Aim 4: To investigate the functional interaction between white adipocytes and HDLECs using complex in vitro models with the special focus on the handling of lipids and VEGF-C activation.
The coculture model, which enables interactions between different cell types, represents a promising tool to create a more physiologically relevant environment that allows a closer understanding of the complex interconnections between cells within tissues. Indirect and direct cell-to-cell contact can engage, turn on/off gene pathways, or modulate cellular stimulation and responses [316, 317].
To better understand cellular interconnections between AT and LS, we have developed human in vitro single and coculture models between white adipocytes and HDLECs. Our results of single culture have shown that secretion products of adipocytes stimulated to lipolysis increase mRNA levels of lipid handling genes (FABP4, CPT1A, CD36) without obvious changes of lymphatic markers in HDLECs. This result goes hand in hand with results of study by Morfoisse et al. [204] showing that exposition of HDLECs to media conditioned by adipocytes elevated gene expression of lipid handling genes together with enhancement of lymphatic sprouting (but not proliferation). Compared to this study, our experimental set up was improved by the generation of adipocyte CM free of the primary lipolytic stimulus (dbcAMP), which could activate cAMP-regulated pathways also in HDLECs [318, 319] and thus bias the obtained results. We also confirmed that lipids provided by adipocytes are uptaken and stored by HDLEC, as shown in our coculture model kept in serum free conditions, where the majority of available FAs are produced de novo by adipocytes. Based on these results, it could be expected that HDLECs can stimulate adipocytes to provide them necessary FFA. These putative stimuli probably affect only basal lipolysis, as CM from HDLEC was unable to enhance lipolysis induced by isoproterenol.
Moreover, cell-to-cell contacts in direct coculture model also significantly increased basal lipolysis. These results thus support our hypothesis that HDLECs are able to stimulate the release of lipolytic substrates from adipocytes. Indeed, Monelli et al. [65] have recently demonstrated the ability of ECs to stimulate lipolysis in vivo in AT via local production of polyamines. Also, our finding of higher basal lipolysis in AT explants from lymphedema patients, whose lymphangiogenesis is enhanced yet unproductive [185], support this
118 functional adipocyte-LECs interplay. Therefore, it can be assumed that LECs may possess, (similary like ECs) lipolytic trigger/s to enhance lipolysis of adipocytes and then use the released FFA to provide energy for lymphangiogenesis. Therefore, new investigations regarding the exact molecular pathway engaged by LECs to stimulte lipolysis are more than needed.
Furthermore, we have observed various cellular structures in our in vitro models, confirming the remarkable plasticity of both adipocytes and HDLECs [43, 101, 254]. For example, in the model of indirect coculture, we have observed the formation of TNTs among HDLECs.
In the direct coculture, it is possible that TNTs could transport lipid droplets between HDLECs (nurtured by adipocytes), similarly as seen in HUVECs [281] but also directly between adipocytes and HDLECs. Thus, TNTs could facilitate the formation of LDs as observed in the cytoplasm of HDLECs in the direct coculture model. This fascinating but purely speculative idea should indeed be tested in future experiments.
Also, the lymphangiogenic process promoted by VEGF-C stimulated the lipid depletion/migration of adipocytes in close contact with the emerging lymphatic vasculature.
This phenomenon was not described before and thus we can only speculate about the pathways necessary to stimulate it. The direct effect of VEGF-C might be excluded, as adipocytes express only very low or no VEGR-2 or -3 mRNA levels (not shown). Recently, LECs have been shown to produce cytokines that dirrectly affect the metabolism of brown adipocytes [211]. Therefore we have also tested the presence of the neurotensin receptor in white adipocytes in vitro, but its mRNA level was undetectable (not shown). Although mature adipocytes are able to migrate, as shown in Drosophila model [320], they migrate towards a source of cytokines (f.e. induced by wounding of tissue), while in our model they would have to be propably migrate away from the source of putative HDLEC-borne stimulus. One possible explanation could be the repulsion of adipocytes by bulky hyaluronan glycocalyx bound to HDLEC via LYVE-1. Hyaluronan is produced by adipocytes and represents a part of ECM that is actively connected to LVs [321]. However, it is not known whether VEGF-C signaling could initiate such profound changes in the glycocalyx resulting in the repulsion of adipocytes. Also, it could be rather the lipid depletion of adipocytes, than the migration. In that case, the direct cell-to-cell interactions are probably crucial to induce the excessive lipolysis. Since all these mentioned hypotheses are speculative, the live cell imaging experiments focused on the behavior of lipid-laden adipocytes could provide important evidence to either support or disprove them.
119 As described above, VEGF-C is a major inducer of lymphangiogenesis. Our results confirm its irreplaceable role during lymphangiogenesis, as lymphatic network formation was observed only in the presence of VEGF-C, whereas the absence of VEGF-C led to the formation of lymphatic clumps of different sizes and shapes during coculturing. On the other hand, HDLEC monocultures in the absence or presence of VEGF-C displayed no morphological alterations. The absence of morphological changes may be explained by the presence of a non-supportive matrix.
As VEGF-C circulates in the blood primarily in inactive pro-VEGF-C form, its proteolytic activation in the local environment is necessary to stimulate lymphangiogenesis. This brought an interesting question: could adipocytes support lymphangiogenesis via the production of suitable VEGF-C proteases? Besided ADAMTS3, an important protease for embryonic cleavage of inactive form of VEGF-C [202]. ADAMTS2 and ADAMTS14 are effective surrogates and play an active role in the proteolytic cleavage of pro-VEGF-C in adults [203]. Based on published single nucleus RNAseq data from human AT [2], we found that each of the adipocyte subsets produced relatively high levels of ADAMTS2 compared with ADAMTS14 or ADAMTS3. However, our results have shown no significant mRNA changes of proteases-mediated activation of VEGF-C during in vitro lipolysis. The necessary proof of concept must be applied via the detection of mature VEGF-C after incubation of pro-VEGF-C with medium enriched by secretory products of adipocytes or directly with adipocytes. This experiment has already been performed, but we are still waiting for a suitable anti-VEGF-C antibody needed to complete the analysis.
Although we did not observe a significant increase in protease mRNA levels during in vitro lipolysis. All analyzed proteases showed a tendency to change mRNA levels in response to insulin. Insulin has not only metabolic function but it can also stimulate angiogenesis linked with AT expansion. AT expansion requires ECM remodeling, which is highly dependent on the biological activities of extracellular proteases [322]. Therefore, it is conceivable that increasing the expression of ADAMTS proteases in response to insulin could contribute not only to ECM remodeling but also to promoting lymphangiogenesis, which appears as important as angiogenesis for AT expansion. As studies analyzing the behavior of the ADAMTS family proteases in AT are scarce, [323-325], our pilot data thus open new perspectives in the search for regulators of AT expansion.
In conclusion, this work has provided new evidence for a close functional interaction between the cellular components of LS and AT.
120
8 CONCLUSIONS
This thesis contributed to the analysis of functional interplay between structures and components of human AT and LS in pathological and normal states. Part of my thesis provided insight into how AT and adipocytes alter in response to the dysfunction of LS expressed by the development of lymphedema. The next part of thesis was dedicated to the analysis of indirect and direct interconnections between cellular components of AT and LS, i.e., adipocytes and LECs, in vitro.
The major conclusions of this thesis are:
Analysis of the role of lymphatic drainage on the lipolytic activity in women showed that less effective lymphatic drainage is accompanied with lower lipolytic reactivity in femoral AT with significant differences in the distribution of AT metabolites between the AT interstitium and circulation, respectively.
Comparison of healthy and lymphedema associated AT from upper limb of women revealed higher expression of lymphangiogenic markers, angiogenic capacity, basal lipolysis and lower FFA re-esterification in LAT. These identified alterations provide a basis for the hypothesis that AT affected by lymphedema is characterized by an unsuccessful effort to restore lymphatic drainage and limit further lipid deposition.
Analysis of the impact of in vitro application of lymphedema-derived fluid on adipogenesis of human preadipocytes suggested that one of the external stimuli leading to AT accumulation affected by lymphedema is present in non-drained lymph/ISF rich in lipolytic products and miRNAs.
Examination of the functional interaction of in vitro coculture model showed that HDLECs stimulated basal lipolysis in adipocytes, when FFA released by adipocytes were uptaken and stored by HDLEC. Adipocytes supported lymphangiogenesis in HDLEC in the presence of VEGF-C. These results support the existence of functional interplay between HDLECs and human adipocytes.
121
9 REFERENCE
1. Tanaka, M. and Y. Iwakiri, The Hepatic Lymphatic Vascular System: Structure, Function, Markers, and Lymphangiogenesis. Cell Mol Gastroenterol Hepatol, 2016. 2(6): p. 733-749.
2. Emont, M.P., et al., A single-cell atlas of human and mouse white adipose tissue. Nature, 2022. 603(7903): p. 926-933.
3. Wells, H.G., Adipose tissue, a neglected subject J Am Med Assoc 1940. 114(22).
4. Yu, P., et al., Adipose tissue, aging, and metabolism. Current Opinion in Endocrine and Metabolic Research, 2019. 5: p. 11-20.
5. Cinti, S., Adipose Organ Development and Remodeling. Compr Physiol, 2018. 8(4): p. 1357- 1431.
6. Wang, T., A.K. Sharma, and C. Wolfrum, Novel insights into adipose tissue heterogeneity.
Rev Endocr Metab Disord, 2022. 23(1): p. 5-12.
7. De Fano, M., et al., Adipose Tissue Plasticity in Response to Pathophysiological Cues: A Connecting Link between Obesity and Its Associated Comorbidities. Int J Mol Sci, 2022.
23(10).
8. Efthymiou, V. and M.E. Patti, It Is Not Just Fat: Dissecting the Heterogeneity of Adipose Tissue Function. Curr Diab Rep, 2022. 22(4): p. 177-187.
9. Bourgeois, C., et al., Specific Biological Features of Adipose Tissue, and Their Impact on HIV Persistence. Front Microbiol, 2019. 10: p. 2837.
10. Symonds, M.E., Adipose Tissue Biology Book 2017.
11. Saely, C.H., K. Geiger, and H. Drexel, Brown versus white adipose tissue: a mini-review.
Gerontology, 2012. 58(1): p. 15-23.
12. Sell, H., Y. Deshaies, and D. Richard, The brown adipocyte: update on its metabolic role. Int J Biochem Cell Biol, 2004. 36(11): p. 2098-104.
13. Stanford, K.I., et al., 12,13-diHOME: An Exercise-Induced Lipokine that Increases Skeletal Muscle Fatty Acid Uptake. Cell Metab, 2018. 27(5): p. 1111-1120 e3.
14. Villarroya, F., et al., Inflammation of brown/beige adipose tissues in obesity and metabolic disease. J Intern Med, 2018. 284(5): p. 492-504.
15. Yang, F.T. and K.I. Stanford, Batokines: Mediators of Inter-Tissue Communication (a Mini- Review). Curr Obes Rep, 2022. 11(1): p. 1-9.
16. Pond, C.M., The Evolution of Mammalian Adipose Tissues, in Adipose Tissue Biology. 2017.
p. 1-59.
17. Guerrero-Juarez, C.F. and M.V. Plikus, Emerging nonmetabolic functions of skin fat. Nat Rev Endocrinol, 2018. 14(3): p. 163-173.
18. Brown, N.K., et al., Perivascular adipose tissue in vascular function and disease: a review of current research and animal models. Arterioscler Thromb Vasc Biol, 2014. 34(8): p. 1621- 30.
19. Zaborska, K.E., et al., Loss of anti-contractile effect of perivascular adipose tissue in offspring of obese rats. Int J Obes (Lond), 2016. 40(8): p. 1205-14.
20. Lee, Y.C., et al., Role of perivascular adipose tissue-derived methyl palmitate in vascular tone regulation and pathogenesis of hypertension. Circulation, 2011. 124(10): p. 1160-71.
21. Westcott, E.D., et al., Perinodal adipose tissue and fatty acid composition of lymphoid tissues in patients with and without Crohn's disease and their implications for the etiology and treatment of CD. Ann N Y Acad Sci, 2006. 1072: p. 395-400.
22. Westcott, E., et al., Fatty acid compositions of lipids in mesenteric adipose tissue and lymphoid cells in patients with and without Crohn's disease and their therapeutic implications. Inflammatory Bowel Diseases, 2005. 11(9): p. 820-827.
122 23. Knight, S.C., Specialized perinodal fat fuels and fashions immunity. Immunity, 2008. 28(2):
p. 135-8.
24. Lago, M.E.L., et al., Skin in vitro models to study dermal white adipose tissue role in skin healing, in Skin Tissue Models for Regenerative Medicine. 2018. p. 327-352.
25. Festa, E., et al., Adipocyte lineage cells contribute to the skin stem cell niche to drive hair cycling. Cell, 2011. 146(5): p. 761-71.
26. Li, X., et al., Perivascular adipose tissue-derived extracellular vesicle miR-221-3p mediates vascular remodeling. FASEB J, 2019: p. fj201901548R.
27. Yu, W., et al., Bone marrow adipogenic lineage precursors promote osteoclastogenesis in bone remodeling and pathologic bone loss. J Clin Invest, 2021. 131(2).
28. Tencerova, M., M. Ferencakova, and M. Kassem, Bone marrow adipose tissue: Role in bone remodeling and energy metabolism. Best Pract Res Clin Endocrinol Metab, 2021. 35(4): p.
101545.
29. Tchernof, A., Visceral Adipocytes and the Metabolic Syndrome. Nutrition Reviews, 2007.
65(6): p. 24-29.
30. Freedland, E.S., Role of a critical visceral adipose tissue threshold (CVATT) in metabolic syndrome: implications for controlling dietary carbohydrates: a review. Nutr Metab (Lond), 2004. 1(1): p. 12.
31. Bodis, K. and M. Roden, Energy metabolism of white adipose tissue and insulin resistance in humans. Eur J Clin Invest, 2018. 48(11): p. e13017.
32. Lamarche, B., et al., Visceral obesity and the risk of ischaemic heart disease: insights from the Quebec Cardiovascular Study. Growth Horm IGF Res, 1998. 8 Suppl B: p. 1-8.
33. Onat, A., et al., Measures of abdominal obesity assessed for visceral adiposity and relation to coronary risk. Int J Obes Relat Metab Disord, 2004. 28(8): p. 1018-25.
34. Ibrahim, M.M., Subcutaneous and visceral adipose tissue: structural and functional differences. Obes Rev, 2010. 11(1): p. 11-8.
35. Frayn, K.N., Adipose tissue as a buffer for daily lipid flux. Diabetologia, 2002. 45(9): p. 1201- 10.
36. Palmer, B.F. and D.J. Clegg, The sexual dimorphism of obesity. Mol Cell Endocrinol, 2015.
402: p. 113-9.
37. White, U.A. and Y.D. Tchoukalova, Sex dimorphism and depot differences in adipose tissue function. Biochim Biophys Acta, 2014. 1842(3): p. 377-92.
38. Zwick, R.K., et al., Anatomical, Physiological, and Functional Diversity of Adipose Tissue. Cell Metab, 2018. 27(1): p. 68-83.
39. Gruss, L.T. and D. Schmitt, The evolution of the human pelvis: changing adaptations to bipedalism, obstetrics and thermoregulation. Philos Trans R Soc Lond B Biol Sci, 2015.
370(1663): p. 20140063.
40. McQuaid, S.E., et al., Femoral adipose tissue may accumulate the fat that has been recycled as VLDL and nonesterified fatty acids. Diabetes, 2010. 59(10): p. 2465-73.
41. Evans, J., et al., Depot- and ethnic-specific differences in the relationship between adipose tissue inflammation and insulin sensitivity. Clin Endocrinol (Oxf), 2011. 74(1): p. 51-9.
42. Piche, M.E., et al., Relevance of human fat distribution on lipid and lipoprotein metabolism and cardiovascular disease risk. Curr Opin Lipidol, 2018. 29(4): p. 285-292.
43. Manolopoulos, K.N., F. Karpe, and K.N. Frayn, Gluteofemoral body fat as a determinant of metabolic health. Int J Obes (Lond), 2010. 34(6): p. 949-59.
44. Buendia, R., et al., Estimation of Arm Adipose Tissue Quotient Using Segmental Bioimpedance Spectroscopy. Lymphat Res Biol, 2018. 16(4): p. 377-384.
45. Crescenzi, R., et al., Upper and Lower Extremity Measurement of Tissue Sodium and Fat Content in Patients with Lipedema. Obesity (Silver Spring), 2020. 28(5): p. 907-915.
46. Deptula, P., et al., Autologous Fat Grafting in the Upper Extremity: Defining New Indications.
Plast Reconstr Surg Glob Open, 2022. 10(8): p. e4469.
123 47. Kane, H. and L. Lynch, Innate Immune Control of Adipose Tissue Homeostasis. Trends
Immunol, 2019. 40(9): p. 857-872.
48. Bapat, S.P., Y. Liang, and Y. Zheng, Characterization of Immune Cells from Adipose Tissue.
Curr Protoc Immunol, 2019. 126(1): p. e86.
49. Cinti, S., et al., Adipocyte death defines macrophage localization and function in adipose tissue of obese mice and humans. J Lipid Res, 2005. 46(11): p. 2347-55.
50. Chung, K.J., et al., Innate immune cells in the adipose tissue. Rev Endocr Metab Disord, 2018. 19(4): p. 283-292.
51. Weisberg, S.P., et al., Obesity is associated with macrophage accumulation in adipose tissue. Journal of Clinical Investigation, 2003. 112(12): p. 1796-1808.
52. Gersh, I. and M.A. Still, Blood Vessels in Fat Tissue. Relation to Problems of Gas Exchange. J Exp Med, 1945. 81(2): p. 219-32.
53. Varzaneh, F.E., et al., Extracellular matrix components secreted by microvascular endothelial cells stimulate preadipocyte differentiation in vitro. Metabolism, 1994. 43(7): p.
906-12.
54. Corvera, S. and O. Gealekman, Adipose tissue angiogenesis: impact on obesity and type-2 diabetes. Biochim Biophys Acta, 2014. 1842(3): p. 463-72.
55. Monelli, E., et al., Angiocrine polyamine production regulates adiposity. Nat Metab, 2022.
4(3): p. 327-343.
56. Wosnitza, M., et al., Plasticity of human adipose stem cells to perform adipogenic and endothelial differentiation. Differentiation, 2007. 75(1): p. 12-23.
57. Hirsch, J. and E. Gallian, Methods for the determination of adipose cell size in man and animals. Journal of Lipid Research, 1968. 9(1): p. 110-119.
58. Stenkula, K.G. and C. Erlanson-Albertsson, Adipose cell size: importance in health and disease. Am J Physiol Regul Integr Comp Physiol, 2018. 315(2): p. R284-R295.
59. Arner, E., et al., Adipocyte turnover: relevance to human adipose tissue morphology.
Diabetes, 2010. 59(1): p. 105-9.
60. Bernstein, R.S., N. Grant, and D.M. Kipnis, Hyperinsulinemia and Enlarged Adipocytes in Patients with Endogenous Hyperlipoproteinemia without Obesity or Diabetes-Mellitus.
Diabetes, 1975. 24(2): p. 207-213.
61. Gumbilai, V., et al., Fat Mass Reduction With Adipocyte Hypertrophy and Insulin Resistance in Heterozygous PPARgamma Mutant Rats. Diabetes, 2016. 65(10): p. 2954-65.
62. Muir, L.A., et al., Adipose tissue fibrosis, hypertrophy, and hyperplasia: Correlations with diabetes in human obesity. Obesity (Silver Spring), 2016. 24(3): p. 597-605.
63. Jernas, M., et al., Separation of human adipocytes by size: hypertrophic fat cells display distinct gene expression. FASEB J, 2006. 20(9): p. 1540-2.
64. Honecker, J., et al., Transcriptome and fatty-acid signatures of adipocyte hypertrophy and its non-invasive MR-based characterization in human adipose tissue. EBioMedicine, 2022.
79: p. 104020.
65. Verboven, K., et al., Abdominal subcutaneous and visceral adipocyte size, lipolysis and inflammation relate to insulin resistance in male obese humans. Sci Rep, 2018. 8(1): p. 4677.
66. Veilleux, A., et al., Visceral adipocyte hypertrophy is associated with dyslipidemia independent of body composition and fat distribution in women. Diabetes, 2011. 60(5): p.
1504-11.
67. Spalding, K.L., et al., Dynamics of fat cell turnover in humans. Nature, 2008. 453(7196): p.
783-787.
68. Drolet, R., et al., Hypertrophy and hyperplasia of abdominal adipose tissues in women. Int J Obes (Lond), 2008. 32(2): p. 283-91.
69. Tchoukalova, Y.D., et al., Regional differences in cellular mechanisms of adipose tissue gain with overfeeding. Proc Natl Acad Sci U S A, 2010. 107(42): p. 18226-31.
124 70. Bluher, M., The distinction of metabolically 'healthy' from 'unhealthy' obese individuals.
Curr Opin Lipidol, 2010. 21(1): p. 38-43.
71. Hardy, O.T., M.P. Czech, and S. Corvera, What causes the insulin resistance underlying obesity? Curr Opin Endocrinol Diabetes Obes, 2012. 19(2): p. 81-7.
72. White, U., R.A. Beyl, and E. Ravussin, A higher proportion of small adipocytes is associated with increased visceral and ectopic lipid accumulation during weight gain in response to overfeeding in men. Int J Obes (Lond), 2022. 46(8): p. 1560-1563.
73. Fukumura, D., et al., Paracrine regulation of angiogenesis and adipocyte differentiation during in vivo adipogenesis. Circ Res, 2003. 93(9): p. e88-97.
74. Kunduzova, O., et al., Apelin/APJ signaling system: a potential link between adipose tissue and endothelial angiogenic processes. FASEB J, 2008. 22(12): p. 4146-53.
75. Scotece, M., et al., Adiponectin and leptin: new targets in inflammation. Basic Clin Pharmacol Toxicol, 2014. 114(1): p. 97-102.
76. Yang, W.H., et al., Leptin promotes VEGF-C production and induces lymphangiogenesis by suppressing miR-27b in human chondrosarcoma cells. Sci Rep, 2016. 6: p. 28647.
77. Rodriguez, A., Novel molecular aspects of ghrelin and leptin in the control of adipobiology and the cardiovascular system. Obes Facts, 2014. 7(2): p. 82-95.
78. Beltowski, J., Leptin decreases plasma paraoxonase 1 (PON1) activity and induces oxidative stress: the possible novel mechanism for proatherogenic effect of chronic hyperleptinemia.
Atherosclerosis, 2003. 170(1): p. 21-29.
79. Ibrahim, H.S., et al., Leptin increases blood pressure and markers of endothelial activation during pregnancy in rats. Biomed Res Int, 2013. 2013: p. 298401.
80. Adya, R., B.K. Tan, and H.S. Randeva, Differential effects of leptin and adiponectin in endothelial angiogenesis. J Diabetes Res, 2015. 2015: p. 648239.
81. Ouchi, N., et al., Association of hypoadiponectinemia with impaired vasoreactivity.
Hypertension, 2003. 42(3): p. 231-4.
82. Chen, H., et al., Adiponectin stimulates production of nitric oxide in vascular endothelial cells. J Biol Chem, 2003. 278(45): p. 45021-6.
83. Sorhede Winzell, M., C. Magnusson, and B. Ahren, The apj receptor is expressed in pancreatic islets and its ligand, apelin, inhibits insulin secretion in mice. Regul Pept, 2005.
131(1-3): p. 12-7.
84. Dray, C., et al., Apelin stimulates glucose utilization in normal and obese insulin-resistant mice. Cell Metab, 2008. 8(5): p. 437-45.
85. Gerst, F., et al., What role do fat cells play in pancreatic tissue? Mol Metab, 2019. 25: p. 1- 10.
86. Ambele, M.A., et al., Adipogenesis: A Complex Interplay of Multiple Molecular Determinants and Pathways. Int J Mol Sci, 2020. 21(12).
87. Rahim, F., et al., Runx2 silencing promotes adipogenesis via down-regulation of DLK1 in chondrogenic differentiating MSCs. J Gene Med, 2020. 22(11): p. e3244.
88. Gupta, R.K., et al., Transcriptional control of preadipocyte determination by Zfp423. Nature, 2010. 464(7288): p. 619-23.
89. Pei, H., et al., Kruppel-like factor KLF9 regulates PPARgamma transactivation at the middle stage of adipogenesis. Cell Death Differ, 2011. 18(2): p. 315-27.
90. Challa, T.D., et al., Regulation of De Novo Adipocyte Differentiation Through Cross Talk Between Adipocytes and Preadipocytes. Diabetes, 2015. 64(12): p. 4075-87.
91. Luo, X., et al., Identification of BMP and activin membrane-bound inhibitor (BAMBI) as a potent negative regulator of adipogenesis and modulator of autocrine/paracrine adipogenic factors. Diabetes, 2012. 61(1): p. 124-36.
92. Rosen, E.D., et al., Transcriptional regulation of adipogenesis. Genes Dev, 2000. 14(11): p.
1293-307.