  Adiponectin ↓ Increases insulin sensitivity and glucose tolerance; anti-atherogenic Leptin ↑ Anorexic; an indicator of total body energy balance and adiposity; affects many neuroendocrine pathways Resistin ↑ Reduces insulin sensitivity and glucose tolerance; pro-inflammatory Angiotensinogen ↑ Increases vascular resistance; promotes new adipocyte production TNFa ↑ Induces insulin resistance; pro-inflammatory; source is unclear (macrophages/ stromal cells/ adipocyes) Plasminogen activator inhibitor-1 ↑ Reduces insulin sensitivity and glucose tolerance; pro-thrombotic; pro-atherogenic; production induced by TNF-a   28 1.7. REFERENCES Agarwal, A.K., and Garg, A. (2006). Genetic disorders of adipose tissue development, differentiation, and death. Annu Rev Genomics Hum Genet 7, 175-199. Anderson, N., and Borlak, J. (2008). Molecular mechanisms and therapeutic targets in steatosis and steatohepatitis. 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Proc Natl Acad Sci U S A 97, 1784-1789.      40 2. DEPOT-SPECIFIC DIFFERENCES IN ADIPOGENIC PROGENITOR ABUNDANCE AND PROLIFERATIVE RESPONSE TO HIGH FAT DIET1 2.1. INTRODUCTION Obesity, characterized by white adipose tissue (fat) expansion, significantly contributes to disease and mortality worldwide (Belanger-Ducharme and Tremblay, 2005; Roth et al., 2004). Fat is chiefly comprised of adipocytes, which represent the smallest functional unit of fat tissue and must change in size and/or number to change fat tissue mass (Hausman et al., 2001). Although overall clinical obesity is linked to metabolic diseases such as type 2 diabetes and dyslipidemia, it is increasingly clear that greater metabolic risks are specifically attributed to the expansion of the visceral fat depot (Wang et al., 2005). Recent evidence suggests that intrinsic characteristics of visceral and subcutaneous fat depots, rather than their anatomical position, may play a role in determining their different effects on metabolism (Gesta et al., 2007; Tran et al., 2008). We hypothesized that one such characteristic may be adipocyte size, which has been proposed as a major determinant of adipocyte function and secretory profile (Bluher et al., 2002; Khan et al., 2009; Rajala and Scherer, 2003).   1A version of this chapter has been provisionally accepted for publication. Joe, A.W.B., Yi, L., Even, Y., Vogl, A.W., Rossi, F.M.V. Depot specific differences in adipogenic progenitor abundance and proliferative response to high fat diet. Stem Cells.  41 In obese individuals increased adipocyte size, particularly in visceral depots, is associated with poorer metabolic parameters, adipose tissue dysfunction and inflammation, and adipocyte death- all of which may attributable to adipocyte stress and systemic lipotoxicity as adipocytes become grossly engorged with lipid (Khan et al., 2009; Livingston et al., 1972; Rajala and Scherer, 2003). In contrast subcutaneous adipocyte hyperplasia, which commonly occurs in type-2 diabetics treated with thiazolidinediones, is associated with improved insulin sensitivity and metabolic profiles (Spiegelman, 1998). Furthermore in ob/ob mice over-expressing adiponectin, a model of severe obesity due mainly to adipocyte hyperplasia, metabolic parameters were much healthier compared to normal ob/ob mice and were similar to that of wild-type animals (Kim et al., 2007). Thus, whether fat mass expands by adipocyte hypertrophy (resulting in large adipocytes) or hyperplasia (resulting in small adipocytes) might ultimately determine a depot’s effect on the development of metabolic disease.  New adipocytes arise from preadipocytes, or adipogenic progenitor (AP) cells, as mature adipocytes are post-mitotic. AP cell abundance and activity, together with the propensity for mature adipocytes to undergo hypertrophy, determine a fat depot’s ability to expand (Gray and Vidal-Puig, 2007). Thus we tested the hypothesis that APs can be prospectively isolated from adipose tissue, and represent a cell population that determines a depot’s ability to generate new adipocytes. Here, we show differences in the mechanisms underlying the expansion of visceral (VAT) and subcutaneous fat (SAT) after exposure to high fat diet, with VAT expanding  42 predominantly by adipocyte hypertrophy, and SAT by adipocyte hyperplasia. To explore whether these differences can be explained by depot-specific characteristics of APs we developed a technique to prospectively identify progenitors from both depots, and show that APs are 8-fold more abundant in SAT than VAT. Furthermore, in vivo BrdU experiments revealed a proliferative response to high fat diet in SAT- resident, but not in VAT-resident APs. Therefore, we propose that AP abundance and activity contribute to a depot’s ability to maintain a pool of small, functional new adipocytes, which may help to prevent the development of metabolic disease associated with obesity.   43 2.2. RESULTS AND DISCUSSION 2.2.1. Different mechanisms underlie SAT and VAT expansion To determine whether subcutaneous and visceral adipose tissue expand via different mechanisms in response to excessive caloric intake, we examined adipocyte size in both depots harvested from male littermate C57BL6/J mice fed either high fat diet (HFD) or standard diet (SD) (n=4 total, 2 per group; Figure 1a-d). High fat diet resulted in an average increase in adipocyte diameter of only 17% in SAT (from 35.2±13.5µm [mean ± SD] to 41.1±19.1µm, n >725), compared to 65% in VAT (from 51.1±15.3µm to 84.2±20.3µm, n> 290) (Figure 2.1 A-D). Thus, since we found a comparable fold increase in the mass of both depots (Figure 2.1E), adipocyte hyperplasia must play a greater role in SAT than VAT expansion. Indeed, histology of SAT from HFD-fed animals revealed regions containing several small adipocytes surrounded by hypertrophic cells, which was never observed in VAT (Figure 2.1 A, B). To further test the hypothesis that more new adipocytes are generated in SAT than in VAT, we exposed a group of mice to high fat diet for 60 days and treated them with BrdU for the final 10 days. Using confocal microscopy and immunohistochemistry, we observed a 148% and 54% increase in BrdU+ adipocyte-associated nuclei of SAT and VAT respectively, in response to HFD (Fig 2.1 F, G). Immunoelectron microscopy against BrdU confirmed labeling in the nuclei of mature vacuolated adipocytes, providing unequivocal evidence that new adipocyte generation had occurred (Appendix A, Figure A1). These results support  44 that greater production of new adipocytes occurs in subcutaneous than visceral fat during HFD-induced adipose tissue expansion.  2.2.2. Prospective identification of adipogenic progenitors In any given tissue, an increase in mature cells (hyperplasia) results from either decreased cell death or increased production from progenitors. Since adipocyte death increases during HFD-induced obesity (Strissel et al., 2007), we sought to directly test the hypothesis that increased progenitor activity is a hallmark of HFD- induced adipocyte hyperplasia. To compare the numbers and proliferative potential of adipogenic progenitors (AP), we developed a fluorescence-activated cell sorting (FACS)-based strategy to isolate murine adipocyte progenitor cells from stromovascular (SV) preparations of SAT and VAT (Figure 2.2; Appendix A, Figure A2). Stromovascular cells are defined as the non-adipocyte containing fraction of enzymatically and mechanically digested adipose tissue. SV cells sorted based on expression of CD45 (hematopoietic), CD31 (endothelial) and α7 integrin (smooth/skeletal muscle) (Sacco et al., 2008) failed to generate adipocytes in vitro and were therefore excluded from further analysis. The CD45- CD31- α7- subset (lin-) was further fractionated based on the expression of Sca-1 and CD34 and cultured for 14 days (Figure 2.2 A, B). From both depots, spontaneous generation of multilocular adipocytes was detected exclusively in cultures of Sca-1+ CD34+ cells. Large, unilocular perilipin-positive adipocytes were observed after one additional week in differentiation conditions (Figure 2.2 C, D). To test the hypothesis that lin- Sca-1+ CD34+ staining specifically identifies all the APs present in stromovascular  45 preparations of SAT and VAT, we performed limiting dilution analysis (LDA) on all four subsets of lin- Sca-1/CD34 expressing cells: Sca-1+CD34+ (R1), Sca-1-CD34+ (R2), Sca-1-CD34- (R3), and Sca-1+CD34- (R4) (Figure 2.3 A-D, Table 2.1). For each of these populations a number of cells ranging from one to 100 was sorted directly into replicate wells of 96 well plates and cultured in growth media for 14 days. To maximize adipocyte differentiation and to reveal latent adipogenic potential in all subsets, LDA cultures were exposed to a differentiation cocktail containing insulin, dexamethasone, isobutylmethylxanthine and troglitazone for an additional week. After 3 weeks we scored individual wells for the presence of colonies (>8 cells) and of perilipin-positive adipocytes (Figure 2.3E). We found that lin- Sca- 1+CD34+ from both SAT and VAT contained the vast majority of clonogenic activity (>94% and >72%, respectively) (Figure 2.3 B, D; Table 2.1). Importantly, even after exposure of all subsets to strong pro-adipogenic conditions, virtually all of the adipogenic activity from both depots (>97% in SAT, >99% in VAT) was detected exclusively in the Sca-1+CD34+ subset, confirming that these markers essentially identify all detectable adipogenic progenitors. When lin- Sca-1+ CD34+ cells were purified from the SAT or VAT of green fluorescent protein (GFP) positive mice and transplanted subcutaneously into unconditioned syngeneic recipients lacking GFP, pockets of GFP+ uni- and multilocular cells expressing the mature marker perilipin were detected in all recipients (n=3 for each depot, Figure 2.4). In support of our purification strategy, others have independently obtained and recently reported similar results from the subcutaneous depot (Rodeheffer et al., 2008) and from SV cells pooled from multiple depots (Tang et al., 2008).  46  2.2.3. APs are more abundant in subcutaneous than visceral fat Since APs are responsible for new adipocyte production during fat tissue growth (Rodeheffer et al., 2008) (Tang et al., 2008), depots that tend to grow by hyperplasia should be associated with higher progenitor numbers and/or activity than those that expand by hypertrophy. Thus, we used limiting dilution analysis to test the hypothesis that adipogenic progenitors are more abundant in SAT than VAT. Our data revealed significant depot-specific differences in the frequency of both clonogenic and adipogenic cells, with SAT-derived lin- Sca-1+ CD34+ cell (S-AP) clonogenicity and adipogenicity at 1 in 9.7-19.2 and 1 in 38.7-49.1 sorted cells, respectively, whereas VAT-derived lin- Sca-1+ CD34+ cell (V-AP) clonogenicity and adipogenicity occurred at 1 in 50.7-62.2 and 1 in 921.5-989.6 sorted cells, respectively (Table 2.1). Furthermore, results from single cell deposition studies were consistent with LDA calculations, confirming that LDA results were not confounded by paracrine signals between seeded cells (Appendix A, Table A1). When numbers of colony-forming adipogenic progenitors (CFU-AP) were calculated in relation to total numbers of lin- SV cells, we found that they were 8-fold more abundant in SAT than VAT (3258±1260 [mean ± SEM] and 406±70 per 106 lin- cells, respectively) (Figure 2.3F).  Finally, in agreement with reports that the stromal fraction from SAT has greater adipogenic potential compared to that from VAT (Adams et al., 1997; Tchkonia et al., 2002), S-AP cultures reliably generated more adipogenic regions and greater numbers of adipocytes than V-AP cultures of  47 comparable density (Figure 2.2 C, D). Altogether, these results suggest that APs are more abundant in subcutaneous fat than visceral fat.  2.2.4. AP proliferation in vivo is highly correlated with SAT, but not VAT expansion Recently, calculations based on atmospheric 14C incorporation into adipocyte genomic DNA suggested that new adipocytes are produced at a constant rate throughout life (Spalding et al., 2008). To date however, no direct analysis of the proliferation of cells fated to become adipocytes has been performed in vivo. Our strategy to prospectively identify APs enabled us to examine their contribution to adipose hyperplasia in healthy adult animals, and thus to directly test the hypothesis that the rate of AP proliferation is modulated by dietary inputs and is a determinant of hyperplastic fat tissue growth. We used a combination in vivo BrdU labeling and flow cytometry to quantify the proliferation of endogenous lin- Sca-1+ CD34+ cells (Appendix A, Figure A3). Animals subjected to either HFD or SD for 60 days (n= 16 per group) received BrdU for the last ten days of this period. We observed a significant linear correlation between the proportion of BrdU+ lin- Sca-1+ CD34+ cells and the weight of the individual depot from which they were extracted, in SAT (R2=0.73, p=<0.0001), but not VAT (R2=0.16, p=0.12) (Figure 2.5 A, B). These data suggest a tight relationship between the rate of AP proliferation and the extent of SAT, but not VAT, expansion. No correlation was found when the analysis was extended to a proliferating non-adipogenic population from either depot (R3 from  48 Figure 2.3 A, C; Appendix A Figure A4), confirming that this relationship between progenitor proliferation and depot growth is specific to adipogenic cells.  2.2.5. Subcutaneous depot APs proliferate in response to high fat diet Since AP proliferation is correlated with the extent of depot expansion in SAT, but not VAT, excessive caloric intake should selectively increase S-AP, but not V-AP, activity. We investigated whether AP proliferation can be modulated by diet in a depot-specific manner, by comparing BrdU labeling of endogenous lin- Sca-1+ CD34+ cells from depots taken from standard diet (SD)-fed and HFD-fed littermates. To avoid confounds due to a described bimodal response to high fat diet in inbred C57BL6/J mice, HFD non-responders were identified based on the method reported by Enriori et al (Enriori et al., 2007). Consistent with our hypothesis, we found a significant increase in the frequency of BrdU+ lin- Sca-1+ CD34+ cells in HFD- responders compared to SD-fed littermates in SAT (p=0.01, Figure 2.5C). In contrast, only a minor increasing trend was observed in samples from VAT (p=0.22, Figure 2.5D), confirming that caloric intake preferentially modulates the proliferation of subcutaneous progenitors over that of visceral progenitors.  2.2.6. Concluding remarks The ability of an organism to store energy is determined by the expandability of its adipose tissue. With respects to metabolic health, it is preferable to expand by hyperplasia to maintain a pool of functional adipocytes. However, during times of  49 large caloric excess, adipose tissue is challenged to accommodate extra lipid, which after chronic exposure can lead to adipocyte hypertrophy and metabolic dysfunction (Vazquez-Vela et al., 2008). Hypertrophy-induced adipocyte dysfunction leads to alterations in glucose homeostasis, changes to the adipocyte secretory profile, and systemic lipotoxicity, resulting in the pathogenesis of metabolic disease (Khan et al., 2009; Livingston et al., 1972; Rajala and Scherer, 2003). Significant differences in the effects of subcutaneous and visceral fat depots on metabolic heath have been described, and while whole depot transplantation experiments have made it clear that such differences are due to intrinsic properties of each depots (Tran et al., 2008), how these properties arise is as yet unclear.  Here we investigated the mechanisms underlying expansion of subcutaneous and visceral fat depots. We show that intrinsic differences in progenitor abundance and activity underlie an higher propensity of SAT versus VAT to grow by new adipocyte production, thereby reducing the fraction of hypertrophic mature adipocytes it contains. Whether production of new adipocytes is constant throughout adult life or it is modulated by changes in caloric intake is highly controversial (Spalding et al., 2008). Our work helps clarify this conundrum by showing that high fat diet leads to significantly increased proliferation of subcutaneous, but not visceral APs. Thus studies focused on the subcutaneous depot support a model in which new adipocyte production can be modulated in adults, while studies of the visceral depot would lead one to believe that it is constant.   50 In summary, we propose that the ability of a depot to expand by hyperplasia determines whether a depot will remain functional, and that a relative lack of progenitor cell activity may explain why depots such as VAT accumulate hypertrophic, dysfunctional adipocytes and are therefore associated with higher risk of disease. It remains to be seen whether differences in AP activity are due to depot- specific extracellular cues or to progenitor cell-autonomous characteristics, and what their impact may be on the risks of disease associated with each depot. Regardless, our work underscores a novel intersection between stem cell biology and metabolism, and will have many important implications on the fields of adipose tissue homeostasis, remodeling, turnover and transplantation.   51 2.3. EXPERIMENTAL PROCEDURES 2.3.1. Animals Mice were maintained in an enclosed, pathogen-free facility, and experiments were performed in accordance with University of British Columbia Animal Care Committee regulations. C57BL/6 mice (>6 weeks) were used in this study. C57BL/6-CMV- βActin-GFP transgenic mice were a generous gift from I. Weissman, Stanford University. For high fat diet experiments, animals were provided Research Diets #D12451 ad libitum (20 kcal% protein [casein and L-cystine], 35kcal% carbohydrate [corn starch, maltodextrin 10 and sucrose], 45kcal% fat [soybean oil and lard]). All mice were bred in-house.  2.3.2. Tissue preparation For all experiments, subcutaneous adipose tissue was obtained from the inguinal fat pad, and visceral adipose tissue was obtained from the perigonadal fat pad. Tissues were minced with scissors until homogeneous. 1mL CollagenaseD (Roche Biochemicals, 1.5u/ml) with Dispase II (Roche Biochemicals, 2.4u/ml), supplemented in total 10mM CaCl2, was applied to each gram of homogenized tissue. The preparation was incubated at 37 °C for 2 hours, washed in phosphate buffered saline (PBS) supplemented with 2% fetal bovine serum (FBS) and 2mM EDTA, and then centrifuged at 200rcf for 10 minutes. The floating adipocyte layer and supernatant were removed, and the remaining stromovascular cell pellet was  52 resuspended in supplemented PBS, and then passed through a 40um cell strainer (Becton Dickenson [BD]) prior to staining for flow cytometry.  2.3.3. Antibodies and flow cytometry / FACS Stromovascular cells were incubated with primary antibodies for 30min at 4C in supplemented PBS containing 2mM EDTA and 2% FBS, at ~3x107 cells per ml. We used the following monoclonal primary antibodies: anti-CD31 (clones MEC13.3 [BD] and 390 [Cedarlane Laboratories]), anti-CD34 (clone RAM34 [eBiosciences]), anti- CD45 (clone 30-F11 [BD]), anti-CD45.1 (clone A20 [BD]), anti-CD45.2 (clone 104 [eBiosciences]), anti-Sca1 (clone D7 [eBiosciences]) and anti-α7integrin (clone CA5.5 [produced in-house]). Where necessary, biotinylated primary antibodies were detected using Streptavidin coupled to either phycoerythrin, allophycocyanin, or fluorescein-isothiocyanate (Caltag). Cells were stained with propidium iodide (1µg/ml) and Hoechst 33342 (2.5µg/ml) and resuspended at ~1x107 cells/ml immediately prior to sorting or analysis. Analysis was performed on LSRII (BD) equipped with 3 lasers. Data was collected FacsDIVA software. Sorts were performed on a FACS Vantage SE (BD) or FACS Aria (BD), both equipped with 3 lasers, using a 100µm nozzle at 18psi to minimize the effects of pressure on the cells. Sorting gates were strictly defined based on machine calibration using single-color antibody staining and isotype control (fluorescence minus one) experiments. Biexponential analysis was performed using FlowJo 8.7 (Treestar) software.   53 2.3.4. Cell culture Cells were grown in high-glucose Dulbecco’s modified eagle medium (DMEM), supplemented with 2.5ng/ml bFGF (Invitrogen) and 20% FBS. This media is referred to in text as ‘growth media’. Cells were seeded in tissue-culture treated plates coated with Collagen type 1 (Sigma). After sorting, cells were allowed to adhere for up to 3 days, after which ½ the medium was changed. Media was changed every 2- 4 days. For adipogenic differentiation, we supplemented bFGF-free growth media with 0.25µM dexamethasone, 0.5mM Isobutylmethylxanthine, 1µg/ml insulin, 5µM troglitazone.  2.3.5. Limiting dilution analysis One to one hundred cells from each Sca-1/CD34 expressing population (R1-R4) were sorted into individual wells of a collagen-coated 96 well plate directly from the sorter. Cells were grown as described in the text. After three weeks, cultures were fixed and stained for perilipin and nuclei. Wells were scored for the presence of colonies (>8 cells) and adipocytes (perilipin-positive cells). A minimum of 30 replicate wells was generated for each cell dose. LDA calculations were based on the single hit Poisson model (see Statistics). Frequency of CFU-APs was determined from wells that scored positive for both colony-forming and adipocyte- forming ability. We determined CFU-AP content in each depot by multiplying the frequency of CFU-APs by the frequency of R1 (lin- Sca-1+ CD34+) cells, and normalizing to 106 lin- cells.   54 2.3.6. Transplantation Host mice (wild type) were anesthetized using isoflurane in accordance with University of British Columbia Animal Care Committee guidelines. Donor cells were isolated from GFP+ mice. Cells were sorted into cold DMEM and collected by centrifugation at 450rcf for 5minutes. Cells were resuspended in 25µl Matrigel and loaded into an ice-cold needle and syringe immediately prior to injection. Cells were injected subcutaneously into the sub-scapular region, with control cells injected on the contralateral side. Tissues were harvested after 3 weeks.  2.3.7. Histology and imaging Tissues were harvested from animals that were perfused transcardially with 20ml PBS/ 2mM EDTA, followed by 20ml 4% paraformaldehyde (PFA). Tissues were processed for cryosectioning or paraffin embedding using standard methods. Cell cultures were fixed in 4% PFA in PBS for 10-20min prior to staining. Immunostaining was performed using monoclonal antibodies against perilipin (Sigma) or BrdU (BD). Briefly, samples were permeabilized in 0.3% Triton-X-100 (Sigma) in PBS, and blocked for 2hrs at room temperature in PBS containing 25% normal goat serum (NGS), 0.3% Triton-X-100, 3% bovine serum albumin (BSA) and 0.1%NaN3. Cells were stained overnight at 4C using antibody diluted in 10% NGS, 0.3% Triton-X-100, 3% BSA and 0.1% NaN3. Primary antibody was detected using goat anti-mouse IgG antibodies conjugated to Alexa 568 or 488 (Molecular Probes), or using the anti- Mouse IgG HRP detection kit (BD).  55 Cells were visualized using a microscope (Axiovert 200 for inverted microscopy, Axioplan2 for conventional microscopy; Carl Zeiss Microimaging) and images were acquired using a charge-coupled device camera (Retiga Exi [Axiovert 200] or Retiga Ex [Axioplan2], qImaging) and OpenLab4 software (Improvision). Adipocyte diameters were determined by using OpenLab. To confirm the specificity of the GFP signal in transplantation experiments, we compared the signal to non-specific autofluorescent background in all other channels (Brazelton and Blau, 2005). Confocal microscopy was performed using a Nikon C1 laser scanning confocal microscope equipped with lasers at 488nm, 568nm, and 633nm. Images were captured using the least exposure time possible, and manipulation of brightness and contrast, coloring adjustments and assembly into figures were performed using ImageJ, OpenLab4, Adobe Illustrator CS3 and Adobe Photoshop CS3 .  2.3.8. In vivo BrdU labeling studies BrdU was administered from days 50-60 of treatment diet in drinking water (0.8g/L in 2% sucrose) and by intraperitoneal injection (100mg/kg) once every 2 days. For flow cytometric analysis, cells were prepared and stained for surface markers as described above. Samples were then fixed in 2% PFA for 20 min, and membranes were permeablized in 0.2% saponin for 10 min. Cells were treated with 30µg DNAse for 1h at 37 °C, and 1:25 dilution of anti-BrdU (clone PRB-1 [Invitrogen]) was applied to the samples for 30min at room temperature. For light microscopy analysis, we prepared 5µm sections from paraffin-embedded adipose tissue. After rehydration and antigen retrieval in citric acid buffer (1.4mM citric acid, 8.7mM Na citrate,  56 pH6.0), slides were treated in increasing concentration of HCl (1, 2 and 4N) for 10min each. Acid was neutralized in borate buffer (80mM Na borate decahydrate, 80mM boric acid, pH 8.5). Slides were stained as described above for immunostaining using anti-BrdU (clone B44 [BD]) at 1:40 dilution. For electron microscopy analysis, tissues were fixed overnight in 3%PFA, then dehydrated through an increasing concentration series of cold (0 to –20oC) ethyl alcohols and embedded in Unicryl. Thin sections were cut using an ultramicrotome and collected on nickel formvar/carbon coated grids. Grids were incubated overnight at 4C in anti- BrdU (clone B44, 0.005 mg/ml in PBS containing 0.05% Tween 20, 0.1% bovine serum albumin [TPBS-BSA], and 1% normal goat serum). Grids were washed and then incubated for 1 hr at room temperature in secondary antibody conjugated to 10- nm colloidal gold diluted 1:25 in TPBS-BSA containing 5% fetal bovine serum. Grids were fixed in 0.1% glutaraldehyde, stained with 1% aqueous uranyl acetate, and then air-dried. Staining was evaluated and photographed on a Philips 300 electron microscope operated at 60 kV.  2.3.9. Statistics Statistical tests were performed using Prism 4 (GraphPad Software). Analysis of limiting dilution data was performed using a web application made available by the Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia (http://bioinf.wehi.edu.au/software/limdil/) (Shackleton et al., 2006). This software tests departures from the single-hit Poisson model using a generalized linear model.  57 2.4. TABLES Table 2.1. Limiting dilution analysis of Sca-1/CD34 expressing populations from SAT and VAT.  Experiments were performed as described in the text. In populations exhibiting clonogenic or adipogenic activity, data was generated from three different sorts, using a minimum of 6 animals each. # of wells indicates the total number of wells analyzed in each sort.  58      Tissue Population Sort # of Wells Clonogenic frequency 1/X (95% CI) Adipogenic frequency 1/X (95% CI)  Subcutaneous Fat R1 (AP) 1 459 9.72 (8.31-22.39) 38.67 (30.12-49.68)   2 480 18.39 (15.42-21.94) 38.49 (30.00-49.44)   3 470 19.21 (16.03-23.05) 49.08 (38.03-63.39)   R2 1 349 562.95 (326.25-971.65) 2538.61 (818.96-7871.43)   2 291 393.45 (198.01-782.29) 3340.25 (474.53- 23530.50)   3 344 143.93 (105.48-196.48) 772.75 (401.39-1488.13)   R3 1 180 infinity infinity   2 60 infinity infinity   R4 1 180 infinity infinity   2 180 infinity infinity  Visceral Fat R1 (AP) 1 460 62.23 (46.72-82.96) 989.62 (319.59-3066.51)   2 340 50.72 (38.10-67.58) 942.11 (300.02-2877.43)   3 480 50.85 (41.45-62.39) 921.54 (480.37-1768.31)   R2 1 290 195.01 (126.96-299.69) Infinity   2 252 685.95 (285.18-1650.93) Infinity   3 253 870.89 (362.65-2092.41) Infinity   R3 1 180 4360.31 (616.35- 30865.11) Infinity   2 180 85.99 (61.27-120.76) 4360.31 (616.35- 30865.11)   3 180 853.06 (353.77-2058.00) Infinity   R4 1 170 infinity infinity   2 180 infinity infinity  59 2.5. FIGURES Figure 2.1. Different mechanisms underlie subcutaneous and visceral fat expansion after exposure to HFD  (A-D) HFD led to a significantly smaller increase in adipocyte size in SAT (A, C) than VAT (B, D). SAT and VAT were collected from male C57BL6/J littermates exposed to HFD or SD for 60 days (total n=4, 2 per group). Adipocyte diameters (C, D) were measured from 5 random fields of H&E stained paraffin sections from SAT (A, n>725) and VAT (B, n>290). Scale bars at 100µm. (E) HFD results in a similar increase in the fat mass of both depots. (F) Detection of new adipocyte production in vivo by BrdU labeling. Confocal immunofluorescence analysis was performed on paraffin sections taken from animals exposed BrdU over the last 10 days of the 60-day treatment diet protocol. Nuclei were stained using TOTO3, and autofluorescence from the green channel was used to detect tissue structures. A single optical section of SAT is shown. Scale bar at 50µm. (G) Quantification of BrdU+ adipocyte-associated nuclei in SAT and VAT sections. Data was normalized to the mean of the control animals (n=4 per group).  60         61 Figure 2.2. Prospective isolation of adipogenic cells from SAT and VAT  (A, B) Viable cells from stromovascular (SV) fractions of SAT (A) and VAT (B) were identified based on forward/side scatter, medium intensity Hoechst staining (to exclude debris) and low intensity PI staining (to exclude dead or dying cells). Markers for hematopoietic (CD45+), endothelial (CD31+) and smooth/skeletal muscle (α7 integrin+) cells were used to exclude mature lineages from analysis (resulting cells called lin-). (C, D) lin- Sca-1+ CD34+ cells sorted from SAT (C) and VAT (D) were cultured for 14 days in growth medium, switched to adipogenic conditions for 7 days, then stained using a purified mouse monoclonal antibody against perilipin. Scale bars at 100µm.  62    63 Figure 2.3. Quantification of adipogenic progenitors using limiting dilution analysis  (A, C) Four distinct Sca-1/CD34 expressing populations were identified in lin- SV cells from SAT (A) and VAT (C) using flow cytometry. (B, D) R1 (Sca-1+CD34+) contains the vast majority of clonogenic and adipogenic potential from both SAT (B) and VAT (D). Clonogenicity (left) and adipogenicity (right) of Sca-1/CD34 expressing populations from both depots was determined using limiting dilution analysis. Dashed lines represent 95% confidence intervals for regression lines. Complete LDA data is shown in Table 2.1. (E) Morphology of adipocyte colonies detected in lin- R1 LDA cultures. Perilipin positivity identifies mature adipocytes. Scale bar at 100µm. (F) Relative content of CFU-APs in the lin- stromovascular fraction of SAT and VAT (n=12).  64       65 Figure 2.4. Transplanted lin- Sca-1+ CD34+ cells produce new adipocytes in wildtype, syngeneic animals  APs were isolated from transgenic mice ubiquitously expressing GFP. Immediately after sorting, 105 GFP+ APs from either SAT or VAT were suspended in 25µl matrigel and transplanted by subcutaneous injection into the sub-scapular region of syngeneic, GFP negative mice (n=3 per group). Tissues were harvested after 3 weeks. Skeletal muscle is labeled “m”, and dermal structures labeled “d”. (A) Low magnification image of a pocket of donor derived GFP+ adipocytes located in the loose connective tissue. Scale bar at 500µm. (B, C) Higher magnification images of engrafted GFP+ SAT APs (B) and VAT APs (C). Scale bars at 100µm. (D) Maximum intensity projection of a confocal optical stack shows that adipocytes arising from transplanted GFP+ APs express the mature marker perilipin. Nuclei were detected using TOTO3. Scale bar at 500µm. (E) High magnification image of a single confocal optical section showing co- localization of GFP signal with perilipin, indicating that donor derived cells generated mature adipocytes in vivo. Scale bar at 50µm.  66      67 Figure 2.5. Greater adipogenic progenitor activity in subcutaneous compared to visceral fat  6-12 week old mice were placed on high fat diet (HFD) for 60 days, and administered BrdU (0.8mg/ml in drinking water and 100mg/kg IP every 2 days) during the last 20 days of HFD treatment. (A, B) In vivo BrdU studies in HFD-fed animals show a strong correlation between the proportion of BrdU+ lin- Sca-1+ CD34+ cells and the extent of SAT (A), but not VAT expansion (B). (C, D) Proliferation of lin- Sca-1+ CD34+ cells from SAT (C) but not from VAT (D) increases in response to HFD treatment. n=16 per group.      68 2.6. REFERENCES Adams, M., Montague, C.T., Prins, J.B., Holder, J.C., Smith, S.A., Sanders, L., Digby, J.E., Sewter, C.P., Lazar, M.A., Chatterjee, V.K., et al. 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Satellite cells, which are found juxtaposed to myofibers beneath the basal lamina, contribute to the growth and regeneration of muscle in the postnatal period (Mauro, 1961; Morgan and Partridge, 2003). Self-renewing satellite cells activate and proliferate in response to muscle injury, and promote regeneration by fusing with damaged myofibers or by producing new myofibres (Charge and Rudnicki, 2004; Collins et al., 2005; Dhawan and Rando, 2005; Kuang et al., 2007; Sacco et al., 2008). Progression of differentiation from quiescent satellite cell to mature post-mitotic myofiber proceeds through discrete cellular intermediates, collectively called myogenic progenitors (MP), which can be distinguished using molecular markers (Buckingham, 2006).  In vivo, this stepwise process is coordinated by specific extracellular signals. Following activation, satellite cells express the Notch ligand Delta1, and paracrine or  2 A version of this chapter has been submitted for publication. Joe, A.W.B., Yi, L., Natarajan, A., Le Grand, F., So, L., Wang, J., Rudnicki, M.A., Rossi, F.M.V. Muscle injury activates resident fibro/adipogenic progenitors that facilitate the terminal differentiation of myogenic cells.  72 autocrine engagement of Notch results in myoblast expansion, partly by direct inhibition of terminal differentiation (Conboy and Rando, 2002). Following myoblast expansion, pro-differentiation signals are required to offset Notch signaling and ensure that dividing myoblasts will differentiate to generate mature myofibers. The precise nature of these signals is the object of intense debate in the field, and several candidate molecules have been proposed for this role, including Wnt family members, IL-6 and IGFs (Bodine et al., 2001; Brack et al., 2007; Serrano et al., 2008), suggesting that multiple factors may be involved in promoting myogenic differentiation. Importantly, the sources of these factors within healing muscle tissue have yet to be determined.  In the context of healing muscle tissue, expanding myoblasts interact with inflammatory and stromal cells. It seems likely that these interactions are important in regulating their activity. The role of inflammatory cells in muscle regeneration has been recently explored (Arnold et al., 2007; Contreras-Shannon et al., 2007; Sonnet et al., 2006); however, nothing is known about the contributions of other tissue- resident populations.  Examining failed muscle regeneration provides clues into which cell types, other than the omnipresent endothelial cells, may be involved in the response to damage. Specifically, after chronic muscle injury and during the late stages of muscular dystrophies, muscle is replaced by a mix of fibrous tissue and white adipocytes in a process termed fatty degeneration. This fibro/adipogenic infiltration compromises  73 muscle function and alters the tissue environment potentially limiting the success of regenerative approaches (Lipton, 1979). Thus, the cells responsible for fibro/adipogenic infiltration likely comprise an important part of the physiological context in which satellite cells function.  It has been suggested that fibrocytes and adipocytes develop from myogenic cells due to alternative lineage choice, which takes place when myogenic differentiation is impaired by cell autonomous defects or by environmental changes associated with aging and disease (Brack et al., 2007; Li et al., 2004; Li and Huard, 2002; Shefer et al., 2004). However, these experiments have not been performed on prospectively identified cell populations. Thus, we decided to test the hypothesis that adipogenic and myogenic activities can arise from a single cell using prospective isolation and lineage tracing analysis. Here, we report the identification of distinct mesenchymal progenitor populations present in skeletal muscle, which can be distinguished from MPs using surface markers. We show that bipotent fibro/adipogenic progenitors (FAPs) represent an abundant population in healthy muscle, and that their developmental origins are different from those of myogenic progenitors. FAPs rapidly enter the cell cycle in response to acute muscle damage, suggesting a role for this population during regeneration. Indeed, we found that FAPs are a source of signals inducing the differentiation of primary myoblasts. Our results identify a novel population of fibro/adipogenic progenitors, distinct from myogenic cells, which proliferates in response to muscle injury to transiently establish an environment that enhances myogenic differentiation.  74 3.2. RESULTS 3.2.1. Purification of mesenchymal progenitors with distinct developmental potential from skeletal muscle Skeletal muscle is a complex tissue containing cells of multiple lineages. Among these, myogenic cells are the best characterized and markers allowing their isolation are known. To identify other progenitor lineages present in muscle we examined for expression of CD34, which is expressed by the large majority of satellite cells (Beauchamp et al., 2000), and Sca-1, which is expressed by progenitor populations of multiple tissues and whose expression on skeletal muscle progenitors is controversial (Mitchell et al., 2005; Polesskaya et al., 2003; Sherwood et al., 2004). We used multiparameter flow cytometry to analyze mononuclear cells from dissociated muscle (Figure 3.1; Appendix B, Figure B1). Although it has been suggested that hematopoietic (CD45+) and/or endothelial (CD31+) cells might be induced to acquire myogenic potential, their role in physiological muscle regeneration is unclear (De Angelis et al., 1999; Polesskaya et al., 2003). Therefore, we focused on the following CD45- CD31-  (lin-) cell populations: Sca-1- CD34-; Sca- 1- CD34+ and Sca-1+ CD34+ (Figure 3.1B). The proliferative and developmental potentials of cells sorted from undamaged muscle were analyzed in vitro (Figure 3.1 C-E, Figure 3.2) and in vivo following transplantation (Figure 3.3).  Double-sorted cells from each population were plated in conditions that allow differentiation along chondrogenic, osteogenic, adipogenic and myogenic lineages,  75 as these mesenchymal tissues are both functionally and anatomically associated with skeletal muscle. Lin- Sca-1- CD34- cells were able to generate mineralized bone nodules and alcian blue-positive cartilage (Figure 3.1C). No adipogenesis was detected, suggesting that these cells represent skeletal progenitors rather than mesenchymal stem cells. Within this subset, clonogenic cells were exceedingly rare, with colony formation observed at a frequency of 1 in 757 sorted cells (Appendix B, Figure B2). Due to their rarity and to the fact that cartilage and bone generation are not common outcomes of muscle healing, we excluded lin- Sca-1- CD34- cells from further analysis.  The lin- Sca-1+ CD34+ cell subset spontaneously and efficiently generated numerous adipocytes in all culture conditions, but failed to give rise to mineralized nodules or cartilage (Figure 3.1D; Appendix B, Figure B3). Adipogenic progenitors expressing identical markers were recently isolated from white adipose depots (Rodeheffer et al., 2008; Tang et al., 2008), suggesting that these cells may be present in multiple tissues. Lin- Sca-1+CD34+ cells were abundant, representing up to 15% of lin- cells present in young, undamaged muscle. Freshly sorted cells expressed genes typical of pre-adipocytes and after differentiation initiated the expression of mature adipocyte markers (Appendix B, Figure B4).  Finally, lin- Sca-1- CD34+ cells gave rise to cultures containing both differentiated myofibers and proliferating mononuclear cells, as expected (Montarras et al., 2005). Despite the presence of 20% fetal serum and bFGF in the growth medium,  76 conditions that efficiently inhibit the differentiation of pre-plating enriched primary myoblasts (Rando and Blau, 1994), sorted lin- Sca-1- CD34+ cells spontaneously generated myofibers in vitro (Figure 3.1E). Early in culture, these cells expressed genes typical of satellite cells, and after 2 weeks in growth media, they initiated expression of markers of terminal myogenic differentiation (Appendix B, Figure B4).  CD34 surface expression is rapidly lost following MP activation, preventing the use of this marker to sort myoblasts from damaged tissue (Beauchamp et al., 2000). However, cell subsets with identical properties to those described above can be isolated with an alternative marker combination based on α7 integrin (Blanco-Bose et al., 2001; Sacco et al., 2008). Indeed, we found that all myogenic activity in both undamaged and damaged muscle originates from lin- α7+ Sca-1- cells (Appendix B, Figure B5). Similarly, we found that lin- α7- Sca-1+ cells were as efficient as lin- Sca- 1+CD34+ cells at generating adipocytes in vitro (Appendix B, Figure B5). Thus, we have developed two strategies, using CD34 or α7 integrin, to prospectively identify identical myogenic progenitors and muscle-resident adipogenic cells, from both undamaged and damaged muscle (Figure 3.1F).  In summary, we identified three distinct lin- cell populations from skeletal muscle, each of which are endowed with the potential for generating specific mesenchymal lineages.   77 3.2.2. Single lin- α7- Sca-1+ cells produce both fibroblasts and adipocytes in vitro We used limiting dilution assays (LDA) to measure the frequency of adipogenic and myogenic progenitors, defined as cells capable of clonal expansion and terminal differentiation into adipocytes or myofibers, in sorted cell populations. We deposited between 1-100 double-sorted lin- α7+ Sca-1- MPs or lin- α7- Sca-1+ cells from undamaged muscle into individual wells of a 96-well plate, with a minimum of 30 replicate wells for each cell dose. After three weeks of culture in growth medium we evaluated each well for: 1) the presence of a colony (>10 cells); 2) the presence of myosin heavy chain (MyHC)-expressing myofibers; and 3) the presence of lipid- laden adipocytes.  We found that 1 in 16 (95% confidence interval (CI) 12.0-21.4) sorted MPs were capable of forming colonies, and that over 95% of these colonies contained MHC+ myofibers, suggesting that all clonogenic cells within this population are myogenic. Lin- α7- Sca-1+ cells formed colonies with similar efficiency (1 in 19; 95% CI 14.2- 22.3) but only 35% of these colonies contained adipocytes, while the remaining 65% contained cells retaining a fibroblastic morphology that did not express myogenic or adipogenic markers, but were positive for fibroblast markers ER-TR7 (Brack et al., 2007), FSP1 (Strutz et al., 1995) and α-smooth muscle actin (SMA) (Tomasek et al., 2002) (Figure 3.2). The frequency of adipogenic colonies did not significantly change even following exposure to strong pro-adipogenic conditions during the last week of culture (containing insulin, dexamethasone, isobutylmethylxanthine and  78 troglitazone). In an attempt to resolve adipogenic and fibrogenic activities we deposited freshly isolated, single lin- α7- Sca-1+ cells into individual wells, directly from the sorter. After three weeks in growth medium, we observed both perilipin- expressing adipocytes and SMA-expressing fibroblasts in single-cell derived colonies, suggesting that the muscle-resident lin- α7- Sca-1+ population contains bipotential progenitors capable of producing both adipocytes and fibroblasts (Figure 3.2C). This notion is consistent with the fact that adipocytic infiltration and fibrosis are invariably linked during fatty degeneration of diseased skeletal muscle. Hence, we will hereafter refer to these cells as fibro/adipogenic progenitors (FAPs).  3.2.3. FAP engraftment is dictated by the environment We tested the developmental potential of MPs and FAPs in vivo by transplanting freshly sorted cells into syngeneic recipients. For each subset, 5x104 cells were purified from GFP expressing donors and injected in the undamaged tibialis anterior (TA) muscle of recipient animals (Figure 3.3 A,B). As previously reported, MP cells efficiently engrafted by fusing to myofibers along the injection needle tract (Figure 3.3 B). In contrast, no donor cells were found in the muscles of animals injected with FAPs three weeks after transplant, suggesting that the environment of adult healthy muscle does not support FAP engraftment. In support of this notion, subcutaneous delivery of GFP+ FAPs cells resulted in the formation of clusters of donor-derived, lipid-laden, perilipin-expressing adipocytes (Figure 3.3 C, D). Together, these data suggest that the survival of FAPs is influenced by environmental signals, and that these signals are not present in healthy skeletal muscle. This is consistent with the  79 fact that fibro/adipocytic infiltration is not observed following healing of young healthy muscle, but only when myogenic regeneration fails due to aging or disease.  3.2.4. FAPs and MPs have distinct developmental potentials and do not arise from a common progenitor Our data indicates that FAPs and MPs represent distinct progenitor populations present in undamaged, young adult muscle and that they are each committed to a specific lineage. In some cases however, a latent myogenic potential is revealed when certain cell types are exposed to differentiating myogenic cells. Recently, adipogenic progenitors isolated from white fat depots were shown to fuse with differentiating C2C12 myoblasts in vitro (Rodeheffer et al., 2008). To test whether muscle derived FAPs can be recruited to the myogenic lineage in this context, we isolated them from GFP+ animals and cultured them together with freshly isolated GFP- MPs for ten days (Figure 3.4A; Appendix B, Figure B6). Immunostaining revealed that none of the GFP+ FAPs acquired expression of MyHC despite being in direct contact with terminally differentiated GFP- myotubes (Figure 3.4A). In fact, a subset of these GFP+ cells had clearly initiated lipid accumulation, confirming their commitment to adipogenic differentiation.  To directly test whether FAPs can respond to damage induced signals by generating myofibers in vivo, we injected 2x104 double-sorted MPs or FAPs freshly-isolated from mice ubiquitously expressing hPLAP, into the TA muscle of either undamaged or notexin-damaged wild-type recipients. After three weeks we detected donor-  80 derived myofibers in all animals receiving MPs, with approximately 4 fold increase in engraftment in the notexin-damaged group (Table 3.1). Conversely, no myofibers were detected in any of the animals receiving FAPs, from either undamaged or damaged groups (Table 3.1). Thus, FAPs do not differentiate along the myogenic lineage or fuse to differentiating myogenic cells in vitro or in vivo.  To further explore the relationship between these two cell types, we asked whether adipogenic cells express Myf5, a transcription factor found in the vast majority of satellite cells. MPs and FAPs cells were sorted from mice expressing nuclear LacZ under the control of Myf5 (Figure 3.4 B,C). As expected, the majority of MPs (60%) were positive for nLacZ staining. No LacZ positive cells were detected in the Sca-1+ CD34+ subset, confirming that FAPs do not express Myf5.  Together, our findings suggest that FAPs originate from a separate developmental source rather than deriving from myogenic cells in response to pathological alterations of the environment. To test this hypothesis directly, we bred knock–in mice carrying CRE recombinase under the control of Myf5 regulatory elements to mice expressing YFP in a CRE-dependent fashion. In this approach, transient Myf5 expression during development leads to the heritable activation of YFP transcription from the ROSA locus, allowing the identification of the progeny of the original Myf5 expressing cell (Figure 3.4D). The same strategy was recently used to show that brown fat and muscle share developmental origins (Seale et al., 2008). FACS analysis of skeletal muscle preparations from these mice revealed that 60% of MPs  81 expressed YFP (Figure 3.4E). Furthermore, the frequency of YFP+ MPs increased to >85% after 5 days of culture (Figure 3.4F). Conversely, >99% Sca-1+ CD34+ FAPs did not express YFP, and remained YFP- after culture (Figure 3.4 E,G). Thus, FAPs and MPs found in adult muscle are unlikely to arise from a common progenitor cell.  3.2.5. Fibro/adipogenic progenitors are rapidly induced to proliferate upon muscle damage In skeletal muscle, fibrosis and adipocyte infiltration in response to damage are only observed in aging or disease. Yet, FAPs are abundant in young, healthy muscle. To investigate a potential physiological role of FAPs in the regeneration of healthy muscle, we analyzed their response to acute damage. To induce damage we injected Notexin (NTX), a myotoxin which acts on terminally differentiated myofibers without damaging mononuclear progenitors (Harris, 2003; Harris et al., 2003). Four days after injection, all lin- α7 integrin+ cells had downregulated CD34 (not shown), confirming that muscle resident progenitors had been uniformly exposed to damage signals.  To determine whether FAPs respond to myofiber damage in vivo by entering the cell cycle we measured the rate at which they incorporate BrdU following NTX injection. BrdU was injected IP every 12 hours starting 24 hours prior to damage induction, and animals harvested at time 0 (no damage) and at 24-hour timepoints thereafter. Flow cytometric analysis revealed significant proliferation in both FAPs and MPs (Figure 3.5 A,B), with a greater frequency of FAPs than MPs incorporating BrdU at  82 early time points (Figure 3.5B). Thus, endogenous FAPs are efficiently activated by signals induced by myofiber damage.   To detect changes in proliferation of endogenous FAPs and MPs over time, we repeated the NTX damage experiment by administering a pulse of BrdU during the 24 hours immediately prior to harvest (Figure 3.5C). This strategy provides a snapshot of proliferation at 24 hours periods after damage, as opposed to measuring all cycling cells starting from time 0. Pulse BrdU data confirmed the observation that a higher fraction of FAPs compared to MPs proliferate during the first 72 hours after damage, and additionally revealed that MPs continue to proliferate after FAP BrdU incorporation has returned to baseline, between six and ten days after damage (Figure 3.5C).  A prediction that can be formulated based on these results is that if FAPs cycle more efficiently than MPs, then the ratio between the two cell types should change substantially during the first few days following damage induction. Indeed, at quiescence MPs are three times as abundant as FAPs (Figure 3.5D). Between 48 and 72 hours after damage FAPs dramatically increase, nearly equaling MPs in numbers. Between four and five days following damage the FAP to MP ratio rapidly returns to pre-damage levels despite the fact that we observed no differences between the proliferation rates of either population, suggesting that FAPs may be actively removed from the tissue at this stage of the healing process. Further work will be required to verify this hypothesis.  83  To explore whether increases in FAP proliferation and abundance correlate with an increase in fibro/adipogenic colony forming cells, we performed limiting dilution assays on cells sorted from TAs at 48hrs post-damage (Figure 3.5E). Consistent with our observation that MP proliferation is limited at this time point, the frequency of myogenic colony forming cells after damage was not significantly different from that obtained from undamaged muscle (Figure 3.5E, right). In contrast, and in good agreement with BrdU incorporation data, we detected a significant increase in the frequency of clonogenic cells within the FAP population 48hrs after damage (Figure 3.5E, left). Importantly, although we sorted our cells after damage, we did not observe any change in the ratio of colonies that generated adipocytes versus fibroblasts. In addition, did we not observe any MyHC positive structure in FAP- seeded wells, nor any perilipin positive adipocytes in MP seeded wells, confirming that despite the rapid induction of proliferation, muscle damage does not alter FAP cell fate or surface phenotype.  To gain further insight in the role of FAPs in regeneration, we investigated their location within the tissue using confocal microscopy. Published in vitro analysis of individual myofibers conclusively showed that adipogenic cells are associated with these structures in undamaged muscle (Shefer et al., 2004). Furthermore, adipogenic progenitors have been localized to the vasculature in fat depots, suggesting they may occupy a similar position in muscle (Tang et al., 2008). We used CD31 and Sca-1 staining to distinguish FAPs from other cells, as FACS  84 analysis indicated that only FAPs and a population of CD31+ cells are positive for Sca-1 in skeletal muscle (Figure 3.1 A, B) (Kafadar et al., 2009). Confocal microscopy identified abundant Sca-1+ mononuclear cells in close relationship with centrally nucleated myofibers (Appendix B, Figure B7) as well as, in areas where damage was less severe, with adjacent blood vessels (Figure 3.6A).  In summary, FAPs are quiescent in undamaged adult muscle, but are highly responsive to signals triggered by myofiber damage. Following damage FAPs proliferate efficiently and are found in close proximity with ailing myofibers. The transient expansion of FAPs, peaking at a time in which commitment of MPs cells to differentiation takes place in vivo, led us to hypothesize that they may play a role in modulating muscle regeneration.  3.2.6. Fibro/adipogenic progenitors provide an environment favoring myogenic differentiation Despite the rapid recruitment of fibro/adipogenic progenitors to proliferate in response to acute muscle damage, neither fibrosis nor adipocytic infiltration is a common outcome of the healing process in young adults. It seems likely that environmental signals present in healthy muscle inhibit FAP differentiation. However, the fact that FAPs are activated at all suggests they may play a role in successful regeneration, and that such role is independent of the formation of fibrocytes or adipocytes.   85 To determine whether FAPs may be a source of signals known to influence myogenic differentiation, we performed qRT-PCR analysis for IGF-1 (Bodine et al., 2001), IL-6 (Serrano et al., 2008), Wnt1, Wnt3A and Wnt5A (Brack et al., 2007) from FAPs and MPs isolated at 0 (undamaged), 2 and 5 days after NTX damage (Figure 3.6B). We found that, compared to MPs, FAPs expressed similar or greater levels of all transcripts at all timepoints. In particular, IGF-1 expression was consistently higher in FAPs than MPs (Figure 3.6B). Also notable was that after NTX damage, IL- 6 expression remained constant in MPs but increased nearly 10 fold in FAPs, suggesting that FAPs represent an inducible source of IL-6 upon activation. Together, these data confirm that FAPs produce several factors known to favor myogenic differentiation.  To test whether signals originating from FAPs regulate the progression of myogenic progenitors toward terminal differentiation, we exposed myogenic cells to FAPs in vitro and measured the effect of co-culture on the spontaneous generation of myofibers. MPs were purified by FACS from GFP+ mice and plated together with an equal number of freshly sorted GFP- FAPs in growth medium. As controls, parallel co-cultures of GFP+ MPs and GFP- MPs were established. After ten days, cultures were stained for MyHC to identify terminally differentiated muscle (Figure 3.7A). The frequency of differentiated cells was calculated as the percentage of GFP+ myonuclei (defined as nuclei present in terminally differentiated MyHC+ GFP+ structures) over total GFP+ cell nuclei. As shown in Figure 3.4A, FAPs do not fuse with differentiating primary myofibers, and are therefore not expected to contribute  86 directly to the number of myonuclei. GFP- MPs will fuse with GFP+ myotubes as syncitial myofibers are generated, thus potentially biasing our analysis toward an overestimation of the number of GFP+ myonuclei in control co-cultures. Despite this confound the frequency of GFP+ myonuclei was drastically increased in the presence of FAPs, reaching an average of 60% compared to 30% in control co- cultures (Figure 3.7A). Thus, FAPs increase the terminal differentiation of myogenic progenitors.  One striking observation stemming from our in vivo studies is that the ratio of FAPs to MPs changes considerably, with FAP numbers peaking 2-3 days following damage, and rapidly falling thereafter. We reasoned that such a change in progenitor numbers might be important if the effects of FAP-derived signals on myogenic cells were dose dependent. To test this hypothesis, we repeated the co- cultivation experiments as described above except that we used 1:1, 1:2, and 1:9 ratios of GFP- FAP to GFP+ MP. To obtain a quantitative molecular readout of the effects of FAPs on myogenic cells, we used flow cytometry to separate GFP+ MP cells from co-cultured FAP or control cells after ten days in culture and measured the expression of myogenic differentiation markers by qRT-PCR. Increasing numbers of FAPs correlated with higher expression of late markers myogenin, Mrf4 and MyHC, revealing a clear dose dependent effect of FAPs on MP differentiation (Figure 3.7B).  Potentially, an increase in progenitor differentiation could be secondary to an increase in culture density and thus to an effect on proliferation. However, no  87 significant difference in proliferation was observed between test and control samples during the first 5 days of coculture (Figure 3.7C). Furthermore despite a small but significant difference in overall co-culture proliferation detected by MTT assay at day 7, the frequency of MPs engaged in cell cycle was comparable in the presence or absence of FAPs (Figure 3.7 C,D). Overall, these data suggest that the increase in myogenic differentiation caused by FAPs is not secondary to an effect on MP proliferation.  The time points at which FAPs cells are most abundant after damage in vivo suggest that they may influence the commitment of proliferating myoblasts to terminal differentiation. This is supported by co-culture data, which suggests that FAPs do not affect proliferation prior to the emergence of highly proliferative myoblasts from days 5-7, yet will significantly increase the number of terminally differentiated MPs at day 10. To test this hypothesis, we analyzed FAP-dependent changes in the expression of myogenic markers by MPs after 7 days of co-culture, a time point that precedes the appearance of myofibers in these cultures. We observed a significant reduction in the expression of early markers Pax3 and Pax7, and a significant increase in myoblast markers MyoD and myogenin in the presence of FAPs (Figure 3.7E). These results suggest that signals originating from fibro/adipogenic progenitors increase the frequency of proliferating primary myoblasts that commit to terminal differentiation. Thus, FAPs generate a transient pro-differentiation niche for MPs. Overall, our findings strongly support a key role of these cells in muscle  88 healing in vivo, suggesting that successful regeneration requires the concerted action of multiple cell types, forming a complex biological system.   89 3.3. DISCUSSION Tissue damage triggers a cascade of events that can lead to regeneration or to repair (Goss, 1992). Regeneration is the outcome of a process that returns the tissue to its normal structure and full functionality, usually by activating local stem/progenitor cells to replace its cellular components. In contrast, repair ensures the continuity of a tissue, but fails to restore its function. In most cases repair is achieved by generating fibrous scar tissue, which in skeletal muscle is often accompanied by adipocytic infiltration. The mechanisms underlying the prevalence of one outcome over the other are still poorly understood. In skeletal muscle, the progenitors responsible for myogenic regeneration are well characterized, as are the mechanisms underlying their activation and differentiation in response to damage. However, very little is known about progenitors involved in repair.  Here, we have used flow cytometry and surface marker expression to identify and characterize non-endothelial, non-hematopoietic progenitors found in normal healthy muscle. We described two new types of progenitors beyond the well-characterized myogenic cells. These mesenchymal progenitors appear to have distinct, non- overlapping developmental potentials. One type is capable of efficient bone and cartilage formation, while the other generates abundant mature adipocytes and fibroblastic cells. Thus in skeletal muscle the chondro-osteogenic, adipogenic and myogenic potential are readily attributable to distinct lineage-committed cell types, suggesting that multipotent mesenchymal stem cells, if present, mainly play a reserve role in this tissue.  90  Of the novel progenitors we describe here, those with chondro/osteogenic potential are rare and do not respond to muscle damage by proliferating, casting doubts on whether they play a significant role in the healing process. Indeed, cartilage and bone are only found within muscle in the context of rare diseases such as fibrodysplasia ossificans progressiva (Shore et al., 2006), and are not a common outcome of healing independently of aging and inflammation.  Fibro/adipogenic infiltration, on the contrary, is a common outcome of chronic muscle disease (Wallace and McNally, 2008). Here we show that fibro/adipogenic progenitors represent a significant fraction of the mononuclear mesenchymal cells found in adult skeletal muscle. These cells are quiescent in adult muscle, and they rapidly enter the cell cycle in response to tissue damage. Their activation does not appear to be the result of direct damage, as the snake toxin NTX used in our studies specifically damages mature myofibers but not mononuclear cells (Harris, 2003; Harris et al., 2003). Recent literature suggests that both adipogenic and fibrogenic cells originate from myogenic cells through alternative lineage choice dictated by a pathological environment (Brack et al., 2007; Li et al., 2004; Li and Huard, 2002; Shefer et al., 2004). Shefer et al. used single myofiber cultures to demonstrate that myogenic and adipogenic cells are associated with the same fiber but stopped short of formally proving that fat and muscle can arise from the same cell (Shefer et al., 2004). Rando’s laboratory has recently reported that, in aging mice, fibrogenic cells can arise from Pax7+ myogenic progenitors in response to altered WNT signaling  91 (Brack et al., 2007). We cannot formally exclude that myogenic cells may become an additional source of fibroblasts when exposed to an aging environment. However the abundance and proliferative capacity of FAPs strongly suggests that they are the main source of fibroblasts and adipocytes in adult tissue. Thus, these cells are excellent candidates for progenitors involved in repair. As such, their study may lead to therapeutic strategies to reduce fibrosis in chronic disease.  Adipogenic progenitors (AP) with a surface phenotype identical to that of FAPs were recently isolated from adipose depots (Rodeheffer et al., 2008; Tang et al., 2008), but it is currently unclear whether FAPs represent the same cells. Unlike APs, single FAPs efficiently generate both fibroblasts and adipocytes. Furthermore, PPARγ2 expression was very low in freshly isolated FAPs from undamaged muscle (Appendix B, Figure B4). This appears to be consistent with data from Tang et al., who used a transgenic PPARγ reporter to identify APs from adipose vasculature but was unable to find similar PPARγ+ cells in muscle. Thus, FAPs may represent an earlier bi-potent progenitor that precedes the initiation of PPARγ expression.  Despite their strong proliferative response to muscle damage and their ability to generate adipocytes in vitro and after subcutaneous transplantation in vivo, activated FAPs disappear from the tissue and do not generate terminally differentiated cells in young muscle. This suggests that local environmental influences control their ability to survive and differentiate. On the other hand, the fact that FAPs are activated at all suggests that they may play a physiological role in regeneration. To test whether  92 signaling molecules produced in FAPs have an effect on MP behavior, we co- cultivated freshly sorted cells of each type. It is important to note that freshly sorted myogenic cells have different properties compared to the commonly used primary myoblasts. Myoblasts are traditionally selected based on adherence and preferential proliferation in specific media (Rando and Blau, 1994). This process yields cultures characterized by a low rate of spontaneous differentiation, likely because of the strong selection for rapidly growing cells. Freshly sorted progenitors on the contrary display a significant rate of spontaneous myofiber formation even in expansion conditions, with up to 30% of the nuclei contained in MyHC+ structures after ten days of culture. This behavior is closer to that observed in vivo, where the proliferating myoblast is a transient state of finite duration. We found that co-culture with FAPs significantly increased this spontaneous rate of differentiation independently of any effects on MP proliferation, and that the magnitude of this effect was dependent on the ratio of FAP to MPs in the co-culture, suggesting that the drastic changes in the relative abundance of these cells observed during regeneration in vivo may play a physiological role. In summary, FAPs transiently expand to provide an environment favoring the commitment of proliferating myoblasts to terminal differentiation.  Specialized environments required for differentiation have been described in tissues characterized by high cellular turnover. Within the thymus for example, progression along the T-cell developmental cascade is spatially regulated, requiring the migration of progenitors to specific locations (Ladi et al., 2006). Here, we report the first  93 example of a transient pro-differentiation niche. It is tempting to speculate that in tissues with very low homeostatic cell replacement such as muscle, pro- differentiation signals may only be required for a short time following acute damage, and thus be provided by short lived, specialized cells. Ascertaining the prevalence of such temporally-regulated niches in low turnover tissues will require further investigation.  Our work describes a novel population of fibro/adipogenic progenitors resident in muscle tissue, and assigns to these cells a role in successful muscle regeneration. These findings introduce the concept of muscle healing as system where the main players involved in regeneration and repair communicate with each other to coordinate their output. If the additional role played by inflammatory cells in directing these processes is considered, a picture of healing as a complex biological system emerges.  We can only hope to understand such complex systems by studying them in their entirety.   94 3.4. EXPERIMENTAL PROCEDURES 3.4.1. Animals All mice were maintained in a pathogen-free facility, and all experiments were performed in accordance with University of British Columbia Animal Care Committee regulations. Adult C57BL/6 mice (>8 weeks) were used in this study. C57BL/6-CMV- βActin-GFP transgenic mice were a generous gift from I. Weissman, Stanford. In vivo lineage tracing was performed in Myf5-Cre-R26R3-YFP mice, generated by breeding heterozygous Myf5-Cre (P. Soriano) with Rosa26-YFP (Srinivas et al., 2001) reporter mice. Muscle damage was induced by intramuscular injection of 0.15µg Notexin snake venom (Latoxan), into the tibialis anterior (TA) muscle.  3.4.2. Tissue preparation Skeletal muscle from both hindlimbs was carefully dissected and then gently torn with tissue forceps until homogeneous. Two milliliters collagenase type 2 (Sigma, 2.5u/ml), in 10mM CaCl2, was added to every 2 hindlimbs, and the preparation was placed at 37 °C for 30 minutes. After washing, a second enzymatic digestion was performed with CollagenaseD (Roche Biochemicals, 1.5u/ml) and Dispase II (Roche Biochemicals, 2.4u/ml), in a total volume of 1ml per mouse, at 37 °C for 60 minutes. Preparations were passed through a 40µm cell strainer (Becton Dickenson [BD]), and washed. Resulting single cells were collected by centrifugation at 400 rcf for 5 min.   95 3.4.3. Flow cytometry / FACS Cell preparations were incubated with primary antibodies for 30min at 4C in supplemented PBS containing 2mM EDTA and 2% FBS, at ~3x107 cells per ml. We used the following monoclonal primary antibodies: anti-CD31 (clones MEC13.3 [BD] and 390 [Cedarlane Laboratories]), anti-CD34 (clone RAM34 [eBiosciences]), anti- CD45 (clone 30-F11 [BD]), anti-CD45.1 (clone A20 [BD]), anti-CD45.2 (clone 104 [eBiosciences]), anti-Sca1 (clone D7 [eBiosciences]) and anti-α7 integrin (produced in-house). For all antibodies we performed fluorescence minus one controls by staining with appropriate isotype antibodies. Where necessary, biotinylated primary antibodies were detected using Streptavidin coupled to phycoerythrin, allophycocyanin, or FITC (Caltag). To assess viability, cells were stained with propidium iodide (1µg/ml) and Hoechst 33342 (2.5µg/ml) and resuspended at ~1x107 cells/ml immediately prior to sorting or analysis. Analysis was performed on LSRII (BD) equipped with 3 lasers. Data was collected using FacsDIVA software. Sorts were performed on a FACS Vantage SE (BD) or FACS Aria (BD), both equipped with 3 lasers, using a 100µm nozzle at 18psi to minimize the effects of pressure on the cells. Sorting gates were strictly defined based on isotype control (fluorescence minus one) stains. Biexponential analysis was performed using FlowJo 8.7 (Treestar) software.  3.4.4. Cell culture Cells were grown in high-glucose Dulbecco’s modified eagle medium (DMEM), supplemented with 2.5ng/ml bFGF (Invitrogen) 20% FBS and 10% heat-inactivated  96 horse serum. This media is referred to in text as ‘growth media’. Cells were seeded in tissue-culture treated plates coated with Matrigel (BD) or Type 1 collagen (Sigma). After sorting, cells were allowed to adhere for 3 days, after which ½ the medium was changed. Media was changed every 2-4 days thereafter. For mesenchymal differentiation, we used reported conditions. Briefly, we cultured cells in DMEM with 20% FBS under the following conditions: for osteogenic differentiation, we supplemented media with 10nM dexamethasone, 5mM β-glycerophosphate and 50ug/ml ascorbic acid; for adipogenic differentiation, we supplemented media with 0.25µM dexamethasone, 0.5mM Isobutylmethylxanthine, 1µg/ml insulin, 5µM troglitazone; for chondrogenic differentiation we pelleted cells by centrifugation at 400rcf for 5min, and grew them in media supplemented with 1ng/ml TGFβ1 and 50µg/ml ascorbic acid.  3.4.5. Limiting dilution analysis One to one hundred cells from each test population was sorted into individual wells of a Matrigel-coated 96 well plate directly from the sorter. Cells were grown as described in the text. After three weeks, cultures were fixed and stained for MyHC and nuclei. Wells were scored for the presence of colonies (>8 cells), cells undergoing terminal myogenic differentiation (MyHC-positive), and lipid laden adipocytes. A minimum of 30 replicate wells was generated for each cell dose. LDA calculations were based on the single hit Poisson model (see Statistics).   97 3.4.6. Gene expression analysis RNA isolation was performed using RNeasy mini kits (Qiagen), and RNA quantification was performed using a ND1000 spectrophotometer (Nanodrop). Reverse transcription was performed using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems). Gene expression analysis was performed using Taqman Gene Expression Assays (Applied Biosystems), on a 7900HT Real Time PCR system (Applied Biosystems). Data was acquired and analyzed using SDS 2.0 and SDS RQ Manager software (Applied Biosystems).  3.4.7. Transplantation Host mice (wild type) were anesthetized using isoflurane. Donor cells were isolated from transgenic mice ubiquitously expressing GFP+. Cells were sorted into cold DMEM and collected by centrifugation at 450rcf for 5minutes. For subcutaneous transplantation, cells were resuspended in 25ul Matrigel and loaded into an ice-cold needle and syringe immediately prior to injection. Cells were injected into the subscapular region, with control cells injected on the contralateral side. For intramuscular transplantation, sorted cells were resuspended in 20ul sterile PBS and injected into the TA. Tissues were harvested after 3 weeks.  3.4.8. Histology and imaging Prior to tissue harvest, animals were perfused transcardially with 20ml PBS/ 2mM EDTA, followed by 20ml 4% paraformaldehyde (PFA). Tissues were processed for  98 cryosectioning or paraffin embedding using standard methods. Immunostaining was performed using monoclonal antibodies against perilipin (Sigma), α smooth muscle actin (Sigma), ER-TR7 (Novus), FSP1 (Novus) myosin heavy chain (in-house) or BrdU (BD). Briefly, samples were permeabilized in 0.3% Triton-X-100 (Sigma) in PBS, and blocked for 2hrs at room temperature in PBS containing 25% normal goat serum (NGS), 0.3% Triton-X-100, 3% bovine serum albumin (BSA) and 0.1%NaN3. Cells were stained overnight at 4C using antibody diluted in 10% NGS, 0.3% Triton- X-100, 3% BSA and 0.1% NaN3. Primary antibody was detected using goat anti- mouse IgG antibodies conjugated to Alexa 568 or 488 (Molecular Probes), or using the anti-Mouse IgG HRP detection kit (BD). Cells were visualized using a microscope (Axiovert 200 for inverted microscopy, Axioplan2 for conventional microscopy; Carl Zeiss Microimaging) and images were acquired using a charge-coupled device camera (Retiga Exi [Axiovert 200] or Retiga Ex [Axioplan2], qImaging) and OpenLab4 software (Improvision). To confirm the specificity of the GFP signal, we compared the signal to non-specific autofluorescent background in all other channels (Brazelton and Blau, 2005). Confocal microscopy was performed using a Nikon C1 laser scanning confocal microscope equipped with lasers at 488nm, 568nm, and 633nm. Images were captured using the least exposure time possible, and manipulation of brightness and contrast, coloring adjustments and assembly into figures were performed using ImageJ, OpenLab4 (Improvision), Illustrator CS3 (Adobe) and Photoshop CS3 (Adobe).   99 3.4.9. BrdU labeling studies For in vivo studies, BrdU was administered in drinking water (0.8g/L in 2% sucrose) and by intraperitoneal injection (100mg/kg). In cell culture experiments, 10µM BrdU was added to culture medium. BrdU treatment regimens are described in the text. For flow cytometric analysis, cells were stained for surface markers as indicated. Samples were fixed in 2% PFA for 20 min, and membranes were permeablized in 0.2% saponin for 10 min. Cells were treated with 30µg DNAse I (Sigma) for 1h at 37°C, after which a 1:25 dilution of anti-BrdU (clone PRB-1 [Invitrogen]) was applied to the samples for 30min at room temperature.  3.4.10. MTT assay Assays were performed in 96-well cell cultures. 50 µl of MTT solution (5 mg/ml dissolved in PBS) was added to each well and samples were incubated at 37°C for 4h. The solution was removed, and the purple formazan salt product resulting from the reduction of the yellow MTT was solublized in 100 µl DMSO, and quantified spectrophotometrically at 570 nm (SpectraMax 190, Molecular Devices).  3.4.11. Statistics Preliminary analysis and data collation was performed using Microsoft Excel. Statistical tests, including t-tests, ANOVA and regression analyses were performed using Prism 4 (GraphPad Software). Analysis of limiting dilution data was performed using a web application made available by the Walter and Eliza Hall Institute of  100 Medical Research, Melbourne, Australia (http://bioinf.wehi.edu.au/software/limdil/) (Shackleton et al., 2006). This software tests departures from the single-hit Poisson model using a generalized linear model.   101 3.5. TABLES Table 3.1. FAP cells do not contribute to skeletal muscle in vivo  2x104 FAPs or MPs were isolated from hPLAP transgenic mice and injected into the undamaged or damaged tibialis anterior (TA) of syngeneic wildtype animals. Damage was induced by notexin injection 48 hours prior to cell transplantation. Tissues were harvested after 3 weeks, and myogenic engraftment was enumerated by histochemistry.  Donor Cells Recipient TA Animal ID Number of hPLAP+ Myofibers detected 2x104 hPLAP+ MP Undamaged 1 77   2 62   3 55   4 35 2x104 hPLAP+ MP NTX-damaged 1 202   2 129   3 192 2x104 hPLAP+ FAP Undamaged 1 0   2 0   3 0 2x104 hPLAP+ FAP NTX-damaged 1 0   2 0   3 0     102 3.6. FIGURES Figure 3.1. Prospective isolation of progenitor populations from skeletal muscle  (A) Viable cells were identified based on forward/side scatter, Hoechst staining to exclude anuclear debris and low PI staining to exclude dead cells.  Hematopoietic (CD45-) and endothelial cells (CD31-) cells were also excluded from analysis. (B) Expression of CD34 and Sca-1 in hoechstmid PIlo CD45- CD31- (lin-) cells. Sca-1- CD34-, Sca-1- CD34+ (MP) and Sca-1+ CD34+ cells were sorted and characterized. (C) Lin- Sca-1- CD34- cells contain osteogenic and chondrogenic activity. Mineralized, multilayered nodules in cultures grown in osteogenic conditions for 10 weeks, and stained with alizarin red (left, scale bar 100µm). Alcian blue positive cartilage in cryosections of cell pellets grown in chondrogenic conditions (right; scale bar 25µm). (D) Lin- Sca-1+ CD34+ cells contain adipogenic progenitors. Sorted cells spontaneously gave rise to multilocular adipocytes (center). Triglycerides were detected by Oil red O staining in unilocular mature adipocytes after 30 days (right). Scale bars at 50µm (left, center) and 100µm (right). (E) MP cells spontaneously differentiate in culture. Myosin heavy chain (MyHC)- expressing myotubes were observed after 15 days in culture (center, right). Scale bars at 50µm (left) and 100µm (center, right).  103 (F) Sca-1- CD34+ (MP, red), but not Sca-1+ CD34+ adipogenic cells (blue) express α7 integrin. The specificity of the α7 staining was confirmed by the “fluorescence minus one” (FMO) control.  104    105 Figure 3.2. Lin- Sca-1+ CD34+ cells generate both fibroblasts and adipocytes  Lin- α7- Sca-1+ cultures were grown for three weeks in growth media and then immunostained using antibodies against fibroblast markers. (A) FSP-1 (B) ER-TR7 (C) Single lin- Sca-1+ α7- cells were deposited into individual wells of a 96-well plate directly from the sorter. After three weeks culture in growth medium, cells were immunostained for smooth muscle actin (SMA) and perilipin. Scale bar: 50µm.      106 Figure 3.3. Developmental potential of sorted progenitor populations in vivo  (A) Lin- Sca-1- CD34+ (MP) and Lin- Sca-1+ CD34+ (FAP) cells were isolated from transgenic mice ubiquitously expressing GFP. (B) MP cells engraft in skeletal muscle. 5x104 freshly isolated MP cells from GFP- expressing mice were injected into the tibialis anterior muscle of syngeneic hosts. Three weeks later, we observed GFP-expressing myofibers along the needle tract. (C) Subcutaneous transplantation of FAP cells. 4x104 freshly isolated FAP cells from GFP-expressing mice were injected subcutaneously into syngeneic GFP- recipients. Three weeks later, confocal microscopy revealed GFP+, perilipin-expressing adipocytes located between the skeletal muscle (m) and dermis (d). GFP expression was detected by immunostaining. (D) High magnification image of transplanted GFP+ FAPs shows co-localization of GFP with the mature adipocyte marker perilipin. A maximum intensity projection image from a confocal optical stack is shown. All scale bars at 50µm.  107     FAP  108 Figure 3.4. Skeletal muscle-derived FAPs and MPs have distinct developmental potentials and do not arise from a common progenitor  (A) FAP and MP co-cultivation confirms that their developmental potentials are non- overlapping. FAPs (A) or MPs (Supplemental Figure 7) sorted from transgenic GFP+ animals were co-cultivated for 14 days with equal numbers of GFP- MPs. Confocal microscopy revealed no contribution of GFP+ FAPs to MyHC+ cells, and no contribution of GFP- MPs to lipid-laden adipocytes (A). (Scale bars at 100µm). (B) The Myf5 locus is expressed in MP, but not FAP cells. MPs (left) and FAPs (right) were sorted from Myf5-LacZ transgenic mice and allowed to adhere overnight. Myf5-LacZ expression was visualized by X-gal staining, and nuclei were visualized using Hoechst dye. Scale bars at 50µm. (C) Quantification of Myf5-expressing MPs and FAPs from (B). No Myf5-expressing FAP cells were detected. (D) Schematic of the lineage tracing strategy. Myf5-Cre-R26R3-YFP mice were generated by crossing Myf5-Cre mice with a reporter strain carrying YFP integrated into the Rosa26 locus downstream of a floxed transcriptional stop sequence. Cre expression results in the heritable and irreversible expression of YFP under control of the Rosa26 locus. (E) MPs, but not FAPs arise from a Myf-5 expressing precursor cell. Analysis of MPs and FAPs from Myf5-Cre-R26R3-YFP mice revealed YFP expression in a large proportion of MPs (top) whereas over 99% of FAPs were YFP negative (bottom).  109 (F, G) YFP expression in sorted MPs and FAPs from Myf5-Cre-R26R3-YFP mice after 5 days in culture. Over 85% of cultured MPs expressed YFP (F); in contrast, no YFP was detected in FAP cultures (G).  110      111 Figure 3.5. FAPs proliferate in response to muscle damage  (A) Detection of BrdU incorporation in endogenous FAPs. Myofiber damage was induced by intramuscular injection of notexin into the TA muscle. Starting 12hrs before damage, BrdU was administered by intraperitoneal (IP) injection (100mg/kg every 12hrs) and in drinking water (0.8mg/ml in 2% sucrose). After 2 days, BrdU incorporation in FAPs was detected by flow cytometry. Graphs show overlaying of BrdU+ events (blue dots) onto Sca-1/CD34 plots (right). The specificity of BrdU staining was confirmed by staining cells from non-BrdU treated animals (left). (B) Comparison of FAP and MP proliferation kinetics after damage. BrdU was administered as in (A). TAs were harvested every 24hrs after damage induction, up to 96hrs. The data is presented as BrdU+ events (blue) overlaid onto Sca-1/α7 integrin plots. Percentages shown indicate the frequency of BrdU+ cells within the gate. (C) Daily analysis of damage-induced progenitor cell proliferation. Damage was induced at time d=0 in all animals. BrdU was administered by IP injection (100mg/kg) at 24 and 12hrs prior to harvest.  Time after damage indicates the period over which BrdU was administered. Data was analyzed by flow cytometry as in (B). (D) Dynamic changes in the ratio of FAP to MP cells after damage. Ratios were determined by comparing the number of events falling into FAP or MP gates using flow cytometric analysis. Day 0 represents undamaged animals. (E) Regression analysis of limiting dilution data from FAPs and MPs sorted from undamaged and damaged TAs. The indicated number of cells were deposited in replicate wells of 96 well plates directly from the FACS sorter. After 21 days, wells  112 were scored for the presence of colonies. Data from damaged TAs is highlighted in red. Data was analyzed based on the single-hit Poisson model for limiting dilution analysis. 95% confidence intervals are represented by dashed lines and are located in brackets in the legend. *p<0.05, **p<0.01  113    114 Figure 3.6. FAPs are in close proximity to myofibers and express pro- differentiation signals after damage  (A) CD31- Sca-1+ cells are found adjacent to both regenerating myofibers and blood vessels after NTX damage. Tissues were harvested 3 days after damage induction, fixed, cryosectioned, then immunostained for CD31 and Sca-1. A single optical slice from a confocal Z-stack is shown. Background autofluorescence from myofibers is shown in the blue channel. Scale bar at 25µm. (B) FAPs express factors known to promote myogenic differentiation. qRT-PCR analyses was performed on cDNA from FAPs and MPs isolated from skeletal muscle at 0 (undamaged), 2 and 5 days after damage, using Taqman probe and primer sets. ND = no signal detected.  115     FAP  116 Figure 3.7. FAPs enhance myogenic differentiation  (A) Immunohistological analysis of co-cultures revealed increased MP differentiation in the presence of FAPs. 5000 MPs were isolated from GFP+ mice, and co-cultivated with 5000 GFP- FAPs or GFP- MPs. After 10 days cultures were fixed and stained for MyHC. The data is expressed as the ratio between nuclei in GFP+ MyHC+ cells (myonuclei) and total nuclei in GFP+ cells. (B) FAPs induced MP differentiation in a dose dependent fashion. GFP+ MPs were co-cultivated with increasing numbers of GFP- FAPs and re-isolated by FACS after ten days. The expression of markers of myogenic differentiation was analysed by qRT-PCR using Taqman probe and primer sets spanning exon-exon boundaries. (TBP: TATA binding protein.) (C) MTT proliferation assay revealed little difference in proliferation between MP- only or MP-AP co-cultures. Assays were performed on freshly sorted cell cultures containing either 5000MP, or 2500MP with 2500 FAP. Data is expressed as absorbance at 570nm (A570). (D) No difference in BrdU incorporation in MPs exposed to equal numbers of GFP+ MPs or GFP+ FAPs. 5000 GFP- MPs were co-cultivated with either 5000 GFP+ MPs or 5000 GFP+ FAPs for a total of 7 days. BrdU (10µM) was applied during the last 24hrs of co-culture. GFP+ cells were removed using FACS, and immunostaining for BrdU was performed on the remaining GFP- MPs. (E) FAPs increase MP commitment to terminal differentiation. 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THESIS SUMMARY It has become clear after over thirty years of work that new adipocyte production, or de novo adipogenesis, indeed occurs in adults; however, the relevance of this phenomenon to adipose physiology and tissue pathogenesis is not well understood (Greenwood and Hirsch, 1974; Johnson et al., 1978; Miller et al., 1984; Spalding et al., 2008). Based on data gathered over the last decade, it has been postulated that smaller adipocytes are more functional than large adipocytes; therefore, the ability to generate new, small, healthy adipocytes may reduce the risk of developing metabolic disease, regardless of an organism’s overall fat mass (Bluher et al., 2002; Gray and Vidal-Puig, 2007; Khan et al., 2009; Kim et al., 2007; Rajala and Scherer, 2003; Vazquez-Vela et al., 2008). However, new adipocyte production in non- adipose tissues, such as skeletal muscle and bone marrow, is associated with aging, chronic disease and tissue decompensation (Rosen and Bouxsein, 2006; Wallace and McNally, 2008). Thus, the ability to generate new adipocytes may be perceived as either physiological or pathological depending on context and tissue. Identifying the cells from which new adipocytes arise in a depot- and tissue- specific manner, and determining the factors that govern their activity and fate, represent major advances towards understanding how, when and where de novo adipogenesis occurs. This knowledge will hopefully allow for control of these mechanisms for therapeutic benefit.   125 Until very recently, cells fated to become adipocytes in vivo had not been identified; therefore, there has only been a very limited understanding of the mechanisms underlying new adipocyte production. The work presented in this thesis is the first to demonstrate the prospective identification and isolation of adipogenic progenitors (AP) from multiple sources. This was achieved using optimized mechanical and enzymatic digestion methods, which allowed us to analyze mononuclear cells from freshly dissociated solid tissues. When combined with multi-parameter flow cytometry and fluorescence-activated cell sorting (FACS), we were enabled to identify and sort distinct cell populations derived from subcutaneous and visceral fat, and skeletal muscle. We isolated a number of populations from each tissue, most notably Hoechstmid PIlo CD31- CD45- α7 integrin- (lin-) Sca-1+ CD34+ (AP) cells, and show using a variety of assays that APs are fated to become adipocytes in vitro and after transplantation in vivo, and represent virtually all of the adipogenic activity arising from their tissues of origin.  My colleagues and I show in Chapter 2 that AP abundance and activity are different between subcutaneous and visceral fat depots, and propose that these differences underlie whether a fat depot expands by hypertrophy or hyperplasia. Although two other groups have very recently identified APs from fat tissues with identical markers, neither had shown that depot-specific differences exist (Rodeheffer et al., 2008; Tang et al., 2008). Furthermore, whether new adipocyte production occurs at a constant rate in adults, as recently proposed, or can be modulated by dietary factors, remains a hotly debated topic in the field (Spalding et al., 2008). Our data  126 provides the first evidence that dietary influences can influence the proliferation of adipogenic progenitors in adults, and that this effect is depot-specific. As a result, we propose that the mechanism of adipose tissue expansion is highly depot-dependent, and is influenced by differences in AP content and activity (Figure 4.1). Thus, according to our model, the presence of active APs determines a depot’s ability to maintain a pool of small, functional adipocytes that may slow the progression of metabolic disease.  On the other hand, new adipocyte production in skeletal muscle occurs as a result of muscle decompensation, and is thought to arise from dysfunctional myogenic progenitors (MP) through alternative lineage choice owing to changes to the normal tissue microenvironment (Shefer et al., 2004; Wallace and McNally, 2008). In Chapter 3, my colleagues and I show that single skeletal muscle resident AP cells are capable of generating both adipocytes and fibroblasts, consistent with the fact that fibrosis and adipogenesis are invariably linked in fatty degeneration of decompensated skeletal muscle. Thus we called these cells fibro/adipogenic progenitors (FAP). Furthermore, contrary to what has been proposed, we show that FAPs do not arise from MPs and are incapable of muscle differentiation in vitro or after transplantation into damaged muscle in vivo. Thus our data provides a fresh new perspective to the field, that FAPs and MPs represent distinct muscle-resident cell populations, both of which are quiescent at steady-state, but are capable of giving rise to different mature lineages when the appropriate signals arise (Figure 4.2).  127  To explore the link between muscle regeneration and fibro/adipogenic infiltration, we examined FAP activation after muscle damage in young, healthy animals, and to our surprise, found that a large proportion of FAPs are activated 24-48hrs after damage, and prior to MPs. Since fibrosis and adipogenesis are not common outcomes of muscle damage in healthy animals, we proposed that FAPs might play a physiological role during muscle regeneration. Through subsequent experiments, we found that FAPs produce several factors capable of inducing myogenic differentiation, and co-cultivation of FAPs with MPs significantly increased the proportion of MPs committing to terminal differentiation. These experiments have led us to hypothesize that FAPs can instruct satellite cells to terminally differentiate into proliferative myoblasts during normal skeletal muscle regeneration, and that this role is completely independent of FAP fibro/adipogenic differentiation (Figure 4.3).   128 4.2. PHYSIOLOGICAL RELEVANCE The models presented in this thesis attempt to explain the contribution of adipogenic progenitors to the physiological state or to the natural progression of disease. Throughout our study design, we made several decisions to ensure that our experiments remained as physiologically relevant as possible. As these studies were all performed in mouse, the applicability of these results to human tissues has not been established. To directly address these questions would require a careful comparison of similar cell populations between mouse and human tissue. Thus, one of the lab’s future goals is to isolate APs from human tissues. Nonetheless, our HFD regimen allowed us to induce adipocyte hypertrophy and hyperplasia in mice, which are relatively more resistant to fat mass expansion than humans due to their relatively high levels of HDL and low levels of LDL and VLDL (Hughes et al., 1997). Thus, it is plausible that our findings may be more easily recapitulated in humans, which have a greater propensity for fat mass expansion due to the relative abundance of LDL and VLDL.  Using flow cytometry / FACS allowed us to work exclusively with pure, primary cell populations, including single cells, that had been freshly isolated from tissues. Furthermore, the results of in vitro differentiation assays were verified by both clonal analysis and by transplantation of freshly sorted cells in vivo, ensuring the highest standard of scientific rigor. Finally, we took advantage of our optimized strategy for flow cytometric analysis of BrdU incorporation by assessing the in vivo proliferation of APs isolated from animals exposed to physiological stressors, such as high fat  129 diet or muscle damage. This was a powerful technique that enabled us to generate easily interpretable, physiologically relevant results. Similar flow cytometric assays using readouts for apoptosis, cell signaling cascades, and lineage tracing are being optimized in the lab, and will provide even more tools with which to interrogate these cells in the future.  Nonetheless, the gold standard for determining physiological relevance remains loss of function / gain of function experiments. The generation of mice that are specifically devoid of APs, or those that have overactive APs, requires the development of genetic tools and specifically, the identification of AP-specific genetic markers. To this end, our lab is currently generating microarray data from APs isolated from subcutaneous and visceral fat, and skeletal muscle; however the development of the appropriate genetic tools remains several years away. In the context of muscle injury, lipodystrophic mice such as the A-ZIP strain, which contains a dominant negative bHLH gene driven by the AP2 promoter, could provide an alternative strategy to test the physiological effects of AP ablation on skeletal muscle regeneration. However, preliminary experiments in our lab revealed that APs are indeed present in A-ZIP animals, likely because APs do not express the AP2 gene, thereby negating this approach.   130 4.3. AGING AS A MODEL TO EXPLORE AP DYSFUNCTION In aging there is a marked redistribution of body fat resulting in smaller fat depots, and increased fat in non-adipose tissues such as skeletal muscle and bone marrow (Brack et al., 2007; Kirkland et al., 2002; Rosen and Bouxsein, 2006). Although the reasons for this are not well understood, the work presented in this thesis suggests that changes in AP activity likely play a role. Experiments in primary pre-adipocytes, which are a heterogenous population cultured from fat depot stromovascular cells, revealed that cells isolated from aged animals were less capable of adipogenic differentiation (Djian et al., 1983; Kirkland et al., 2002). However, it is unclear whether this is due to a loss in the relative number of adipose forming cells in those cultures, or whether cell autonomous changes occurred with age. Our ability to prospectively identify APs and assay pure cell populations allows us to address this question.  One hypothesis to explain the effects of aging on whole body fat distribution is that fat depot expandability declines with age due an inability of APs to replace dead adipocytes or to generate new ones as required. Thus, the prolonged effects of the resulting metabolic dysregulation, combined with the life-long stresses in non- adipose tissues (injury, inflammation, etc), leads to activation and differentiation of non-adipose tissue-resident APs such as skeletal muscle FAPs. In a preliminary experiment using tissues pooled from aged mice (>20 months old), we observed a clear reduction in the number of lin- Sca-1+ CD34+ AP cells in the visceral fat when compared to young animals (<4 months old); however, we observed little difference  131 in subcutaneous fat (Appendix C, Figure C1). Although the data is preliminary, it is tempting to speculate that there are depot-specific differences in the rate of AP exhaustion with age that may be important in determining the age-dependent physiological function of a particular depot.  We performed similar experiments in the context of aged skeletal muscle, and found that the ratio of APs to myogenic progenitors (MP) is significantly higher in aged animals (Appendix C, Figure C2). However, this appears to be caused by an age- dependent reduction in the number of MPs and not due to an increase in the number of APs (Appendix C, Figure C2). Clearly, these results will require significant additional work to allow appropriate interpretation. Notably, limiting dilution analyses and careful in vitro quantification of adipogenic activities must be performed on these cells to ensure that AP surface marker phenotypes do not change with age. Nonetheless, our results suggest that AP biology changes with age, and likely plays a role in the age-related redistribution of fat.   132 4.4. CELL-AUTONOMOUS DIFFERENCES BETWEEN APS HARVESTED FROM DIFFERENT DEPOTS AND TISSUES Although a careful comparison between APs isolated from different anatomical sites is beyond the scope of this thesis, it is a logical and necessary extension of our work. We have described striking differences in the frequency and activity of AP cells isolated from visceral (V-AP) and subcutaneous (S-AP) fat, but we cannot say whether these differences could be due to cell-autonomous effects. We first attempted to answer this question by comparing the size and adipogenicity of single- cell derived V-AP and S-AP colonies, and to date we have not been able to note any significant difference. More work is needed to address these issues.  A distinct difference can be observed in the frequency of lin- cells from both fat depots that fall within the AP (Sca-1+ CD34+) gate—this represents 16% of lin- cells in subcutaneous fat, and 56% of lin- cells in visceral fat (Figure 2.2 A, B). Despite that the V-AP gate captures greater numbers of lin- cells than the S-AP gate, only about 1 in 1000 V-APs generate adipocyte colonies, whereas 1 in 40 S-APs can do the same. Although loss of cell viability due to tissue disruption and FACS could have led us to underestimate the true adipogenicity of this population, it is unlikely to explain the discrepancy we observed between the frequency of V-APs and S-APs as both tissues were treated using identical methods. Thus, it appears that other cell types are present in the visceral fat lin- Sca-1+ CD34+ population, suggesting that an additional marker is required to further purify APs. Current candidates under investigation include CD24, Thy-1, and PDGFr (both α and β isoforms). In particular,  133 CD24 was used by Friedman’s lab to select for a rare subset S-APs (~3%) that had greater in vivo adipogenic capacity than CD24- APs (Rodeheffer et al., 2008); however, preliminary experiments in our lab using CD24 were unsuccessful at enriching adipogenic cells from either S-APs or V-APs. Regardless, whether this marker will be found on APs from both depots, or only on APs from one depot, will provide valuable insight about whether cell-autonomous differences exist between S-APs and V-APs. Furthermore, the identification of another marker may allow for isolation of APs from human tissues as our current strategy uses Sca-1, which has no human homologue (Bradfute et al., 2005).  It is not surprising that we found adipogenic progenitors in fat tissue; however, finding them in skeletal muscle was initially quite startling. Nonetheless, our data clearly shows that these muscle-derived APs are associated with muscle fibres, are distinct from myogenic cells, and are capable of generating adipocytes at an equal or greater efficiency than S-APs or V-APs. Furthermore, based on our hypothesis that these cells may be involved in fibro/adipogenic infiltration of diseased muscle, we have explored the fibrogenic potential of these cells and show that they are capable of generating both adipocytes and cells that express typical fibroblast markers; thus we call them FAPs. Whether S-APs or V-APs also generate fibroblasts is unclear, but based on the morphology of their single-cell derived cultures, and the fact that only about half of the cells from single S-AP or V-AP derived colonies become adipocytes, it is highly probable. Further, microarray experiments comparing FAPs to S-APs and V-APs are under progress, and we hope  134 the results will clarify whether cell-autonomous differences exist between these populations.   135 4.5. APS FROM FAT AND SKELETAL MUSCLE ARE CLOSELY ASSOCIATED WITH BLOOD VESSELS Histological examination of adipose tissue development performed over several decades revealed a close spatial and temporal association between blood vessel formation and adipocyte formation (Hausman et al., 1980). Experiments performed in Graff’s lab confirmed these early findings, and revealed that adipogenic progenitors are associated with blood vessels in fat tissue (Tang et al., 2008). This is in accordance with our data, and data from others, which also show that muscle- derived adipogenic cells are vessel-associated (Figure 3.6) (Shefer et al., 2004). As these cells occupy a perivascular position, it is conceivable that APs are pericytes. However, populations of pericytes have been shown to be capable of multilineage differentiation (Dellavalle et al., 2007), which despite our greatest efforts we have not been able to demonstrate in any of our AP populations. In fact, our data strongly suggests otherwise, demonstrating that S-APs, V-APs and FAPs are only capable of adipogenic and fibrogenic differentiation, and casting doubt over the hypothesis that they may be multipotent stem cells or pericytes. Nonetheless, APs may represent a lineage-committed daughter cell of a multipotent pericyte. This hypothesis remains to be tested.   136 4.6. IDENTIFICATION OF APS IN OTHER TISSUES Since APs are found close to vessels, they may form an integral part of all vessels and thus can be found in all tissues, or they may be tissue-specific cells that simply reside near the vasculature. To begin to address this question, we sought to isolate APs from other tissues in which adipocyte infiltration has been described: bone marrow and tendon (Hashimoto et al., 2003; Jarvinen et al., 1997; Rosen and Bouxsein, 2006). In preliminary FACS experiments on bone marrow and crushed bone isolated from wild-type mice, we were unable to isolate significant numbers of lin- Sca-1+ CD34+ cells. Furthermore, cultivation of these rare cells did not yield cells capable of adipocyte differentiation. This data is not completely surprising given that adipocyte infiltration into bone marrow is a common occurrence in humans but is only observed in mutant mice, including SAMP6 osteoporotic animals and congenic 6T animals that have increased body fat and reduced bone mineral density (Rosen et al., 2004; Rosen and Bouxsein, 2006; Uchiyama et al., 1994). On the other hand, abundant lin- Sca-1+ CD34+ cells were identified from mouse achilles tendon cell preparations. Cultivation of different Sca-1 / CD34 expressing populations from lin- tendon cells revealed that differentiating adipocytes arose exclusively in the Sca-1+ CD34+ population, confirming the presence of APs in lin- Sca-1+ CD34+ tendon- derived cells  (Appendix C, Figure C3). Although the data is preliminary, together they suggest that APs are not found in all tissues, but can be found in tissues that are known to undergo fatty degeneration due to new adipocyte production.   137 4.7. DEVELOPMENTAL ORIGINS OF AP CELLS Although it is accepted that adipogenic cells arise from mesenchymal precursors, very little is known about the specification and commitment to this lineage, largely because strategies to identify adipocyte precursors had not been developed (Billon et al., 2007; Gesta et al., 2007). Now that we have established a method to isolate APs from multiple adult tissues, it will be important to determine whether our techniques can be used to prospectively identify similar cells from primordial adipose tissues in the fetus or developing embryo. Identification of specific genetic markers for these cells will allow for the development of important tools, including lineage tracing strategies, which have already provided valuable information about the developmental relationships between fat and other tissues. For example, we now know that a subset of white fat (particularly around the head) arises from the neural crest, and that brown fat, which was recently thought to derive from a bi-potent white-brown fat progenitor, actually shares a common lineage with skeletal muscle and not white fat (Billon et al., 2007; Gesta et al., 2007; Seale et al., 2008). Using a similar strategy combined with the prospective identification of APs, others in our lab have confirmed that APs from different depots arise from different embryonic germ layers. Although the physiological relevance of these findings is unclear, this is currently under investigation. Finally, these results underscore the complexity underlying the ontogeny of adipose tissues. It is hoped that the contributions outlined in this thesis will lead an expansion of knowledge in this area.   138 4.8. ADIPOGENIC PROGENITORS OR ADIPOGENIC STEM CELLS? Stem cells are defined by the ability to differentiate and to self-renew in vivo. To date, self-renewal capacity has not been demonstrated in our adipogenic cell population; thus throughout this work I referred to our cells as adipogenic progenitors. Whether these cells are true stem cells capable of life-long contribution to de novo adipogenesis will have important implications on their effect on metabolism. Currently, others in the lab are investigating the in vivo self-renewal capacity of adipogenic progenitors isolated from different tissues and depots using serial transplantation studies similar to those performed in the field of hematopoiesis. Whether APs can self renew is unclear; nonetheless, based on the work described in Chapter 2 we expect to find depot-specific differences in this ability. Clearly, depots that contain primarily adipogenic stem cells rather than progenitors will have a greater capacity for new adipocyte production and tissue expandability, and thus be better able to maintain a pool of small, functional adipocytes. Furthermore, the utility of adipogenic cells for therapeutic transplantation (potentially for tissue reconstruction or correction of metabolic disorders), may benefit from using true adipogenic stem cells that generate ectopic fat depots capable of maintaining long- term tissue homeostasis.  Distinct cellular intermediates of adipocyte differentiation have been described in 3T3-F422A and 3T3-L1 cell lines; thus, it is feasible that similar intermediates exist in vivo (Gesta et al., 2007). Since our prospective isolation strategy purifies virtually all the adipogenic cells arising from fat and skeletal muscle cell preparations, we  139 may actually be isolating a heterogenous population containing several types of adipogenic precursors, including stem cells, at different stages of differentiation. In support of this notion, we observed a large variation in the adipogenic ability of colonies derived from single APs—some produced very few adipocytes, whereas others produced large ‘blast-like’ adipocyte colonies. We aim to improve on our purification strategy by introducing additional surface markers that may allow us to resolve these differences, which may also permit the isolation of true adipogenic stem cells. Finally, identifying distinct marker combinations that specifically label the cellular intermediates of adipogenesis will reveal even greater insights into the ontogeny and physiology of adipose depots, as well as of infiltrating adipocytes that arise in tissues undergoing fatty degeneration.   140 4.9. FINAL REMARKS Adipose tissue is often spoken of as a single organ, yet it is found in a multitude of anatomical sites, arises from different embryonic germ layers, can expand as a result of both physiological or pathological stimuli, and functions differently depending on where it is found. Since the field is dominated by work performed on immortalized preadipocyte cell lines, there has been a prevailing assumption that all adipocytes are the same. Although there are obvious similarities shared between all adipocytes, disparities in the ontogeny and physiology of different fat depots dictate that differences between their adipocytes must exist. Thus, physiology of adipose tissue is an extremely complex subject requiring careful consideration of the biology of each fat pad.  The work presented in this thesis describes a strategy to interrogate the adipogenic lineage, allowing for careful experimentation on cells fated to become adipocytes from multiple tissues. Importantly, our work uncovers a previously unforeseen link between stem cell biology and adipose tissue physiology. Remarkably, our strategy allows us to identify AP cells from anatomically and functionally distinct tissues using identical markers. Although we show that APs share a common fate, whether these cells are functionally identical, and exactly how they contribute to their resident tissue, remains to be seen. Nonetheless, the enabling technology described in this work begs intriguing questions, the answers to which may one day allow for the therapeutic manipulation of de novo adipogenesis, and may provide a source of adipocyte-forming cells for transplantation therapies.  141 4.10. FIGURES Figure 4.1. Proposed model for the effect of progenitor content and activity on fat depot expansion  (A) Fat depots containing low progenitor content and activity grow mainly by adipocyte hypertrophy, potentially resulting in dysfunctional adipocytes. (B) Fat depots containing high progenitor content and activity can grow by new adipocyte production, thereby maintaining a pool of smaller, functional adipocytes.     142 Figure 4.2. Models of stem/progenitor cell contribution to fibro/adipogenic infiltration of skeletal muscle.  (A) The current model of fibro/adipogenic infiltration in skeletal muscle dictates that multipotent satellite cells contribute to muscle regeneration in normal situations; however, prolonged exposure to the environment of diseased muscle instructs satellite cells to acquire the ability to differentiate into adipocytes and fibroblasts. (B) The proposed model of fibro/adipogenic infiltration in skeletal muscle, based on the work presented in this thesis, suggests that there are two distinct progenitors present in skeletal muscle at steady state: 1) the monopotent satellite cell, only capable of myogenic differentiation; and 2) the fibro/adipogenic progenitor (FAP), capable of differentiation into adipocytes and fibroblasts. When damage occurs in healthy tissue, satellite cell differentiation predominates and results in normal regeneration. When damage occurs in diseased tissue, FAP differentiation predominates and leads to fibro/adipocytic deposition and/or production of scar tissue.  143       144 Figure 4.3. Proposed model for the role of fibro/adipogenic progenitors in normal and impaired skeletal muscle regeneration  (A) At rest, fibro/adipogenic progenitors (FAPs) are found adjacent to blood vessels and satellite cells / myogenic progenitors (MPs) reside beneath the basal lamina, next to the myofiber. (B) 0-48 hours after muscle damage FAPs proliferate, and MPs become activated from their quiescent state. (C, D) Normal regeneration. MPs proliferate between 48-120 hours after damage, and FAPs release factors promoting myogenic commitment to terminal differentiation, including IGF-1 and IL-6 (C). Also during this time, FAPs stop proliferating— and preliminary data from our lab indicates that MPs may negatively regulate FAP expansion. After several weeks, the newly regenerated myofiber contains centrally located myonuclei (D). The resulting mechanical activity from the functional muscle tissue inhibits FAP differentiation. (E, F) Impaired regeneration. In the context of chronically diseased or aged skeletal muscle, MP proliferation is compromised. Thus, FAP expansion prevails (E). After several weeks, the lack of myofiber regeneration results in loss of mechanical activity (F). As a result, FAP differentiation is no longer inhibited and production of new adipocytes and fibroblasts ensues.  145     146 4.11. REFERENCES Billon, N., Iannarelli, P., Monteiro, M.C., Glavieux-Pardanaud, C., Richardson, W.D., Kessaris, N., Dani, C., and Dupin, E. (2007). The generation of adipocytes by the neural crest. Development 134, 2283-2292. Bluher, M., Michael, M.D., Peroni, O.D., Ueki, K., Carter, N., Kahn, B.B., and Kahn, C.R. (2002). Adipose tissue selective insulin receptor knockout protects against obesity and obesity-related glucose intolerance. Dev Cell 3, 25-38. Brack, A.S., Conboy, M.J., Roy, S., Lee, M., Kuo, C.J., Keller, C., and Rando, T.A. (2007). Increased Wnt signaling during aging alters muscle stem cell fate and increases fibrosis. Science 317, 807-810. Bradfute, S.B., Graubert, T.A., and Goodell, M.A. (2005). Roles of Sca-1 in hematopoietic stem/progenitor cell function. Exp Hematol 33, 836-843. Dellavalle, A., Sampaolesi, M., Tonlorenzi, R., Tagliafico, E., Sacchetti, B., Perani, L., Innocenzi, A., Galvez, B.G., Messina, G., Morosetti, R., et al. (2007). Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat Cell Biol 9, 255-267. Djian, P., Roncari, A.K., and Hollenberg, C.H. (1983). Influence of anatomic site and age on the replication and differentiation of rat adipocyte precursors in culture. J Clin Invest 72, 1200-1208. Gesta, S., Tseng, Y.H., and Kahn, C.R. (2007). Developmental origin of fat: tracking obesity to its source. Cell 131, 242-256. Gray, S.L., and Vidal-Puig, A.J. (2007). Adipose tissue expandability in the maintenance of metabolic homeostasis. Nutr Rev 65, S7-12. Greenwood, M.R., and Hirsch, J. (1974). Postnatal development of adipocyte cellularity in the normal rat. J Lipid Res 15, 474-483.  147 Hashimoto, T., Nobuhara, K., and Hamada, T. (2003). Pathologic evidence of degeneration as a primary cause of rotator cuff tear. Clin Orthop Relat Res, 111- 120. Hausman, G.J., Campion, D.R., and Martin, R.J. (1980). Search for the adipocyte precursor cell and factors that promote its differentiation. J Lipid Res 21, 657-670. Hughes, S.D., Verstuyft, J., and Rubin, E.M. (1997). HDL deficiency in genetially engineered mice requires elevated LDL to accelerated atherogenesis. Arterioscler. Thromb. Vasc. Biol. 17. 1725-1729. Jarvinen, M., Jozsa, L., Kannus, P., Jarvinen, T.L., Kvist, M., and Leadbetter, W. (1997). Histopathological findings in chronic tendon disorders. Scand J Med Sci Sports 7, 86-95. Johnson, P.R., Stern, J.S., Greenwood, M.R., and Hirsch, J. (1978). Adipose tissue hyperplasia and hyperinsulinemia on Zucker obese female rats: a developmental study. Metabolism 27, 1941-1954. Khan, T., Muise, E.S., Iyengar, P., Wang, Z.V., Chandalia, M., Abate, N., Zhang, B.B., Bonaldo, P., Chua, S., and Scherer, P.E. (2009). Metabolic dysregulation and adipose tissue fibrosis: role of collagen VI. Mol Cell Biol 29, 1575-1591. Kim, J.Y., van de Wall, E., Laplante, M., Azzara, A., Trujillo, M.E., Hofmann, S.M., Schraw, T., Durand, J.L., Li, H., Li, G., et al. (2007). Obesity-associated improvements in metabolic profile through expansion of adipose tissue. J Clin Invest 117, 2621-2637. Kirkland, J.L., Tchkonia, T., Pirtskhalava, T., Han, J., and Karagiannides, I. (2002). Adipogenesis and aging: does aging make fat go MAD? Exp Gerontol 37, 757-767. Miller, W.H., Jr., Faust, I.M., and Hirsch, J. (1984). Demonstration of de novo production of adipocytes in adult rats by biochemical and radioautographic techniques. J Lipid Res 25, 336-347.  148 Rajala, M.W., and Scherer, P.E. (2003). Minireview: The adipocyte--at the crossroads of energy homeostasis, inflammation, and atherosclerosis. Endocrinology 144, 3765-3773. Rodeheffer, M.S., Birsoy, K., and Friedman, J.M. (2008). Identification of White Adipocyte Progenitor Cells In Vivo. Cell 135, 240-249. Rosen CJ, Ackert-Bicknell CL, Adamo ML, Shultz KL, Rubin J, Donahue LR, Horton LG, Delahunty KM, Beamer WG, Sipos J, et al. (2004). Congenic mice with low serum IGF-I have increased body fat, reduced bone mineral density, and an altered osteoblast differentiation program. Bone 35, 1046–1058. Rosen, C.J., and Bouxsein, M.L. (2006). Mechanisms of disease: is osteoporosis the obesity of bone? Nat Clin Pract Rheumatol 2, 35-43. Seale, P., Bjork, B., Yang, W., Kajimura, S., Chin, S., Kuang, S., Scime, A., Devarakonda, S., Conroe, H.M., Erdjument-Bromage, H., et al. (2008). PRDM16 controls a brown fat/skeletal muscle switch. Nature 454, 961-967. Shefer, G., Wleklinski-Lee, M., and Yablonka-Reuveni, Z. (2004). Skeletal muscle satellite cells can spontaneously enter an alternative mesenchymal pathway. J Cell Sci 117, 5393-5404. Spalding, K.L., Arner, E., Westermark, P.O., Bernard, S., Buchholz, B.A., Bergmann, O., Blomqvist, L., Hoffstedt, J., Naslund, E., Britton, T., et al. (2008). Dynamics of fat cell turnover in humans. Nature 453, 783-787. Tang, W., Zeve, D., Suh, J.M., Bosnakovski, D., Kyba, M., Hammer, R.E., Tallquist, M.D., and Graff, J.M. (2008). White fat progenitor cells reside in the adipose vasculature. Science 322, 583-586. Uchiyama, Y., Miyama, K., Kataginri, T., Yamaguchi, A., Takamori, H., Nkashima, K., Sato, T., Suda, T. (1994). Adipose conversion is accelerated in bone marrow cells of congenitally osteoporotic SAMP6 mice. J Bone Miner Res 9 (Suppl 1), B365.  149 Vazquez-Vela, M.E., Torres, N., and Tovar, A.R. (2008). White adipose tissue as endocrine organ and its role in obesity. Arch Med Res 39, 715-728. Wallace, G.Q., and McNally, E.M. (2008). Mechanisms of Muscle Degeneration, Regeneration, and Repair in the Muscular Dystrophies. Annu Rev Physiol.  150 5. APPENDICES 5.1. APPENDIX A: SUPPLEMENTARY INFORMATION FOR CHAPTER 2  Appendix A contains additional data for Chapter 2, and a version of this appendix has been submitted for publication as supplementary information for Chapter 2.   151 5.1.1. Appendix A- Tables Table A1. Single lin- Sca-1+ CD34+ cell deposition  Lin- Sca-1+ CD34+ cells isolated from SAT or VAT were deposited into single wells of a 96 well plate at a frequency of 1 cell per well. Individual wells were visually inspected to ensure that only one cell was deposited per well. Cells were grown as described in the text, and after 21 days wells were scored for the presence of colonies and adipocytes.  Tissue Sort # of Wells # of colonies # with adipocytes  Subcutaneous Fat 1 180 16 2  2 180 6 2  3 180 10 4  Visceral Fat 1 180 4 1  2 160 3 0  3 360 5 1     152 5.1.2. Appendix A- Figures Figure A1. Detection of BrdU labeling in adipocyte nuclei by immuno-electron microscopy (IEM)  Fat tissue was harvested and prepared for IEM from animals that were placed on high fat diet HFD for 60 days, and administered BrdU (0.8mg/ml in drinking water and 100mg/kg IP every 2 days) during the last 10 days of HFD treatment. Anti-BrdU antibodies were detected using gold-labeled secondary antibodies. (A) Low magnification IEM image revealed typical adipocyte appearance, with a large lipid vacuole and a think layer of cytoplasm.  The presence of electron dense gold particles in the adipocyte nucleus unequivocally demonstrates BrdU incorporation. (B) High magnification IEM image from inset in (A) shows gold labeling at the periphery of the nucleus (asterisks). L:  lipid vacuole; c: cytoplasm; n: nucleus.  153      154 Figure A2. Confirmation of Sca-1 and CD34 stain specificity using isotype control antibodies  Stromovascular preparations from subcutaneous (A-C) or visceral (D-F) adipose tissue were gated as described in the text (Hoechstmid PIlo CD45- CD31- α7-). Isotype stains were the same as normal stains, except that one antibody was substituted with an isotype-matched, non-specific antibody. (A, D) Gated cells were analyzed for the expression of Sca-1 and CD34 (also shown in the far right panels of Fig. 1a and Fig. 1d). (B, E) Sca-1 isotype control. (C, F) CD34 isotype control.      155 Figure A3. Detection of BrdU labeling of endogenous lin- Sca-1+ CD34+ cells by flow cytometry  (A) 6-12 week old mice were placed on high fat diet HFD for 60days, and administered BrdU (0.8mg/ml in drinking water and 100mg/kg IP every 2 days) during the last 10 days of HFD treatment. Fat tissue was harvested at the end of the experiment, weighed, and then fractionated and prepared for flow cytometry. (B) APs were gated from SV cells as described in the text, and then analyzed for BrdU incorporation using a directly conjugated BrdU antibody. A robust BrdU signal is evident in BrdU-treated, but not control animals, which allowed the setting of precise gates for quantification.  156      157 Figure A4. Proliferation of non-adipogenic R3 (lin- Sca-1+ CD34+) cells is not correlated to fat depot size  Animals were placed on standard or high fat diet for 60 days.  During the last 10 days of treatment diet animals were administered BrdU by intraperitoneal injection (100mg/kg) every 2 days, and given BrdU in drinking water (0.08g/L in 20% sucrose). On day 60, SAT (A) and VAT (B) were harvested, weighed, and then prepared for multiparameter flow cytometry for surface marker staining and BrdU labeling. Correlation analysis of a non-adipogenic population (R3: lin-, Sca-1-, CD34-) taken from the same samples as those from Fig. 2.5 A and B, revealed no significant relationship between the frequency of BrdU+ R3 cells and the fat depot from which they came. The data presented in this figure is a control experiment for the data presented in Fig. 2.5 A, B.  158      159 5.2. APPENDIX B: SUPPLEMENTARY INFORMATION FOR CHAPTER 3  Appendix B contains additional data for Chapter 3, and a version of this appendix has been submitted for publication as supplementary information for Chapter 3.   160 5.2.1. Appendix B- Figures Figure B1. Confirmation of Sca-1 CD34 stain specificity using isotype-matched control antibodies  Skeletal muscle preparations were stained and gated as described in the text (Hoechstmid PIlo CD45- CD31-). Isotype stains were the same as normal stains except that one antibody was substituted with an isotype-matched, non-specific antibody, conjugated to the same fluorophore. (A) CD34 PE Isotype control (B) Sca-1 PECy7 Isotype control      161 Figure B2. Linear regression of colony forming data from freshly isolated skeletal muscle DN (Sca-1- CD34-) cells  Hoechstmid PIlo CD45- CD31- Sca-1- CD34- cells were sorted and seeded at the indicated densities in a 5cm matrigel-coated dish and cultivated in growth medium for 2 weeks.  After fixation, colonies (defined as clusters of more than 10 cells) were counted, and then plotted against the number of cells inoculated. Dashed lines represent the 95% confidence regression band. Clonogenicity was determined from the slope of the curve, and the 95% confidence interval is shown in brackets.      162 Figure B3. Skeletal muscle DP (Sca-1+ CD34+) cells do not form bone or cartilage.  Cells were previously gated for Hoechstmid PIlo CD45- CD31-. (A) DP cells were grown in osteogenic conditions for 10 weeks, and then stained with alizarin red for visualization of calcified nodules. Despite the presence of alizarin red staining, we observed no areas that could be identified as osteogenic nodules. Sparse adipocytes were observed throughout the cultures. Scale bar at 50µm. (B) DP cell pellets were grown in chondrogenic conditions for 8 weeks, cryosectioned (10µm), and then stained for glycosaminoglycans with alcian blue. Although non-specific staining could be observed on the periphery of the pellet, no staining was observed throughout the pellet, and no characteristic chondrocyte morphology was observed. (Scale bar at 50µm).       163 Figure B4. Quantitative gene expression in sorted cells  Muscle-resident adipogenic (lin- Sca-1+ CD34+) and myogenic (lin- Sca-1- CD34+) cells were sorted and cultured as described in text. Differentiated cells arose spontaneously in growth media after >2 weeks culture, and did not require additional factors. qPCR analyses was performed using Taqman probe and primer sets spanning exon-exon boundaries. All values are expressed relative to TBP expression. ND= no signal detected. (A) Freshly isolated adipogenic cells express preadipocyte markers (B) Adipogenic cells express early markers of adipogenesis immediately after sorting, and markers of mature adipocytes after terminal differentiation. (C) Adipogenic cells do not express skeletal muscle or brown fat genes (D) Myogenic cells express markers common to satellite cells immediately after sorting, and markers of mature and regenerating muscle after terminal differentiation. (E) Myogenic cells do not express genes common to white adipocytes.  164      165 Figure B5. An alternative strategy using α7 integrin to identify adipogenic and myogenic cells from dissociated skeletal muscle  Skeletal muscle preparations were gated for Hoechstmid PIlo CD45- CD31- (lin-) cells, and then analyzed for the expression of Sca-1 and α7 integrin. Lin- Sca-1- α7- cells did not differentiate into myosin heavy chain (MyHC)-expressing myotubes (grey gate, left image), revealing that all myogenic activity was found exclusively in the lin- α7+ Sca-1- population (red gate, middle image). All adipogenic activity was found exclusively in the lin- α7- Sca-1+ population (blue gate, right image), and over 99% of these cells expressed CD34. Greyed out histogram represents the CD34-PE isotype control. All scale bars at 100µm.  166  Figure B6. Co-cultivation control experiment  This figure shows data from a control experiment for Figure 3.3A. Equal numbers of MPs sorted from transgenic GFP+ and wildtype animals were co-cultivated for 14 days. Confocal microscopy confirmed participation of both GFP+ and GFP- MPs to the myogenic lineage. Scale bar at 100µm. (A) GFP+ myotubes (B) Myosin Heavy Chain immunostain (C) Merge       167 Figure B7. Sca-1+ mononuclear cells are juxtaposed to skeletal muscle fibers after damage  Numerous Sca-1+ mononuclear cells are found adjacent to regenerating myofibers in areas of severe damage. Tissues were harvested 3 days after NTX damage induction, fixed, cryosectioned, then immunostained for CD31 and Sca-1. A single optical slice from a confocal Z-stack is shown. Background autofluorescence from myofibers is shown in the green channel. Scale bar at 25µm. (A) Nuclear staining using TOTO3 (B) Sca-1 immunostaining (C) Merge        168 5.3. APPENDIX C. PRELIMINARY DATA  Appendix C contains preliminary data discussed in Chapter 4.   169 5.3.1. Appendix C- Figures Figure C1. The effect of aging on adipogenic progenitor (AP) abundance in subcutaneous and visceral fat depots  Tissues were harvested from young (< 4 months) and old (>20 months) B6 mice. Adipogenic progenitors were identified as Hoechstmid PIlo CD45- CD31- α7- (lin-) Sca- 1+ CD34+ cells, and were enumerated by flow cytometry using beads. Data for young animals came from one experiment generated from pooled tissue preparations of 3 animals. Data for old animals came from two experiments, each arising from pooled tissue preparations of 2-3 animals.       170 Figure C2. The effect of aging on adipogenic (AP) and myogenic (MP) progenitor abundance in skeletal muscle  Skeletal muscle was harvested from young (< 4 months) and old (>20 months) B6 mice and prepared for flow cytometry. Adipogenic progenitors were identified as Hoechstmid PIlo CD45- CD31- (lin-) Sca-1+ CD34+ cells, and myogenic progenitors as lin- Sca-1- CD34+ cells. (A) The effect of aging on the ratio of APs to MPs, as determined from flow cytometry data. (B) The effect of aging on the frequency of APs (left) and MPs (right), determined as the percentage of lin- cells.  171        172 Figure C3. The prospective isolation of adipogenic progenitor cells from tendon  (A) Viable cells from digested tendon tissue were identified based on forward/side scatter, medium intensity Hoechst staining (to exclude debris) and low intensity PI staining (to exclude dead or dying cells). Markers for hematopoietic (CD45+), endothelial (CD31+) and smooth/skeletal muscle (α7 integrin+) cells were used to exclude mature lineages from analysis (resulting cells called lin-). (B) Isotype control experiments to verify staining specificity. Isotype stains were the same as normal stains except that one antibody was substituted with an isotype- matched, non-specific antibody, conjugated to the same fluorophore. The plots shown are from the lin- gate of a anti-CD34 isotype control (left) and anti-Sca-1 isotype control (right). (C) Lin- cells from tendon preparations were analyzed for Sca-1 and CD34 expression and labeled P1 (Sca-1+ CD34-), P2 (Sca-1+ CD34+) and P3 (Sca-1- CD34-). Sorted cells were cultured for 10 days in growth medium, then switched to adipogenic conditions for 2 weeks. Adipogenic differentiation was detected exclusively in P2.  173       174 5.4. APPENDIX D. UBC RESEARCH ETHICS BOARD CERTIFICATES OF APPROVAL  Appendix D contains the University of British Columbia Animal Care Committee Certificate of Approval for projects performed under guidance of Dr. Fabio Rossi, my PhD Supervisor.  175   176   177   178 

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