Adipose Tissue in Health and Disease

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Edited by Todd Leff and James G. Granneman Adipose Tissue in Health and Disease

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Edited by Todd Leff and James G. Granneman

Adipose Tissue in Health and Disease

The Editors Todd Leff Wayne State University Department of Pathology 540 E. Canﬁeld Detroit, MI 48201 USA

All books published by Wiley-VCH are carefully produced. Nevertheless, authors, editors, and publisher do not warrant the information contained in these books, including this book, to be free of errors. Readers are advised to keep in mind that statements, data, illustrations, procedural details or other items may inadvertently be inaccurate. Library of Congress Card No.: applied for

James G. Granneman Wayne State University Department of Psychiatry 550 E. Canﬁeld Detroit, MI 48201 USA Cover Whole mount micrograph of brown adipose tissue. Capillaries (red) were labeled with Griffonia simplicifolia I-B4 isolectin and brown adiipocytes were labeled with antibodies to perilipin (green). The image is a 3D projection of numerous confocal optical slices. Printed with kind permission by J. Granneman.

British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library. Bibliographic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliograﬁe; detailed bibliographic data are available on the Internet at http://dnb.d-nb.de. # 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim All rights reserved (including those of translation into other languages). No part of this book may be reproduced in any form – by photoprinting, microﬁlm, or any other means – nor transmitted or translated into a machine language without written permission from the publishers. Registered names, trademarks, etc. used in this book, even when not speciﬁcally marked as such, are not to be considered unprotected by law. Cover Design Adam Design, Weinheim Typesetting Thomson Digital, Noida, India Printing and Binding betz-druck GmbH, Darmstadt Printed in the Federal Republic of Germany Printed on acid-free paper ISBN: 978-3-527-31857-5

V

Contents Preface XIX List of Contributors

XXI

Part One Adipose Tissue Development and Morphology 1 1 1.1 1.2 1.2.1 1.2.2 1.2.3 1.2.4 1.2.5 1.2.6 1.3 1.3.1 1.3.2 1.3.3 1.3.4 1.3.5

2

2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.2

Transcriptional Control of Adipogenesis and Fat Cell Gene Expression Ursula A. White and Jacqueline M. Stephens Introduction 3 Transcriptional Control of Adipogenesis 4 AP-1 Transcription Factors 4 Signal Transducers and Activators of Transcription 5 Krüppel-Like Factors 6 SREBPs 7 C/EBP 7 PPAR-c 8 Identiﬁcation of Adipocyte Transcription Factor Target Genes 9 C/EBP Target Genes 9 SREBP-1 Target Genes 10 PPAR-c Target Genes 11 STAT-5 Target Genes 12 Summary 13 References 14 Cellular and Molecular Basis of Functional Differences among Fat Depots 21 Thomas Thomou, Tamara Tchkonia, and James L. Kirkland Introduction 21 Fat Tissue Function 21 Diversity in Fat Distribution 23 Regional Differences in Fat Tissue Growth 24 Disease Associations 26 Physiology 27

3

VI

Contents

2.2.1 2.2.2 2.2.3 2.2.4 2.2.5 2.2.6 2.2.7 2.3

3 3.1 3.2 3.3 3.3.1 3.3.2 3.4 3.5 3.6 3.6.1 3.6.2 3.6.3

3.6.4 3.7

4 4.1 4.2 4.3 4.3.1 4.4 4.4.1

Contribution of Inherent Cell Dynamic Mechanisms to Regional Differences 28 Preadipocyte Function 29 Preadipocyte Replication 29 Differences in Adipogenesis among Depots 30 Regional Variation in Susceptibility to Apoptosis 32 Differences in Preadipocyte Subpopulations among Fat Depots 32 Differences in Preadipocyte Gene Expression Proﬁles among Depots 33 Conclusions 35 References 36 Plasticity of the Adipose Organ 49 Saverio Cinti Introduction 49 Gross Anatomy Demonstrates that WAT and BAT are Mixed Together in the Adipose Organ 50 Light and Electron Microscopy show that White and Brown Adipocytes have a Well-Deﬁned and Distinct Morphology 51 WAT 51 BAT 52 WAT and BAT have a Different Vascular and Nerve Supply 53 WAT and BAT have a Different Physiology 54 Phenotype of the Adipose Organ is Variable: Plasticity of the Adipose Organ 55 Transformation of the Phenotype: Cold and Warm Exposure and Acclimatization 55 Transformation of the Phenotype: Pregnancy and Lactation 58 Transformation of the Phenotype: Hypertrophy and Hyperplasia (Positive Energy Balance: Overweight and Obesity) 59 Transformation of the Phenotype: Hypoplasia (Negative Energy Balance: Caloric Restriction and Fasting) 60 Adipose Organ of Humans 61 References 63 Biology of Adipose Tissue Stem Cells 69 Jeffrey M. Gimble, Bruce A. Bunnell, and Farshid Guilak Introduction 69 In Situ Localization and Embryology 69 Isolation Methods 70 Yield, Proliferation Rate, Depot, and Aging Inﬂuences Characterization 71 Immunophenotype 71

70

Contents

4.4.2 4.4.3 4.4.4 4.4.5 4.5 4.5.1 4.5.2 4.6

Immunogenicity 71 Proteomic and Transcriptomic Analysis 72 Cytokine Proﬁle 72 Clonality 72 Differentiation and Potential Utility for Regenerative Medicine Mesodermal Lineages 73 Endodermal and Ectodermal Lineages 73 Conclusions 74 References 74

Part Two Metabolic Functions of Adipose Tissue 5 5.1 5.2 5.2.1 5.2.2 5.2.2.1 5.2.2.2 5.2.2.3 5.2.2.4 5.3 5.3.1 5.3.2 5.3.3 5.3.4 5.3.5 5.3.6 5.4

6

6.1 6.2 6.3 6.4 6.5 6.6

81

Molecular Mechanisms of Adipocyte Lipolysis 83 James G. Granneman and Hsiao-Ping H. Moore Introduction 83 Key Players in Adipocyte Lipolysis 83 Lipid Droplets and Droplet Scaffold Proteins 84 Lipases 86 HSL 86 Adipose Triglyceride Lipase 87 CGI-58 87 Other Lipases and the Biological Signiﬁcance of HSL versus ATGL 88 Lipolytic Protein Trafﬁcking 88 PLIN Subcellular Targeting 88 Interactions with CGI-58 88 Interactions with HSL 89 Interactions with ATGL 89 Disruption and Dispersion of Lipid Droplets Following PKA Activation 90 Additional Interactions 90 Working Model and Unresolved Issues 91 References 93 New Developments in the Lipolytic Processing of Triglyceride-Rich Lipoproteins 97 André Bensadoun, Anne P. Beigneux, Loren G. Fong, and Stephen G. Young Introduction 97 LPL 98 Functional Domains of LPL 99 Regulation of LPL Activity by Angiopoietin-Like Proteins Role of GPIHBP1 in the Lipolysis of Triglyceride-Rich Lipoproteins 101 Role of Apo-AV in Lipolysis 103

100

73

VII

VIII

Contents

6.7

Newly Discovered Regulators of LPL Activity and their Physiological Signiﬁcance 104 References 105

7

Intracellular Fatty Acid Transport, Activation, and Trafficking 109 Doug Mashek Introduction 109 Fatty Acid-Binding Protein Family 109 Function and Regulation of FABP-4 110 Function and Regulation of FABP-5 112 Fatty Acid Activation and Channeling: Role of Long-Chain Acyl-CoA Synthetases and Fatty Acid Transport Proteins 113 Role of Acyl-CoA-Binding Protein in Acyl-CoA Metabolism 116 Regulation and Function of Distinct Fatty Acid and Acyl-CoA Pools 117 Contribution of Fatty Acid and Acyl-CoA Metabolism to Metabolic Diseases 119 Conclusions 121 References 121

7.1 7.2 7.2.1 7.2.2 7.3 7.4 7.5 7.6 7.7

8

8.1 8.2 8.3 8.3.1 8.3.2 8.3.3 8.3.4

8.4 8.4.1 8.4.2 8.5 8.6 8.7

Aquaporins and Adipose Tissue: Lesson from Discovery to Physiopathology and to the Clinic of Aquaporin Adipose (AQP7) 129 Ken Kishida Introduction 129 Characteristics of Adipocytes and Gycerol Metabolism in the Mammalian Body 130 Adipose Glycerol Channel: AQP7 132 AQP7: A Putative Adipose-Speciﬁc Glycerol Channel 132 Function and Regulation of AQP7 in Adipocytes 134 Human AQP7 Genetic Mutation 137 Adipose-Derived Glycerol and Gluconeogenesis through AQP7 – Lessons from AQP7-Deﬁcient Mice and Cells 139 Hepatic Glycerol Channel: AQP9 141 AQP9: A Putative Hepatic-Speciﬁc Glycerol Channel 141 Gluconeogenesis through AQP9 – Lessons from AQP9-Deﬁcient Mice 142 Coordination of Adipose Glycerol Channel, AQP7, and Hepatic Glycerol Channel, AQP9 143 Dysregulation of AQP7 and AQP9 in Obesity with Insulin Resistance 143 Conclusions 144 References 144

Contents

9

9.1 9.2 9.2.1 9.2.2 9.3 9.4 9.5 9.6 9.6.1 9.6.2 9.6.3 9.6.4 9.7 9.7.1 9.7.2 9.7.3 9.7.4 9.8

Signaling Pathways Controlling Lipolysis and Lipid Mobilization in Humans 149 Max Lafontan Introduction 149 Role of Lipases in the Regulation of Hydrolysis of Fat Cell Triacylglycerols 151 Hormone-Sensitive Lipase 151 Adipose Tissue Triglyceride Lipase 151 Adrenergic Control of cAMP Production, Lipolysis and Lipid Mobilization 152 Control of cAMP Production by Adenylyl Cyclase Inhibitors – Inhibition of Lipolysis 157 Insulin: A Major Antilipolytic Agent Controlling cAMP Degradation 158 Natriuretic Peptides Control cGMP Production, Lipolysis, and Lipid Mobilization in Humans 159 Natriuretic Peptides 159 Lipolytic Effect of Natriuretic Peptides 160 Induction of Lipid Mobilization by Administration of Pharmacological Doses of ANP 162 Contribution of ANP to the Physiological Control of Lipid Mobilization in Humans 162 Other Lipolytic Pathways 163 Growth Hormone 163 IL-6 164 TNF-a 165 Other Lipolytic Peptides 166 Future Trends and Pharmacological Prospects 167 References 168

Part Three Endocrine Functions of Adipose Tissue 181 10 10.1 10.2 10.2.1 10.2.2 10.2.3 10.3 10.4 10.5 10.6 10.6.1 10.6.2

Leptin Secretion and Action 183 Robert V. Considine Introduction 183 Leptin Synthesis 184 Gender and Body Fat Distribution Determine Serum Leptin 184 Caloric Intake, Insulin, and Glucose Inﬂuence Serum Leptin 185 Transcriptional Regulation of Leptin Synthesis in Adipocytes 186 Leptin Receptors 187 Leptin Action in the Central Nervous System 187 Leptin Resistance in Obesity 189 Metabolic Complications of Hyperleptinemia in Obesity 190 Leptin and Obesity-Related Hypertension 190 Other Possible Pathologic Effects of Leptin 191

IX

X

Contents

10.7 10.7.1 10.7.2 10.7.3 10.7.4 10.8

Leptin Therapy in Humans 192 Leptin, Weight Loss, and Human Obesity Congenital Leptin Deﬁciency 192 Lipodystrophic Leptin Deﬁciency 193 Hypothalamic Amenorrhea 193 Conclusions 194 References 194

11

Adiponectin 201 Jonathan P. Whitehead and Ayanthi A. Richards Introduction 201 Adiponectin Structure and Post-Translational Modiﬁcations 202 Signiﬁcance and Bioactivity of Adiponectin Multimers 204 Adiponectin and Liver 205 Adiponectin and Skeletal Muscle 206 Adiponectin and the Vasculature 206 Adiponectin and the Brain 207 Adiponectin Expression and Secretion 208 Adiponectin Secretion 209 Ectopic Adiponectin Expression 211 Regulation of Expression and Secretion 212 Oxidative stress 212 Activators of PPARc – TZDs and Fish Oils 213 Weight Loss 213 Other Agents 214 Adiponectin Clearance 214 Adiponectin Receptors and Downstream Effectors 215 Adiponectin Signaling 216 Conclusions 217 References 218

11.1 11.2 11.3 11.4 11.5 11.6 11.7 11.8 11.9 11.10 11.11 11.11.1 11.11.2 11.11.3 11.11.4 11.12 11.13 11.14 11.15

12

12.1 12.2 12.3 12.4 12.5 12.6 12.7 12.8

192

Preadipocyte factor-1 and Adipose Tissue-Specific Secretory Factor/Resistin – Two Secreted Factors from Adipose Tissue: Role in Adipogenesis and Insulin Resistance 231 Hei Sook Sul, Yuhui Wang, and Carolyn Hudak Introduction 231 Pref-1 Structure 232 Pref-1 Inhibition of Adipocyte Differentiation 233 Mechanism for Pref-1 Function 234 In Vivo Effect of Pref-1 on Adipogenesis and Glucose/Insulin Homeostasis 235 ADSF/Resistin: Identiﬁcation and Structure 236 ADSF/Resistin Expression and Function 237 Conclusions 239 References 240

Contents

13 13.1 13.2 13.2.1 13.2.2 13.2.3 13.2.4 13.2.5 13.2.6 13.2.7 13.2.8 13.3 13.3.1 13.3.2 13.4

14 14.1 14.2 14.3 14.4 14.5 14.5.1 14.5.2 14.6 14.7 14.8

Adipose Tissue and Blood Pressure Regulation 245 Lisa A. Cassis and Sara B. Police Introduction 245 Adipose Tissue Changes with Obesity: Relation to Blood Pressure Control 245 Adipocyte RAS in Obesity-Related Hypertension 247 Leptin in Obesity-Related Hypertension 249 Adiponectin in Obesity-Related Hypertension 250 Insulin and Obesity-Related Hypertension 251 Plasminogen Activator Inhibitor-1 and Obesity-Related Hypertension 252 Free Fatty Acids and Obesity-Related Hypertension 252 Resistin and Obesity-Related Hypertension 253 11b-HSD-1 and Obesity-Related Hypertension 254 Regional Adipose Deposition and Blood Pressure Regulation 254 Changes in Visceral Adipose Tissue in Obesity-Related Hypertension 255 Potential Role for Perivascular Adipose Tissue in Obesity-Related Hypertension 255 Conclusions 256 References 257 Adipokines, Inflammation, and Obesity 265 Karine Clément Introduction 265 Contribution of Adipose Tissue in Systemic Inﬂammation during Obesity 266 Adipose Tissue Depots and Adipokine Production 268 Adipokines and Adipose Tissue Cell Types 269 Adipokines, Macrophages, and the Biology of Adipocytes 270 Chemoattraction 271 Paracrine Cross-Talk in the Adipose Tissue via Adipokines 272 Adipokines and Complications of Obesity 274 Adipokines and Weight Loss 275 Conclusions 276 References 276

Part Four Adipose Tissue and Disease 15

15.1 15.2

283

Depot-Specific Biology of Adipose Tissues: Links to Fat Distribution and Metabolic Risk 285 Mi-Jeong Lee and Susan K. Fried Introduction 285 Adipose Depots: Deﬁnitions 286

XI

XII

Contents

15.3 15.3.1 15.3.2 15.4 15.4.1 15.4.2 15.4.3 15.5

15.6 15.7 15.8 15.9 15.10

16

16.1 16.2 16.2.1 16.2.2 16.2.3 16.2.4 16.2.4.1 16.2.4.2 16.2.4.3 16.2.4.4 16.2.4.5 16.3 16.4 16.5 16.6 16.7 16.8 16.9

Physiological and Anatomical Differences among Depots may Drive Functional Heterogeneity 286 Depot Differences in Cellular Composition 286 Deﬁnition of Visceral Fat Depots 287 Heterogeneity in Adipocyte Function among Adipose Depots 288 Lipolysis 288 Triglyceride Deposition 289 Glucose Uptake and Insulin Action 290 Regional Differences in Adipose Tissue Gene Expression and Protein Production: Relationship to the Metabolic Syndrome 291 Search for Novel Adipokines with Depot-Speciﬁc Expression that Link Regional Adiposity to Metabolic Risk 291 Importance of Adipose Tissue Macrophages and other Immunocytes in Regional Adipose Tissue Dysfunction 293 Gene Expression Proﬁles are Providing New Insights on Regional Adipose Growth and Function 295 Depot Differences in Cell Proliferation and Differentiation Capacity 296 Conclusions and Future Directions 297 References 298 Viral Induction of Obesity and Adipogenesis 307 Magdalena Pasarica, Rohan N. Dhurandhar, Nazar Mashtalir, and Nikhil V. Dhurandhar Introduction 307 Viruses 313 Canine Distemper Virus 313 Rous-Associated Virus-7 314 Borna Disease Virus 314 Adenoviruses 315 SMAM-1 315 Ad-36 316 Ad-5 319 Ad-37 319 Adipogenic Potential of other Adenoviruses 319 Chlamydia pneumoniae 320 Gut Microbiota 320 Gut Parasites 321 Scrapie Agents 322 Interaction of Pathogens and Adipose Tissue 323 Adipogenic Pathogens and Humans 324 Conclusions 324 References 325

Contents

17 17.1 17.2 17.3 17.4 17.5 17.6 17.7 17.8

18 18.1 18.2 18.3 18.4 18.4.1 18.4.2 18.4.3 18.5 18.5.1 18.5.1.1 18.5.2 18.5.2.1 18.5.2.2 18.5.2.3 18.6 18.6.1 18.6.1.1 18.6.1.2 18.6.2 18.6.2.1 18.7 18.8

Adipose Tissue Cachexia 333 Michael John Tisdale 333 Introduction 333 Changes in Adipose Tissue in Cachexia 333 Energy Expenditure in Cancer Patients 335 Factors Governing Adipose Tissue Mass 336 Mechanism of Loss of Adipose Tissue in Cachexia 337 Requirements of Tumor-Bearing Animals for Lipids 338 Fat-Mobilizing Substances in Cancer Cachexia 338 Conclusion 340 References 342 Obesity and Diabetes: Lipotoxicity 347 Christopher J. Lelliott, Matej Ore4si4c, and Antonio J. Vidal-Puig Introduction 347 White Adipose Tissue at the Center of Lipid Homeostasis and Delivery 348 Insulin Resistance in Adipocytes Disrupts the Balance between Lipid Storage and Secretion 348 Scenarios that may Result in Ectopic Fat Deposition 348 Altered Plasticity of the Adipose Tissue: A Shift in Expansion Towards Hypertrophy 349 Impaired Fat Deposition Capacity in Adipose Tissue 350 Inappropriate Balance of Substrate Uptake and Oxidative Capacity in Peripheral Tissue 353 Mechanisms Contributing to the Lipotoxicity in the Peripheral Organs 355 Lipotoxicity in Skeletal Muscle 355 Randle Hypothesis and its Successors 355 Molecular Mechanisms for the Generation of Muscle Lipotoxicity 356 Type of Lipids is More Important than the Amount of Fat Deposited 356 Diacylglycerols and Insulin Resistance 358 Ceramides and Insulin Resistance 358 Impaired Oxidation as a Trigger for Lipotoxicity 360 Adipocytokines Proinﬂammatory Activity Contributes to Lipotoxicty in Skeletal Muscle 361 Adipose Tissue Macrophages as Key Players for Lipotoxicity 361 Signaling Effector Pathways in Lipotoxicty 361 Lipotoxicity and Insulin Resistance Affecting Liver Metabolism 362 Hepatic Lipotoxicity 362 Pancreatic b-Cell as a Target for Lipotoxicity 363 New Analytical and Computational Methods to Identify Lipotoxicity-Related Metabolic Networks 363

XIII

XIV

Contents

18.9

Lessons from Lipotoxicity – Potential Antilipotoxic Therapeutic Strategies 365 References 365

19

Obesity and Cancer 369 Andrew G. Renehan Introduction 369 Epidemiology 369 Excess Body Weight and Cancer Risk 369 Excess Body Weight and Cancer Mortality 370 Biological Mechanisms 371 Candidate Mechanisms 371 Insulin and IGFs 372 Insulin-Cancer Hypothesis 372 Insulin and C-Peptide 372 IGFs 376 Sex Steroids 376 Estrogen and Breast Cancer 376 Androgens and Breast Cancer 377 Sex Steroids and Endometrial Cancer 377 Adipokines 378 Overview 378 Leptin and Cancer in Humans 378 Adiponectin and Cancer Risk in Humans 379 Adipokines, Animal Models, and Cancer Risk 380 Other Biological Candidates 380 Obesity-Related Inﬂammatory Markers 380 Nuclear Factor-kB System 381 Oxidative Stresses 381 Mechanical Mechanisms 381 New Research Areas 382 References 382

19.1 19.2 19.2.1 19.2.2 19.3 19.3.1 19.3.2 19.3.2.1 19.3.2.2 19.3.2.3 19.3.3 19.3.3.1 19.3.3.2 19.3.3.3 19.3.4 19.3.4.1 19.3.4.2 19.3.4.3 19.3.4.4 19.4 19.4.1 19.4.2 19.4.3 19.5 19.6

20 20.1 20.2 20.2.1 20.2.1.1 20.2.1.2 20.2.2 20.2.2.1 20.2.2.2 20.3

Overview of Acquired and Genetic Lipodystrophies 385 Tisha Joy and Robert A. Hegele Introduction 385 Congenital Lipodystrophies 386 Congenital Generalized Lipodystrophy (Berardinelli–Seip Syndrome) 386 Clinical Features 386 Molecular Genetics 390 Familial Partial Lipodystrophy 390 Clinical Features 390 Molecular Genetics 391 Acquired Lipodystrophies with a Possible Genetic Component 393

Contents

20.3.1 20.3.1.1 20.3.1.2 20.3.2 20.3.2.1 20.3.2.2 20.3.3 20.4 20.4.1 20.4.1.1 20.4.1.2 20.4.2 20.4.2.1 20.4.2.2 20.4.3 20.4.3.1 20.4.3.2 20.4.4 20.4.4.1 20.4.4.2 20.4.5 20.4.5.1 20.4.5.2 20.5

Acquired Generalized Lipodystrophy 393 Clinical Features 393 Molecular Genetics 393 Acquired Partial Lipodystrophy (Barraquer–Simons Syndrome) 393 Clinical Features 393 Molecular Genetics 394 HIV-Related Lipodystrophy 394 Lipodystrophy Associated with other Syndromes 395 Mandibuloacral Dysplasia 395 Clinical Features 395 Molecular Genetics 395 SHORT Syndrome 396 Clinical Features 396 Molecular Genetics 396 Neonatal Progeroid Syndrome 396 Clinical Features 396 Molecular Genetics 396 Hutchinson–Gilford Progeria Syndrome 396 Clinical Features 396 Molecular Genetics 397 Werner Syndrome 397 Clinical Features 397 Molecular Genetics 397 Conclusions 397 References 398

21

Mouse Models of Lipodystrophy 403 Jimmy Donkor and Karen Reue Introduction 403 Physiological Mechanisms of Lipodystrophy in Mouse Models 403 Lipodystrophic Models with Impaired Adipogenesis 407 A-ZIP/F1 Transgenic Mouse 408 aP2- SREBP-1c Transgenic Mouse 408 Mouse Models with Altered PPAR-c Levels 409 C/EBPa-Deﬁcient Mouse 410 Zmpste24-Deﬁcient Mice 410 Lipodystrophic Models with Impaired Triacylglycerol Biosynthesis 411 GPAT1-Deﬁcient Mouse 411 AGPAT6-Deﬁcient Mouse 412 DGAT1-Deﬁcient Mouse 412 Lipin-1-Deﬁcient Mouse 412 Lipodystrophic Models with Enhanced Energy Expenditure 414 Leptin Transgenic Mouse 414 PPAR-d Transgenic Mouse 415

21.1 21.2 21.3 21.3.1 21.3.2 21.3.3 21.3.4 21.3.5 21.4 21.4.1 21.4.2 21.4.3 21.4.4 21.5 21.5.1 21.5.2

XV

XVI

Contents

21.5.3 21.6 21.6.1 21.6.2 21.6.3 21.6.4 21.6.5 21.7

22 22.1 22.2 22.3 22.4 22.5 22.6 22.7 22.8 22.8.1 22.8.2 22.8.3 22.8.4 22.9 22.9.1 22.9.2 22.10 22.11

23

23.1 23.2 23.2.1 23.2.2 23.3 23.3.1 23.3.2 23.4 23.5

FOXC2 Transgenic Mouse 415 Mouse Models with Acquired or Conditional Lipodystrophy 416 aP2-DTA Transgenic Mouse 416 FAT-ATTAC Transgenic Mouse 416 RSK2 Deﬁcient Mouse 417 Drug-Induced Lipoatrophy 417 Diet-Induced Lipoatrophy 417 Conclusions 418 References 419 Caloric Restriction, Longevity, and Adiposity 423 Leanne M. Redman and Eric Ravussin Introduction 423 Physiological Changes with Aging 424 Aging and Caloric Restriction 424 Energy Restriction may Alter the ‘‘Rate of Living’’ 425 CR and Oxidative Stress 426 CR and Cardiovascular Disease 427 CR and Insulin Resistance/Type 2 Diabetes Mellitus 427 What is Known from Humans? 427 Centenarians from Okinawa 427 Vallejo Study 428 Unexpected CR in Biosphere 2 428 Randomized Controlled Trials of Prolonged CR in Humans 429 Could CR Increase Longevity in Humans? 431 How Much CR? 432 How Long is CR Required? 432 CR Mimetics 433 Conclusions 434 References 434 Peroxisome Proliferator-Activated Receptor-c: A Key Regulator of Adipose Tissue Formation, Remodeling, and Metabolism 441 Olga Astapova and Todd Leff Introduction 441 Molecular Biology of PPAR-c 442 PPAR-c Structure and DNA Binding 442 Transcriptional Regulation by PPAR-c 444 PPAR-c is a Master Regulator of Adipose Tissue Development 447 Role of PPAR-c in Adipogenesis – Cell Culture Studies 447 PPAR-c is Required for Adipose Tissue Development In Vivo 448 Metabolic Functions of PPAR-c 449 White versus Brown Fat-Speciﬁc Functions of PPAR-c 450

Contents

23.6 23.7

24 24.1 24.2 24.3 24.3.1 24.3.2 24.4 24.4.1 24.4.2 24.4.3 24.5 24.5.1 24.5.2 24.5.3 24.6 24.6.1 24.6.2

25 25.1 25.2 25.3 25.3.1 25.3.2 25.3.3 25.4 25.5 25.6 25.7

PPAR-c Function in Adipose Tissue Maintenance and Remodeling 452 Conclusions 454 References 454 Early-Life Programming of Adipogenesis and Adiposity 459 Roselle L. Cripps and Susan E. Ozanne Introduction 459 Theories for the Developmental Origins of Obesity 460 Evidence for the Developmental Origins of Obesity 460 Data from Humans 461 Data from Animal Models 461 Adipogenesis 462 Adipogenesis In Vitro 462 Control of Adipogenesis 463 Developmental Alterations to Adipogenesis 464 Potential Mechanisms? 465 Glucocorticoids 465 Leptin 465 Epigenetic Alterations 467 Future Perspectives 467 Optimizing Early Life Nutrition? 467 Interventions? 467 References 468 Evolutionary Aspects of Obesity and Adipose Tissue Function 473 Jonathan C. K. Wells Introduction 473 Thrifty Genotype and Phenotype Hypotheses 474 Ethological Approach 476 Ontogeny 477 Fitness Value 478 Evolutionary History 480 Signiﬁcance of Agriculture 482 Signiﬁcance of Colonizing 483 Signiﬁcance of Social Inequality 485 New Obesogenic Environment 486 References 487 Index

491

XVII

XIX

Preface We are in the midst of a world-wide epidemic of obesity. The causes of this epidemic are complex and multifaceted, and reﬂect an unintended synergy between our evolutionary legacy and the current postindustrial environment we have collectively created. Despite these complexities, one thing is clear: the wide-spread prevalence of obesity has contributed directly and fundamentally to several chronic diseases, foremost of which is diabetes. The nature of the connection between adiposity and disease is incompletely understood. However, experience with human and animal lipodystrophies (abnormalities of adipose tissue quantity and distribution), indicates that the relationship between obesity and diabetes is not due to the magnitude of fat accumulation per se, but rather the functioning of the adipose organ. Thus, the aim of this collection is to outline our current understanding of the basic metabolic, endocrine and immune functions of adipose tissues, and how dysregulation of adipose tissue physiology contributes to disease. The collection is organized along four interrelated topics. Part 1 addresses aspects of the molecular and cellular development of adipocytes, and morphology of adipose tissue. A theme developed in these chapters concerns the remarkable heterogeneity and plasticity of the adipose tissue organ. Indeed, growing evidence demonstrates that various fat depots have distinct developmental origins, cellular composition, and unique endocrine and metabolic proﬁles that differentially contribute to health and disease. Part 2 focuses on metabolic functions that are classically associated with adipose tissue, such as lipid storage and mobilization. Here, too, recent work has identiﬁed novel and unexpected complexities in the transmembrane and intracellular mechanisms by which the balance of lipid storage and mobilization is maintained. Adipose tissue secretes a diverse set of hormones and cytokines, collectively known as adipokines, that greatly impact both local and systemic metabolism. Part 3 addresses recent developments in our understanding of the now ‘‘classic’’ endocrine functions of adipocytes (i.e., leptin and adiponectin), and touches upon the potentially important relationships between the production of inﬂammatory cytokines in adipose tissue during hypertrophic obesity and systemic insulin sensitivity.

XX

Preface

Lastly, nearly half of the collection is devoted to the topic of adipose tissue in health and disease. Chapters in Part 4 explore the developmental, evolutionary and social origins of human obesity. In addition, these chapters address the physiological consequences of exceeding adipose tissue capacity, as occurs in both obese and lipodystrophic states, as well as the speciﬁc risks to health conferred by obesity and potential mechanisms involved. Finally, this section touches upon adipose tissue as a therapeutic target for approaches to promote health and longevity. Detroit, January 2010

Todd Leff and James G. Granneman

XXI

List of Contributors Olga Astapova Wayne State University School of Medicine Department of Pathology 540 E. Canﬁeld Detroit, MI 48201 USA Anne P. Beigneux University of California David Geffen School of Medicine Department of Medicine Division of Cardiology 650 Charles E. Young Drive South A2-237 CHS Bldg. Los Angeles, CA 90095 USA André Bensadoun Cornell University Division of Nutritional Sciences 321 Savage Hall Ithaca, NY 14853 USA Bruce A. Bunnell Tulane Center for Gene Therapy Tulane University Health Sciences Center J. Bennett Johnston Building 1324 Tulane Avenue, SL-99 New Orleans, LA 70112-2699 USA

Lisa A. Cassis University of Kentucky Graduate Center for Nutritional Sciences Lexington, KY 40536 USA Saverio Cinti University of Ancona Department of Molecular Pathology and Innovative Therapies-Anatomy via Tronto 10/a 60020 Ancona Italy Karine Clément Hôpital Pitié-Salpêtrière INSERM, Cordelier Research Center Endocrinology and Nutrition department Batiment Husson Mourrier Boulevard de lhôpital 75013 Paris France Robert V. Considine Indiana University School of Medicine Division of Endocrinology 541 N. Clinical Drive Indianapolis, IN 46202 USA

XXII

List of Contributors

Roselle L. Cripps University of Cambridge Metabolic Research Laboratories Level 4, Institute of Metabolic Science Box 289, Addenbrookes Hospital Cambridge CB2 OQQ UK Nikhil V. Dhurandhar Louisiana State University System Pennington Biomedical Research Center Department of Infection and Obesity 6400 Perkins Road Baton Rouge, LA 70808 USA Rohan N. Dhurandhar Louisiana State University System Pennington Biomedical Research Center Department of Infection and Obesity 6400 Perkins Road Baton Rouge, LA 70808 USA Jimmy Donkor University of California David Geffen School of Medicine Departments of Human Genetics and Medicine 695 Charles E. Young Drive South Los Angeles, CA 90095 USA Loren G. Fong University of California David Geffen School of Medicine Department of Medicine Division of Cardiology 695 Charles E. Young Drive South Gonda Bldg., Rm4524 Los Angeles, CA 90095 USA

Susan K. Fried University of Maryland School of Medicine Department of Medicine Division of Endocrinology, Diabetes and Nutrition 660 W. Redwood Street Baltimore, MD 21201 USA and Baltimore Veterans Affairs Medical Center Geriatric Research Education and Clinical Center 10 N. Greene Street Baltimore, MD 21201 USA Jeffrey M. Gimble Louisiana State University System Pennington Biomedical Research Center 6400 Perkins Road Baton Rouge, LA 70808 USA James G. Granneman Wayne State University School of Medicine Center for Integrative Metabolic and Endocrine Research Detroit, MI 48201 USA Farshid Guilak Duke University Medical Center Department of Surgery Division of Orthopedics MSRB Rm 375, Box 3093 Durham, NC 27710 USA

List of Contributors

Robert A. Hegele University of Western Ontario Schulich School of Medicine and Dentistry 100 Perth Drive London, Ontario N6A 5K8 Canada

Max Lafontan Institut de Médecine Moléculaire de Rangueil INSERM Unit 858 BP 84225 31432 Toulouse Cedex 4 France

Carolyn Hudak University of California Department of Nutritional Science and Toxicology Morgan Hall Berkeley, CA 94720 USA

Mi-Jeong Lee University of Maryland School of Medicine Department of Medicine Division of Endocrinology, Diabetes and Nutrition 660 W. Redwood Street Baltimore, MD 21201 USA

Tisha Joy University of Western Ontario Division of Endocrinology 100 Perth Drive London, Ontario N6A 5K8 Canada James L. Kirkland Mayo Clinic Robert and Arlene Kogod Center on Aging 200 First St., S.W. Rochester, MN 55905 USA Ken Kishida Osaka University Graduate School of Medicine Department of Metabolic Medicine 2-2-B5 Yamada-oka, Suita Osaka 565-0871 Japan

Todd Leff Wayne State University School of Medicine Department of Pathology Detroit, MI 00000 USA Christopher J. Lelliott AstraZeneca R&D Cardiovascular/Gastrointestinal Research Area Department of Biosciences Pepparedsleden 1 43183 Mölndal Sweden Doug Mashek University of Minnesota Department of Food Science and Nutrition 1334 Eckles Avenue St. Paul, MN 55108 USA

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List of Contributors

Nazar Mashtalir Louisiana State University System Pennington Biomedical Research Center Department of Infection and Obesity 6400 Perkins Road Baton Rouge, LA 70808 USA

Eric Ravussin Louisiana State University System Pennington Biomedical Research Center Department of Human Physiology 6400 Perkins Road Baton Rouge, LA 70808 USA

Hsiao-Ping H. Moore Lawrence Technological University College of Arts and Sciences Southﬁeld, MI 48075 USA

Leanne M. Redman Louisiana State University System Pennington Biomedical Research Center Department of Human Physiology 6400 Perkins Road Baton Rouge, LA 70808 USA

Matej Ore4si4c VTT Technical Research Center of Finland Tietotie 2 02044 Espoo Finland Susan E. Ozanne University of Cambridge Metabolic Research Laboratories Level 4, Institute of Metabolic Science Box 289, Addenbrookes Hospital Cambridge CB2 OQQ UK Magdalena Pasarica Louisiana State University System Pennington Biomedical Research Center Department of Infection and Obesity 6400 Perkins Road Baton Rouge, LA 70808 USA Sara B. Police University of Kentucky Graduate Center for Nutritional Sciences Lexington, KY 40536 USA

Andrew G. Renehan University of Manchester School of Cancer, Enabling Sciences and Technology Manchester Academic Health Science Center The Christie NHS Foundation Trust Wilmslow Road Manchester M20 4BX UK Karen Reue University of California David Geffen School of Medicine Departments of Human Genetics and Medicine 695 Charles E. Young Drive South Los Angeles, CA 90095 USA Ayanthi A. Richards University of Queensland Princess Alexandra Hospital Diamantina Institute for Immunology, Cancer and Metabolic Medicine Brisbane Queensland, QLD 4102 Australia

List of Contributors

Jacqueline M. Stephens Louisiana State University Department of Biological Sciences 202 Life Sciences Building Baton Rouge, LA 70803 USA Hei Sook Sul University of California Department of Nutritional Science and Toxicology Morgan Hall Berkeley, CA 94720 USA Tamara Tchkonia Mayo Clinic Robert and Arlene Kogod Center on Aging 200 First St., S.W. Rochester, MN 55905 USA Thomas Thomou Mayo Clinic Robert and Arlene Kogod Center on Aging 200 First St., S.W. Rochester, MN 55905 USA Michael John Tisdale Aston University School of Life and Health Sciences Nutritional Biomedicine Birmingham B4 7ET UK Antonio J. Vidal-Puig University of Cambridge Addenbrookes Hospital Department of Clinical Biochemistry Hills Road Cambridge CB2 2QR UK

Yuhui Wang University of California Department of Nutritional Science and Toxicology Morgan Hall Berkeley, CA 94720 USA Jonathan C.K. Wells UCL Institute of Child Health Childhood Nutrition Research Centre 30 Guilford Street London WC1N 1EH UK Ursula A. White Louisiana State University Department of Biological Sciences 202 Life Sciences Building Baton Rouge, LA 70803 USA Jonathan P. Whitehead University of Queensland Princess Alexandra Hospital Diamantina Institute for Immunology, Cancer and Metabolic Medicine Brisbane Queensland, QLD 4102 Australia Stephen G. Young University of California David Geffen School of Medicine Department of Medicine Division of Cardiology 650 Charles E. Young Drive South A2-237 CHS Bldg. Los Angeles, CA 90095 USA

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Part One Adipose Tissue Development and Morphology

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1 Transcriptional Control of Adipogenesis and Fat Cell Gene Expression Ursula A. White and Jacqueline M. Stephens 1.1 Introduction

Adipocytes are highly specialized cells that play a major role in energy homeostasis in vertebrate organisms. Excess adipocyte size or number is a hallmark of obesity, which is currently a global epidemic. Obesity is not only the primary disease of fat cells, but a major risk factor for the development of non-insulin-dependent diabetes mellitus, cardiovascular disease, and hypertension. Obesity and its related disorders result in dysregulation of the mechanisms that control the expression of metabolic genes in adipocytes. Therefore, understanding adipocyte differentiation is relevant not only for understanding the pathogenesis of metabolic diseases, but also for identifying proteins or pathways that might be appropriate targets for pharmacological interventions. In the last 15 years, signiﬁcant advances towards an understanding of the regulatory processes involved in adipocyte differentiation have largely been made by the identiﬁcation of transcription factors that regulate the differentiation of fat cells and/or are involved in the induction and maintenance of adipocyte gene expression. Interestingly, the majority of studies that have identiﬁed transcriptional regulators of adipogenesis have been performed in vitro. These studies have been primarily conducted in the 3T3-L1 or 3T3-F442A preadipocyte cell lines that were originally generated in the laboratory of Howard Green at Harvard University [1, 2]. In the last 32 years, these cells lines have been used by thousands of investigators worldwide. In vivo, adipocytes have three primary characteristics, which include lipid storage, insulin sensitivity, and endocrine properties. The 3T3-L1 cells have all three of these notable characteristics of fat cells. In addition, many adipocyte speciﬁc genes have been identiﬁed using this cell line. Many cell types cannot be adequately studied in vitro because the cultured cells do not have all the properties that the cells have in vivo. However, the preadipocyte cell lines that were developed by Green have been an extremely useful model system for adipocyte biologists and the data obtained in these cells has been validated from less mechanistic in vivo studies in the last decade.

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In the late 1980s, a commentary by Steven McKnight and Dan Lane indicated that CAAAT/enhancer-binding protein (C/EBP)-a was a key metabolic regulator of energy metabolism [3]. Numerous studies have since conﬁrmed the role of C/EBP-a and other C/EBP family members in energy balance, and deﬁned roles for these transcription factors in adipocyte differentiation (reviewed in [4]). In 1994, two prominent laboratories independently identiﬁed peroxisome proliferator-activated receptor-c (PPAR)-c as an important modulator of adipocyte differentiation. Studies by Mitch Lazars group observed the induction of PPAR-c during adipogenesis [5], and experiments by Bruce Speigelmans laboratory revealed that PPAR-c was a transcriptional factor that bound to an enhancer element in the fatty acid-binding protein, aP2, promoter and conferred its fat-speciﬁc expression [6]. The early 1990s was the time that Brown and Goldstein identiﬁed sterol regulatory element-binding protein (SREBP)-1 [7], also termed adipocyte differentiation and determination factor (ADD)-1, whose expression was observed to play a role in adipocyte determination by the Spiegelman laboratory [8]. Since this time, several other transcription factors have been found to play an important role in adipocyte differentiation. Section 1.2 focuses on the transcription factors mentioned above and other important transcription factors that are regulated during adipocyte differentiation and are known to contribute to this process. Section 1.3 focuses on target genes that have been shown to be modulated by several of these transcription factors in both differentiating and mature adipocytes.

1.2 Transcriptional Control of Adipogenesis

Studies of the aP2 gene or use of its regulatory sequences have led to signiﬁcant discoveries in adipocyte biology and metabolic diseases. aP2 is an abundantly expressed adipocyte gene that was ﬁrst discovered in 1984 [9]. One of the earliest studies on PPAR-c identiﬁed it as a transcription factor that bound to an enhancer element in the aP2 promoter and conferred its fat-speciﬁc expression [8]. Since its discovery, the aP2 promoter has been used by hundreds of laboratories to construct transgenes to have fat-speciﬁc expression. One lesser known fact is that c-Fos was also shown to bind to the aP2 promoter just 124 bp upstream of the transcriptional start site [10]. As a means of introduction, this was the ﬁrst study to propose that activating protein (AP)-1 proteins were regulators of adipocyte gene expression. 1.2.1 AP-1 Transcription Factors

Members of the AP-1 family of transcription factors are well-known regulators of cellular proliferation and differentiation. AP-1 is a collective term referring to dimeric transcription factors composed of c-Jun, JunB, JunD and c-Fos, FosB, Fra-1, or Fra-2 subunits that bind to a common DNA site – the AP-1-binding site (reviewed in [11]). As indicated above, studies by the Spiegelman laboratory indicated that c-Fos was

1.2 Transcriptional Control of Adipogenesis

involved in the modulation of aP2 expression [10]. Several years later, it was shown that the expression of c-Jun, c-Fos, Jun-B, Fos-B, and Fra-1 was induced immediately after the induction of adipocyte differentiation [12, 13]. Although the role of individual AP-1 family members in adipogenesis has not been elucidated, there is strong evidence to indicate these transcription factors are important in vivo. Transgenic mice were generated that express a dominant-negative protein that prevents the DNA binding of B-ZIP transcription factors of both the C/EBP and Jun families under the control of the adipose-speciﬁc aP2 enhancer/promoter. These mice have no white fat tissue throughout life [14]. Collectively, these studies suggest that the induction and expression of AP-1 transcription factors are important in fat cell differentiation. 1.2.2 Signal Transducers and Activators of Transcription

In the last 10 years, several groups have studied the modulation and function of signal transducers and activators of transcription (STAT) proteins during adipogenesis and in mature fat cells. The STAT family of mammalian transcription factors is comprised of seven proteins (STAT-1, -2, -3, -4, -5A, -5B, and -6) that, in response to stimulation of various receptors, mainly those for cytokines, are phosphorylated on tyrosine residues causing their translocation to the nucleus. Each STAT family member shows a distinct pattern of activation by cytokines and upon nuclear translocation can regulate the transcription of particular genes [15]. STATs have been shown to bind distinct DNA sequences and this binding regulates the transcription of speciﬁc genes [15, 16]. Since the tissue distribution and function of each STAT is unique, the regulation of tissue-speciﬁc genes appears to be a physiological role for these proteins [17]. This hypothesis is supported by numerous reports that demonstrate that speciﬁc STATs are activated differently by growth factors and cytokines, and STAT activation can be cell-type dependent. In addition, transgenic knockout experiments have revealed crucial roles for each known mammalian STAT [15] and cell-speciﬁc functions for STAT family members have been identiﬁed [18]. The ﬁrst studies on STATexpression in 3T3-L1 cells revealed that STAT-1, -5A, and 5B were highly induced during murine adipogenesis [19]. Similar results were observed during the in vitro differentiation of human preadipocytes [20]. In addition, the ectopic expression of C/EBP-b and -d in nonprecursor cells results in an induction of adipogenesis [21] that is accompanied by an induction in STAT-5A and -5B protein levels [22]. These two STATproteins are also coordinately regulated with both PPAR-c and C/EBP-a in differentiating 3T3-L1 cells under a variety of different conditions [23]. In 3T3-F442A preadipocytes, the ability of growth hormone to modulate adipogenesis was attenuated by STAT-5 antisense oligonucleotides [24]. Also, constitutively active STAT-5 is capable of replacing the requirement for growth hormone in adipogenesis of these cells [25]. Moreover, ectopic expression of STAT-5A has been shown to confer adipogenesis in 3T3-L1 preadipocytes [26] and in two different nonprecursor cell lines [27]. Interestingly, STAT-5B was not capable of conferring adipogenesis in nonprecursor cells [27]. Transgenic deletion of STAT-5A, STAT-5B, or

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AP 1 (fos, jun, etc.) STAT 5 phosphorylaon C/EBP and C/EBP KLF15 SREBP 1/ADD1 PPAR C/EBP STAT5A and STAT 5B

Figure 1.1 Induction of transcription factors during adipogenesis.

both STAT-5 genes in mice resulted in signiﬁcantly reduced fat pad sizes compared to wild-type mice [28]. Also, in primary cultures of adipose tissue from these animals, growth hormone did not stimulate lipolysis as it did in adipocytes from wild-type animals [29], suggesting that some of the effects of growth hormone on fat metabolism are dependent on STAT-5 proteins. It should be noted that the increased expression of STAT-5 proteins is not typically observed until after the induction of C/ EBP-a and PPAR-c (refer to Figure 1.1), yet the activation of STAT-5 proteins in preadipocytes occurs prior to the induction in expression of PPAR-c in 3T3-L1 cells [27]. In fact, both STAT-5 proteins are tyrosine phosphorylated and translocate to the nucleus within 15 min after the induction of adipogenesis of preadipocytes [27, 30]. Coupled with the observations in STAT-5 null mice, which have fat pads one-ﬁfth normal size [28], the data suggest that activation of STAT-5 proteins may be an important driver of adipogenesis both in vitro and in vivo. This hypothesis is also supported by work indicating that one of the PPAR-c promoters can be modulated by STAT-5 [31], suggesting that STAT-5 activation might drive adipogenesis by inducing PPAR-c expression. In summary, work by a variety of laboratories has demonstrated that STAT-5 proteins, particularly STAT-5A, are activated and induced during adipogenesis, and play an important role in adipose tissue development. 1.2.3 Kr€ uppel-Like Factors

Kr€ uppel-like zinc ﬁnger transcription factors (KLFs) are known to play diverse roles in cell differentiation and development in mammals. One protein in the KLF family, KLF-15, was shown to be highly induced during the differentiation of 3T3-L1 preadipocytes into adipocytes [32]. Inhibition of KLF-15 function or expression with a dominant-negative mutant or via RNA interference results in an inhibition of adipogenesis in 3T3-L1 cells [32]. These studies also revealed that KLF-15 could confer adipogenesis in nonprecursor cells and result in the induction of PPAR-c expression. Similar to KLF-15, KLF-5 expression is also highly induced during adipocyte differentiation in 3T3-L1 cells and embryonic ﬁbroblasts obtained from heterozygote KLF-5 mice exhibit reduced adipogenesis [33]. Another member of this family, KLF-2, has been shown to be a negative regulator of adipogenesis and has the ability to attenuate PPAR-c expression [34]. KLF-s are not only involved in lipid accumulation, but also appear to play a role in the ability of the adipocyte to

1.2 Transcriptional Control of Adipogenesis

be insulin-sensitive, as indicated by studies showing that KLF-15 is important in the expression of glucose transporter 4 (GLUT4) [35]. 1.2.4 SREBPs

In 1993, studies by the Spiegelman group identiﬁed a basic helix–loop–helix transcription factor that was expressed in adipocytes and whose expression was increased during adipogenesis [8]. The protein was called ADD-1. Two months later, this protein was labeled SREBP-1 by the Brown and Goldstein laboratories, who named the transcription factor for its ability to bind sterol-responsive elements within the promoter of the low-density lipoprotein receptor gene [7]. It is now known that there are three SREBP isoforms (SREBP-1a, -1c, and -2) that have been well characterized. SREBP-1a and -1c are transcribed from the same gene, each by a distinct promoter, and the predominant SREBP-1 isoform in liver and adipose tissue is SREBP-1c. SREBP-2 is relatively selective in transcriptionally activating cholesterol biosynthetic genes and SREBP-1c has a greater role in regulating genes associated with fatty acid synthesis (reviewed in [36]). Although there is clear evidence that SREBP is an insulin-modulated transcription factor involved in the regulation of genes associated with cholesterol and lipid metabolism, there are less convincing studies to indicate that SREBPs are critical for adipogenesis. Mice deﬁcient in SREBP-1 do not have a signiﬁcantly decreased amount of white adipose tissue, but SREBP-2 levels were increased suggesting it might compensate for SREBP-1 in this animal model [37]. These in vivo studies are supported by additional transgenic studies where SREBP-1 deﬁcient mice were crossed with ob/ob (leptin-deﬁcient) mice and it was found that SREBP-1 was not required for the development of obesity [38]. These observations concluded that SREBP-1 regulation of lipogenesis was highly involved in the development of fatty livers, but was not a determinant of obesity in this animal model [38]. However, ectopic expression of a dominant-negative SREBP-1c was shown to attenuate adipocyte differentiation [39]. In addition, overexpression of SREBP-1c enhanced the adipogenic activity of PPAR-c [39] and other studies suggest that SREBP-1c contributes to the generation of PPAR-c ligands [40]. In summary, in vitro studies support a role for SREBP-1 in adipogenesis, whereas in vivo studies indicate that SREBPs are not required for the production or expansion of adipose tissue. 1.2.5 C/EBP

C/EBP transcription factors were the ﬁrst family of transcription factors shown to play a critical role in the differentiation of fat cells in vitro. Today, we know that transgenic mice lacking both C/EBP-b and C/EBP-d or C/EBP-a alone have defective adipocyte differentiation [41, 42]. Prior to these in vivo observations, the cascade of induction of these three C/EBP family members was revealed by McKnight and collaborators who showed that C/EBP-b and -d were induced immediately after the

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induction of differentiation, whereas C/EBP-a expression did not occur until 4–5 days after the initiation of differentiation [43]. This group also demonstrated that C/ EBP-b and -d were responsible for inducing C/EBPa expression [44]. Ectopic expression studies conducted by several laboratories demonstrated the adipogenic capabilities of C/EBP-a or -b alone, or in the presence of C/EBP-d [21, 44–46]. Today, C/EBP-a and PPAR-c are considered the two primary transcription factors that mediate adipogenesis. However, cells lacking C/EBP-a are capable of adipogenesis, but are not insulin-sensitive [47, 48]. There is also evidence to indicate that C/EBPs may play a role in the induction of PPAR-c ligands [49]. In summary, both in vitro and in vivo studies indicate a substantial role for C/EBP-b, -d, and -a in adipogenesis. Although C/EBP-a may not be required for lipid accumulation, this transcription factor clearly plays a role in conferring insulin sensitivity in adipocytes. 1.2.6 PPAR-c

Although a number of transcription factors, including those mentioned above, have been shown to have profound effects on fat cell differentiation and the expression of adipocyte genes, only one adipocyte transcription factor has been shown to be necessary foradipogenesis.PPAR-c isa memberofthenuclearhormonereceptorsuperfamilythat is required for the development of adipocytes, and deletion of PPAR-c in mice results in placentaldysfunctionandembryonic lethality [50,51]. Asmentioned above,PPAR-c was originally identiﬁed as a transcription factor induced during differentiation that bound an enhancer element within the aP2 promoter [6]. Since that time, there have been multiple studies on the role of PPAR-c in adipocytes. A remarkable ﬁnding in 1995 was that the insulin-sensitizing drugs thiazolidinediones were ligands for PPAR-c [52]. These were some of the ﬁrst molecular studies to indicate the importance of adipose tissue in insulin resistance. Many investigators were surprised to learn that activating a transcription factor whose expression was highly enriched in fat cells could contribute to whole-animal insulin sensitivity. It is now known that adipocytes secrete several hormones that can affect the activity of other tissues; however, the studies of PPAR-c have revealed that modulation of this transcription factor can contribute to systemic insulin resistance. Although the PPAR-c null mice are embryonic lethal [51], transgenic mice lacking PPAR-c speciﬁcally in adipose tissue exhibit greatly reduced sized fat pads, and insulin resistance in fat and liver [53]. However, PPAR-c heterozygote mice have enhanced insulin sensitivity [54]. Together, these studies suggest the amount of PPAR-c in adipose tissue is physiologically relevant. In the last several years, several studies have examinedpathways that are involved inregulating the levels of PPAR-c. In particular, the ubiquitin–proteasome system has emerged as an important regulator of PPAR-c proteins [55, 56]. In addition, a role for the ubiquitin-like protein SUMO (small ubiquitin-like modiﬁer) in regulating PPAR-c has been demonstrated by several groups [57–60]. The phosphorylation of PPAR-c by mitogen-activated protein kinases is also an important modulator of the activity of this transcription factor (reviewed in [61]). A very recent study conﬁrmed that PPAR-c knockdown prevented adipocyte differentiation, but also suggested that PPAR-c was not required for maintenance of the

1.3 Identification of Adipocyte Transcription Factor Target Genes

differentiated state after the cells had undergone adipogenesis [62]. These observations are supported many anecdotal observations that indicate that PPAR-c is decreased as adipocytesageinvitro.Inaddition,theincreaseoflifespanviacaloricrestrictionresultsin the induction of sirtuin (SIRT)-1 – a transcriptional modulator with deacetylase activity that represses PPAR-c activity in vivo [63]. This study also demonstrated that the repression of PPAR- c by SIRT1 was evident in 3T3-L1 adipocytes. Collectively, studies bynumerous laboratories demonstratethe adipogenic capabilities ofPPAR-c. However, theroleofthistranscriptionfactorinmatureadipocytesisnotwellunderstoodandrecent studies suggest that this transcription factor is not required for maintenance of the adipocyte following adipogenesis [62]. Nonetheless, PPAR-c expression and activity is controlled at multiple levels, including alternative promoter usage, tissue-limited expression, phosphorylation, acetylation, ubiquitylation, and SUMOylation. The multiple levels of regulation of this transcription factor suggest that controlling the amount and activity of PPAR-c is important. The role of PPAR-c in the development and treatment of diabetes is well established (reviewed in [64]), and the importance of PPARc in humans is indicated by several loss-of-function mutations in the PPAR-c gene that cause lipodystrophy and diabetes in humans ([65–67], reviewed in [68]). In addition to the transcription factors described above that promote adipogenesis, there are several other transcription factors that have been shown to have positive or negative effects on adipocyte differentiation (reviewed in [4]). Section 1.3 reveals speciﬁc target genes that have been identiﬁed as targets of the positive adipogenic transcriptional activators described above. The identiﬁcation and characterization of these target genes has provided critical information in understanding the role of these proteins in both differentiating and mature fat cells.

1.3 Identification of Adipocyte Transcription Factor Target Genes

As stated in Section 1.2, adipogenesis occurs as a result of a transcriptional cascade that involves the tightly regulated induction of key transcription factors, including C/EBP-b, C/EBP-d, C/EBP-a, SREBP-1c, STAT-5, and PPAR-c. These adipogenic factors, in turn, induce the expression of various adipocyte genes that are important in conferring lipid accumulation, insulin sensitivity, and endocrine properties to mature adipocytes. In addition, many of these adipogenic factors possess the ability to regulate one anothers gene expression. These interactions add complexity to the pathways that regulated lipid and glucose metabolism. Hence, it is critical to identify the target genes of these adipogenic transcription factors in order to understand adipocyte gene expression and fat cell function. 1.3.1 C/EBP Target Genes

The three isoforms of the C/EBP family are differentially regulated and expressed at speciﬁc times in accordance with each proteins regulatory role in adipogenesis.

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C/EBP-b and -d are expressed very early in adipocyte differentiation in 3T3-L1 adipocytes, in contrast to C/EBP-a, whose expression is limited to the late phase of differentiation [43]. Both C/EBP-b and -d play important roles in the induction of PPAR-c gene expression [21], as functional C/EBP regulatory elements have been identiﬁed in the promoter of the PPAR-c gene [69]. In addition, investigators have shown that these C/EBP regulatory elements are also in the promoter of the C/EBP-a gene [70], supporting a role of C/EBP-b and -d in regulating the expression of C/EBPa [43, 44]. Hence, C/EBP-b and -d can modulate the transcriptional regulation of other primary players in adipogenesis (i.e., PPAR-c and C/EBP-a). C/EBP-b has also been shown to regulate other fat-speciﬁc genes, as the binding of C/EBP-b to C/EBP response elements in the adiponectin promoter is necessary for its transcription [71]. C/EBP-a plays a more diverse role in adipocyte gene expression, as it regulates more target genes that are inﬂuential not only in adipogenic pathways, but also genes that are important in insulin signaling and the production of endocrine hormones. C/EBP-a has the ability to affect the expression of other transcription factors that are necessary for adipogenesis. For example, C/EBP-a can directly modulate the transcription of PPAR-c through C/EBP response elements within the PPAR-c2 promoter [69, 72]. C/EBP-a can also transactivate the promoters of both aP2 [73, 74] and stearoyl-CoA desaturase (SCD) genes [73] during 3T3-L1 differentiation via speciﬁc sequence elements in the promoter. In addition, C/EBP-a can transactivate the insulin-responsive GLUT4 promoter in vitro [75] and the insulin receptor gene in vivo [76]. These studies clearly demonstrate the importance of this transcription factor in affecting glucose metabolism and insulin signaling. Adipocytes also have important endocrine functions, including the production and secretion of leptin and adiponectin. One study has demonstrated that C/EBP-a is a transcriptional activator of the leptin promoter via a functional C/EBP-binding site [77]. Similar to C/EBP-b, C/EBP-a can activate transcription of the murine adiponectin promoter via C/EBPbinding sites [78] and can modulate adiponectin expression in mature human adipocytes via interactions with C/EBP response elements in an intronic enhancer [79]. In summary, these studies indicate C/EBP-a can modulate the expression of important adipocyte transcription factors, key endocrine products of adipocytes, and genes associated with lipid and glucose metabolism. 1.3.2 SREBP-1 Target Genes

The identiﬁcation of ADD-1 [8], later termed SREBP-1 due to its importance in cholesterol metabolism [7], is an important modulator of adipocyte gene expression. SREBP-1/ADD-1 appears to be responsible for the differentiation-dependent induction of fatty acid synthase (FAS) during adipogenesis [39]. In addition, one study has shown that C/EBP-b is under the direct control of SREBP-1c in 3T3-L1 adipocytes. In this study, SREBP-1c was shown to activate transcription via SREBP-binding sites in the C/EBP-b promoter [80]. Adiponectin gene expression has also been shown to be directly regulated by SREBP-1c, which binds sterol regulatory elements in the adiponectin promoter in mouse adipocytes in vitro and in vivo [81]. These data

1.3 Identification of Adipocyte Transcription Factor Target Genes

support SREBP-1cs regulatory involvement in mature adipocytes by mediating the expression of several adipogenic genes. Since SREBP-1 null mice are not defective in adipogenesis [37], the primary function of this transcription factor in adipocytes appears to be the regulation of genes associated with lipid accumulation and the endocrine properties of adipocytes. 1.3.3 PPAR-c Target Genes

PPAR-c, which is considered the master regulator of adipogenesis, exerts its effects directly in adipocytes, but is also known to be important in other cell types. PPAR-c can modulate the expression of a variety of genes that differ in their metabolic function. The ﬁrst PPAR-c target gene identiﬁed was the aP2 gene. The aP2 gene contains a functional peroxisome proliferator response element (PPRE) within its promoter that plays a central role in its induction during adipogenesis [6]. Several studies also support a role for PPAR-c in the induction of other adipocyte transcription factors, including C/EBP-a and STAT-5A. It is known that PPAR-c and C/EBP-a can regulate each others expression during adipogenesis [48], and the crossregulation of PPAR-c and C/EBP-a is required for efﬁcient adipogenesis [82]. Recently, PPAR-c was also found to regulate expression of STAT-5A through several identiﬁed PPREs [83]. Taken together, these studies demonstrate cross-talk between three transcription factors that are highly induced during adipogenesis and are known to play a role in adipose tissue development in vivo [28, 42, 50]. Through its transcriptional regulation of various genes, PPAR-c is functional in many biological processes, including lipid metabolism. PPAR-c regulates genes involved in lipoprotein metabolism, including lipoprotein lipase (LPL) whose expression is modulated in adipocytes via a PPRE within its promoter [84]. In addition, PPAR-c can directly modulate the transcription of hormone-sensitive lipase (HSL) [85], which is an important contributor to lipolysis. The malic enzyme, whose product is involved in lipogenesis, is a PPAR-c target gene [86]. The expression of acyl-CoA synthetase, which converts fatty acids to acyl-CoA for subsequent b-oxidation, is induced in differentiated adipocytes [87] and its expression can be modulated by a PPRE [88]. A functional PPRE in the murine fatty acid transport protein (FATP) gene is modulated by PPAR-c in 3T3-L1 adipocytes [89]. Gene expression of another fatty acid transporter gene, fatty acid translocase (FAT) [90], is induced by PPAR-c in adipose tissue in vivo [91]. The expression of SCD, an enzyme involved in fatty acid biosynthesis, is modulated by a PPRE promoter element [92] and may be regulated by PPAR-c in adipose tissue in vivo [90]. Two lipid droplet-associating proteins are also known PPAR-c target genes in adipocytes. A functional PPRE has been characterized in the murine perilipin gene promoter [93] and in the lipid droplet-associating protein, S3-12 [94]. Hence, PPAR-c regulates genes that play a role in lipogenesis, lipolysis, and lipid droplet formation. PPAR-c can also directly control genes whose expression contributes to glucose homeostasis and insulin signaling. The expression of phosphoenolpyruvate

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carboxykinase (PEPCK), a key enzyme in gluconeogenesis, is modulated by a PPAR-c-binding site in its promoter [95]. The insulin-stimulated glucose transporter, GLUT4, is upregulated by PPAR-c activation in rat adipose tissue in vivo [90] and in NIH-3T3 cells [96]. Rev-Erba (NR1D1) is another nuclear receptor whose expression is also induced during adipocyte differentiation [97]. Rev-Erba expression is induced by PPAR-c activation in rat adipose tissues in vivo as well as in 3T3-L1 adipocytes in vitro and this transcriptional induction occurs via a PPAR response element in the Rev-Erba promoter [98]. Therefore, Rev-Erba is a target gene of PPAR-c in adipose tissue, and is another member of the nuclear receptor superfamily that is modulated during adipogenesis and may contribute to the regulation of adipocyte gene expression. Although many studies on PPAR-c focus on the identiﬁcation of target genes in white adipocytes, studies in brown adipocytes have shown that the uncoupling protein (UCP)-1 gene contains a PPRE that is bound and activated by PPAR-c [99]. It has been known for several years that PPAR-c ligands can increase the expression and plasma concentrations of adiponectin [100], and more recent study have observed that PPAR-c induces adiponectin in adipocytes via a PPRE in the human adiponectin promoter [101]. All of these results indicate that PPAR-c is not only required for regulating transcription factors that drive adipogenesis, but this nuclear receptor is important for modulating the adiponectin gene, which confers an important endocrine function to adipocytes. Overall, the identiﬁcation of PPAR-c target genes in adipocytes has revealed that this nuclear receptor regulates a variety of genes that are involved in adipogenesis, insulin signaling, glucose homeostasis, and lipid metabolism. 1.3.4 STAT-5 Target Genes

It is widely accepted that STATproteins have cell-speciﬁc functions. Studies on STAT5 proteins have revealed their importance in adipogenesis in vitro and in vivo [27, 28]. Recent studies have focused on the identiﬁcation of STAT-5 target genes in adipocytes. The promoter for acyl-CoA oxidase, the rate-limiting enzyme in peroxisomal fatty acid b-oxidation, contains a STAT-5-binding site that modulates its gene expression in fat cells [102]. Another study indicates that growth hormone also exerts stimulatory effects on adipogenesis through STAT-5A and -5B by enhancing the transcriptional activity of C/EBP-b/d and PPAR-c [31]. Transfection studies have demonstrated that aP2 promoter activity can be activated by STAT-5 [103]. Yet other studies have shown that STAT-5 mediates the inhibition of aP2 expression in rat primary preadipocytes [104]. This was the ﬁrst study to suggest that STAT-5 proteins could act as transcriptional repressors. This observation has been supported by another study that revealed STAT-5A could act as a transcriptional repressor in adipocytes. A STAT-5A-binding site in the murine FAS promoter has been shown to mediate the repression of FAS transcription that occurs with prolactin treatment [105]. In addition to modulation of genes associated with lipid metabolism, STAT-5 can also modulate a gene associated with glucose metabolism. The gene for

1.3 Identification of Adipocyte Transcription Factor Target Genes

pyruvate dehydrogenase kinase (PDK)-4, a known regulator of glycolysis, is highly induced in adipocytes by growth hormone or prolactin (PRL) in a STAT-5-dependent manner [106]. Under these conditions, the induction of PDK-4 is accompanied by insulin resistance. It is well known that PRL and growth hormone are important modulators of lipid metabolism, and are also potent inducers of STAT-5 in adipocytes [103, 107]. Hence, many of the metabolic actions of these hormones could be mediated by STAT-5s direct modulation of target genes. Unfortunately, relatively few STAT-5 target genes have been identiﬁed in adipocytes. Nonetheless, we hypothesize that several other STAT-5A target genes will be identiﬁed that play a role in lipid or glucose metabolism. 1.3.5 Summary

It is clear that there are several transcription factors that are highly expressed during adipogenesis and are critical not only for lipid accumulation, but to confer insulin sensitivity and endocrine properties to mature adipocytes. The primary function of these proteins is to directly modulate target genes that contribute to the adipocyte phenotype (reviewed in Table 1.1). Although much emphasis has been placed on C/EBP-a and PPAR-c, there are clearly several other transcription factors that are important, including STAT-5 proteins and KLFs. However, very few target genes for these transcription factors have been identiﬁed. Interestingly, it is still not known which transcription factor(s) confer the adipocyte speciﬁc expression leptin. In the future, the identiﬁcation of new or known adipocyte transcription factors and their target genes will be essential in enhancing our knowledge of adipogenesis and indicating what transcriptional modulators are required for the maintenance of lipid laden, insulin-sensitive, hormone-secreting fat cells.

Table 1.1 Target genes of key adipocyte transcription factors.

Adipogenic transcription factors

Characterized target genes

C/EBP-b C/EBP-d C/EBP-a

PPAR-c [21, 69], C/EBP-a [43, 70], adiponectin [71] PPAR-c [21, 69], C/EBP-a [43, 70] PPAR-c [69, 72], aP2 [73, 74], SCD [73], GLUT4 [75], insulin receptor [76], leptin [77], adiponectin [78, 79] FAS [39], C/EBP-b [80], adiponectin [81] aP2 [6], C/EBP-a [48], STAT-5A [83], LPL [84], HSL [85], malic enzyme [86], acyl-CoA synthetase [88], FATP [89], FAT [91], SCD [90, 92], perilipin [93], S3-12 [94], PEPCK [95], GLUT4 [90–96], Rev-Erba [97, 98], UCP-1 [99], adiponectin [100, 101] acyl-CoA oxidase [102], C/EBP-b/d [31], PPAR-c [31], aP2 [103, 104], FAS [105], PDK [106]

SREBP-1 PPAR-c

STAT-5

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PPARgamma: a uniﬁed pathway. Genes Dev., 16, 22–26. Olsen, H. and Haldosen, L.A. (2006) Peroxisome proliferator-activated receptor gamma regulates expression of signal transducer and activator of transcription 5A. Exp. Cell Res., 312, 1371–1380. Schoonjans, K., Peinado-Onsurbe, J., Lefebvre, A.M., Heyman, R.A., Briggs, M., Deeb, S., Staels, B., and Auwerx, J. (1996) PPARalpha and PPARgamma activators direct a distinct tissue-speciﬁc transcriptional response via a PPRE in the lipoprotein lipase gene. EMBO J., 15, 5336–5348. Yajima, H., Kobayashi, Y., Kanaya, T., and Horino, Y. (2007) Identiﬁcation of peroxisome-proliferator responsive element in the mouse HSL gene. Biochem. Biophys. Res. Commun., 352, 526–531. Castelein, H., Gulick, T., Declercq, P.E., Mannaerts, G.P., Moore, D.D., and Baes, M.I. (1994) The peroxisome proliferator activated receptor regulates malic enzyme gene expression. J. Biol. Chem., 269, 26754–26758. Martin, G., Schoonjans, K., Lefebvre, A.M., Staels, B., and Auwerx, J. (1997) Coordinate regulation of the expression of the fatty acid transport protein and acyl-CoA synthetase genes by PPARalpha and PPARgamma activators. J. Biol. Chem., 272, 28210–28217. Schoonjans, K., Watanabe, M., Suzuki, H., Mahfoudi, A., Krey, G., Wahli, W., Grimaldi, P., Staels, B., Yamamoto, T., and Auwerx, J. (1995) Induction of the acyl-coenzyme A synthetase gene by ﬁbrates and fatty acids is mediated by a peroxisome proliferator response element in the C promoter. J. Biol. Chem., 270, 19269–19276. Frohnert, B.I., Hui, T.Y., and Bernlohr, D.A. (1999) Identiﬁcation of a functional peroxisome proliferator-responsive element in the murine fatty acid transport protein gene. J. Biol. Chem., 274, 3970–3977. Singh, A.H., Liu, S., Crombie, D.L., Boehm, M., Leibowitz, M.D., Heyman, R.A., Depre, C., Nagy, L., Tontonoz, P.,

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and Davies, P.J. (2001) Differential effects of rexinoids and thiazolidinediones on metabolic gene expression in diabetic rodents. Mol. Pharmacol., 59, 765–773. Motojima, K., Passilly, P., Peters, J.M., Gonzalez, F.J., and Latruffe, N. (1998) Expression of putative fatty acid transporter genes are regulated by peroxisome proliferator-activated receptor alpha and gamma activators in a tissue- and inducer-speciﬁc manner. J. Biol. Chem., 273, 16710–16714. Miller, C.W. and Ntambi, J.M. (1996) Peroxisome proliferators induce mouse liver stearoyl-CoA desaturase 1 gene expression. Proc. Natl. Acad. Sci. USA, 93, 9443–9448. Nagai, S., Shimizu, C., Umetsu, M., Taniguchi, S., Endo, M., Miyoshi, H., Yoshioka, N., Kubo, M., and Koike, T. (2004) Identiﬁcation of a functional peroxisome proliferator-activated receptor responsive element within the murine perilipin gene. Endocrinology, 145, 2346–2356. Dalen, K.T., Schoonjans, K., Ulven, S.M., Weedon-Fekjaer, M.S., Bentzen, T.G., Koutnikova, H., Auwerx, J., and Nebb, H.I. (2004) Adipose tissue expression of the lipid droplet-associating proteins S3-12 and perilipin is controlled by peroxisome proliferator-activated receptor-gamma. Diabetes, 53, 1243–1252. Tontonoz, P., Hu, E., and Spiegelman, B.M. (1995) Regulation of adipocyte gene expression and differentiation by peroxisome proliferator activated receptor gamma. Curr. Opin. Genet. Dev., 5, 571–576. Wu, Z., Xie, Y., Morrison, R.F., Bucher, N.L., and Farmer, S.R. (1998) PPARgamma induces the insulindependent glucose transporter GLUT4 in the absence of C/EBPalpha during the conversion of 3T3 ﬁbroblasts into adipocytes. J. Clin. Invest., 101, 22–32. Chawla, A. and Lazar, M.A. (1993) Induction of Rev-ErbA alpha, an orphan receptor encoded on the opposite strand of the alpha-thyroid hormone receptor gene, during adipocyte differentiation. J. Biol. Chem., 268, 16265–16269.

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98 Fontaine, C., Dubois, G., Duguay, Y.,

Helledie, T., Vu-Dac, N., Gervois, P., Soncin, F., Mandrup, S., Fruchart, J.C., Fruchart-Najib, J., and Staels, B. (2003) The orphan nuclear receptor RevErbalpha is a peroxisome proliferatoractivated receptor (PPAR) gamma target gene and promotes PPARgammainduced adipocyte differentiation. J. Biol. Chem., 278, 37672–37680. 99 Sears, I.B., MacGinnitie, M.A., Kovacs, L.G., and Graves, R.A. (1996) Differentiation-dependent expression of the brown adipocyte uncoupling protein gene: regulation by peroxisome proliferator-activated receptor gamma. Mol. Cell. Biol., 16, 3410–3419. 100 Maeda, N., Takahashi, M., Funahashi, T., Kihara, S., Nishizawa, H., Kishida, K., Nagaretani, H., Matsuda, M., Komuro, R., Ouchi, N., Kuriyama, H., Hotta, K., Nakamura, T., Shimomura, I., and Matsuzawa, Y. (2001) PPARgamma ligands increase expression and plasma concentrations of adiponectin, an adipose-derived protein. Diabetes, 50, 2094–2099. 101 Iwaki, M., Matsuda, M., Maeda, N., Funahashi, T., Matsuzawa, Y., Makishima, M., and Shimomura, I. (2003) Induction of adiponectin, a fatderived antidiabetic and antiatherogenic factor, by nuclear receptors. Diabetes, 52, 1655–1663.

102 Coulter, A.A. and Stephens, J.M. (2006)

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STAT5 activators modulate acyl CoA oxidase (AOX) expression in adipocytes and STAT5A binds to the AOX promoter in vitro. Biochem. Biophys. Res. Commun., 344, 1342–1345. Nanbu-Wakao, R., Fujitani, Y., Masuho, Y., Muramatu, M., and Wakao, H. (2000) Prolactin enhances CCAAT enhancerbinding protein-beta (C/EBP beta) and peroxisome proliferator-activated receptor gamma (PPAR gamma) messenger RNA expression and stimulates adipogenic conversion of NIH-3T3 cells. Mol. Endocrinol., 14, 307–316. Richter, H.E., Albrektsen, T., and Billestrup, N. (2003) The role of signal transducer and activator of transcription 5 in the inhibitory effects of GH on adipocyte differentiation. J. Mol. Endocrinol., 30, 139–150. Hogan, J.C. and Stephens, J.M. (2005) The regulation of fatty acid synthase by STAT5A. Diabetes, 54, 1968–1975. White, U.A., Coulter, A.A., Miles, T.K., and Stephens, J.M. (2007) The STAT5Amediated induction of pyruvate dehydrogenase kinase 4 expression by prolactin or growth hormone in adipocytes. Diabetes, 56, 1623–1629. Balhoff, J.P. and Stephens, J.M. (1998) Highly speciﬁc and quantitative activation of STATs in 3T3-L1 adipocytes. Biochem. Biophys. Res. Commun., 247, 894–900.

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2 Cellular and Molecular Basis of Functional Differences among Fat Depots Thomas Thomou, Tamara Tchkonia, and James L. Kirkland 2.1 Introduction

Variation in fat distribution is remarkable. Subjects matched for body mass index (BMI) sometimes differ vastly in how much fat is subcutaneous or visceral, or even the particular subcutaneous regions affected. Fat redistribution, especially from subcutaneous to visceral fat depots, has received increasing attention as a risk factor for atherosclerosis and other serious morbidities. The case for considering fat cells and their progenitors, preadipocytes, from different fat depots to be distinct cell subtypes that contribute to regional variation in function will be considered, as will molecular mechanisms responsible (Figure 2.1). 2.1.1 Fat Tissue Function

Different fat depots have distinct roles (Table 2.1). With respect to energy storage, subcutaneous fat, which can expand outward and is not restricted by the anatomic constraints that limit visceral fat growth, is specialized to provide long-term fuel storage. It is the main source of leptin, which signals the state of long-term fat stores to the brain [1–5]. The greater development of subcutaneous fat in females than males may have evolved to facilitate steady access to calories during pregnancy [6]. Visceral fat provides short-term storage and rapid access to calories that can be processed quickly by the liver. Larger visceral fat stores may have evolved in males to allow rapid access to energy required during hunting [6]. Fat is important in immunity. Complement components, cytokines, and chemokines, including tumor necrosis factor (TNF)-a, interleukin-6, and macrophage chemotactic factors, are released by adipose cells. High local fatty acid concentrations that are toxic to pathogenic bacteria, fungi, and other cell types contribute to the protective function of fat tissue. Preadipocytes and fat cells are resistant to cytotoxic levels of fatty acids [7]. Perhaps because of their secretory proﬁle and high local fatty

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Mesenchymal Progenitor Cell Determination

Circulating Progenitors

Replication

Preadipocyte (2 subtypes) Differentiation Apoptosis

Dedifferentiation

Senescence Lipid Accumulation

Lipolysis Fat Cell Apoptosis/Necrosis

Figure 2.1 Fat tissue cell dynamics. Preadipocytes arise from multipotent mesenchymal progenitor cells, which may also be able to develop into muscle, bone, cartilage, and macrophages, and possibly from circulating progenitors. After becoming committed to the fat cell lineage through GATA, Wnt, and Notch signaling components, preadipocytes can replicate, differentiate into fat cells or possibly reverse determination to become multipotent progenitors again. There are at least two preadipocyte subtypes, one being more capable

of replication, differentiation into fat cells, and resisting apoptosis than the other. Cells of each subtype can convert into the other. Preadipocytes can be induced to differentiate by IGF-1, glucocorticoids, a burst of cAMP generation, fatty acids, and other factors. Differentiated fat cells can accumulate more lipid, release fatty acids through lipolysis, be removed through apoptosis, senescence, or necrosis, or possibly dedifferentiate into preadipocytes. The preadipocyte cell dynamic processes that vary among fat depots are underlined.

acid levels, infections and metastases are uncommon in fat depots. Subcutaneous fat acts as a barrier to infection entering through the skin. The greater omentum can migrate and wall off infection from ruptured viscera. The omentum hangs down from the stomach in bipeds, migrating around the abdomen during body movement Table 2.1 Fat tissue function.

Subcutaneous

Omental

Regeneration/wound response

long-term/protection against lipotoxicity ü mechanical ( ! growth to provide cushioning) insulation signaling long-term stores replication to ﬁll voids

Anatomic constraints

none

rapid fatty acid release to central circulation üüü patching (adhesion/attachment, hemostasis) — injury response/maintain blood pressure bandage function/creeping fat/" other cell types very restricted, especially during pregnancy

Fuel storage/metabolic Innate immune Protective Temperature Endocrine/paracrine

2.1 Introduction

and peristalsis. In quadrupeds, epididymal or paraovarian fat, which is attached at the highest point in the abdomen, serves a function analogous to omental fat in bipeds. Omental fat releases attachment, pressor, and hemostatic factors, such as plasminogen-activated inhibitor-1, in addition to inﬂammatory cytokines and chemokines. It acts as a medicated bandage in the event of a perforated viscus or other intraabdominal catastrophe. Perhaps these protective functions of omental fat switch into contributing to metabolic syndrome when this depot is enlarged in obesity – a state rare in nature. Mesenteric fat may help to prevent systemic spread of enteral infections [8]. Mesenteric fat responds to inﬂammation by growing, perhaps to wall inﬂamed viscera off from the abdominal cavity. Indeed, radiologists and surgeons use creeping fat to identify segments of bowel affected by inﬂammatory bowel disease (Crohns disease, ulcerative colitis). Certain fat depots have a cushioning function, including fat over the joints, palms, and soles (the depots preserved in Berardinelli–Seip lipodystrophy [9]), as well as fat around the heart and kidneys, and retro-orbital fat. Fat tissue can grow in response to mechanical stress, adding to its cushioning function, reminiscent of the close association between preadipocytes and bone progenitors [10]. White fat has an insulating effect and is ideally situated under the skin to prevent heat loss. Brown fat, which is present mainly in central, deep fat depots surrounding the great blood vessels in infants, generates heat. Bone marrow fat cells produce paracrine regulators of blood cell generation [11, 12]. Bone marrow preadipocytes and osteoblasts can turn into each other under particular conditions [13–15]. Thus, in addition to storage of calories, fat depots are specialized to carry out distinct protective and regenerative functions. 2.1.2 Diversity in Fat Distribution

Environmental, epigenetic, genetic, and disease- and drug-associated processes likely combine to cause diversity in fat distribution. Environmental factors associated with increased central fat include physical inactivity, low socioeconomic status (or the stress associated with this), and poor education [16, 17]. Smoking leads to development of central obesity that is reversed after smoking cessation [16–19]. Moderate alcohol consumption is associated with decreased central obesity [17]. Drugs that alter fat distribution include sex steroids, glucocorticoids, and some AIDS protease inhibitors, which increase visceral fat while causing loss of subcutaneous fat from some areas [20–22]. Antidiabetic thiazolidinediones (TZDs; e.g., rosiglitazone, pioglitazone) increase total body fat, but do so by increasing subcutaneous fat, causing relative redistribution of fat from visceral to peripheral depots [23–25]. Although environmental factors have a substantial effect on fat distribution, heritable genetic and epigenetic mechanisms make an even stronger contribution. Of genetic factors, gender has the largest impact. Visceral fat constitutes approximately 20% of total fat in males, but only 6% in premenopausal females [26, 27]. Hormonal manipulation to convert males to females causes redistribution of fat

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from visceral to subcutaneous depots, while the opposite occurs in female-tomale transsexuals [28, 29]. Although the postmenopausal state and male gender are associated with increased visceral fat, gender itself does not directly inﬂuence the relation between intra-abdominal fat and insulin sensitivity, underscoring the close ties between regional fat distribution and insulin responsiveness [30]. Effects of sex steroids on fat distribution are not straightforward. While testosterone administration causes visceral obesity in females, it reverses visceral obesity in hypogondal males [31]. Among subjects of the same gender, genetic factors account for over 60% of variation in visceral fat mass [32]. In monozygotic twins, 75% of variance in the increase of visceral fat after overfeeding is within twin pairs and only 25% among sets of twins [33], supporting a strong role for heritable factors. Over 50% of the variance in visceral fat mass has been attributed to single gene effects in segregation analyses, and only 20% to multifactorial effects [32]. While the genes most commonly accounting for altered regional fat distribution have not been identiﬁed, rare single gene mutations cause syndromes that profoundly affect distribution. The mutations that cause the Berardinelli–Seip and Dunnigan–Kobberling segmental lipodystrophy syndromes lead to altered fat distribution, diabetes, and hyperlipidemia [20, 34]. Berardinelli–Seip syndrome may be caused by a recessive mutation of the retinoid X receptor-a [35, 36], resulting in extensive subcutaneous fat loss except over the palms, soles, periarticular areas, and orbits [9, 35]. The Dunnigan–Kobberling syndrome results from a dominant mutation of lamin A/C nuclear envelope proteins, causing loss of subcutaneous fat except around the face and neck, with preserved or increased visceral fat. Thus, genetic factors, including gender and single- and multigene products, likely account for much of the variation in fat distribution. Fat tissue distribution changes throughout the lifespan, with increases in overall fat mass through middle age. In advanced old age, extensive loss of particular fat depots occurs, together with deposition of fat in marrow, liver, muscle, and other ectopic sites [37–44]. In old age, subcutaneous fat, especially in the extremities, tends to be lost ﬁrst, with thinning of the skin over the dorsa of the hands and development of spindly legs. Subcutaneous fat loss proceeds centrally in old age and visceral fat tends to be lost last. This relative preservation of visceral compared to subcutaneous fat in older individuals is associated with increased metabolic syndrome prevalence [45]. 2.1.3 Regional Differences in Fat Tissue Growth

Certain fat depots enlarge during development, in obesity, or after partial lipectomy, principally through increases in the size of existing fat cells. Others enlarge through both fat cell enlargement and differentiation of preadipocytes into new fat cells [44, 46, 47]. Subcutaneous fat, with its long-term storage function, tends to grow by increasing fat cell number, with fat cell sizes approximating the most thermodynamically stable size for triolein droplets (Figure 2.2). Visceral fat, with its short-term

2.1 Introduction

SUBCUTANEOUS

OMENTAL

Figure 2.2 Mechanisms of fat tissue growth during progression of obesity vary among depots. Human subcutaneous fat, specialized to provide long-term nutrient storage and not encumbered by the anatomic constraints faced by visceral fat, grows through increases in fat cell number. This is facilitated by the higher

potential of subcutaneous than visceral preadipocytes for replication and adipogenesis. Visceral fat enlarges through increases in fat cell size rather than number, consistent with its role in storing and releasing nutrients rapidly and its limited space for growth.

lipid storage function, changes in size principally through increases or decreases in fat cell size, rather than changes in fat cell number. After fat tissue is removed, fat reappears in remaining depots [48–50]. For example, after liposuction, breast enlargement occurs in women [49]. Subcutaneous fat increases when visceral fat is removed from experimental animals [51]. Conversely, when subcutaneous fat is removed, visceral fat increases. This raises concerns about potential effects of liposuction on visceral fat and perhaps contributes to visceral fat enlargement associated with the subcutaneous fat loss in lipodystrophies and aging. In experimental animals, not every fat depot enlarges to the same extent after removal of fat from other sites. For example, after removal of other fat pads, inguinal fat mass increases, but epididymal fat mass remains unchanged [51]. Unilateral removal of epididymal or inguinal fat pads from animals results in disproportionate enlargement of ipsilateral compared to contralateral depots [52]. Hormones are unlikely to be responsible for this asymmetric response, but it is plausible that neurological events are. The sympathetic nervous system appears to be particularly relevant. Adrenergic stimulation promotes lipolysis through activating b-receptors or inhibits lipolysis through a-receptors. Ratios of b- to a-receptors vary among different fat depots [53, 54]. Following sympathetic drive, fat stores are greatest in depots with high numbers or afﬁnities of a- relative to b-receptors. Regional variation in receptor densities could result from differences in nerve supply or the inherent properties of adipocytes. Indeed, these mechanisms may combine to cause functional differences among depots. Norepinephrine inhibits replication of cultured preadipocytes [55], while sympathetic denervation of fat tissue results in increased fat cell number [56]. Thus, although total fat mass increases toward a set point, individual fat depots exhibit distinct responses to lipectomy.

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2.1.4 Disease Associations

Accumulation of central fat is associated with elevated risk of diabetes, hypertension, atherosclerosis, dyslipidemia, and cancers [6, 27, 50–64]. Central obesity is more closely associated with insulin resistance than total body fat [24, 62–64]. Even lean subjects (with a nonobese BMI) who have a high ratio of central to peripheral fat are insulin resistant [30]. Whether impaired insulin sensitivity associated with central fat is related mainly to increased visceral fat [24, 62–64] or increased abdominal subcutaneous together with visceral fat [57, 58] has been debated. The deep layer of abdominal subcutaneous fat is more strongly associated with insulin resistance than the superﬁcial layer [59, 60]. Furthermore, increased visceral fat in computed tomography and magnetic resonance imaging studies is related to decreased antilipolytic effect of insulin, but this relation does not hold for abdominal subcutaneous fat [61]. Femoral subcutaneous fat might actually be protective [62]. Thus, increased visceral fat may be a powerful risk factor in central obesity, with further risk being added by abdominal subcutaneous fat [63]. Which of the major visceral fat depots (mesenteric and omental) is most closely associated with metabolic disease is not yet known, but venous drainage and preadipocyte and fat cell characteristics vary between these intra-abdominal depots [64–66]. The adverse metabolic impact of visceral obesity may be due to (i) an underlying process that predisposes both to visceral obesity and metabolic dysfunction, (ii) the location of visceral fat depots (with venous drainage through the liver and pancreas, increased intra-abdominal pressure), or (iii) inherent differences among fat depots in adipocyte characteristics, including their cytokine secretory proﬁles [31, 67]. In support of the ﬁrst mechanism, both central obesity and insulin resistance occur in mice with mutations that interfere with insulin receptors in muscle [68, 69]. Injection of estrogen into the central nervous system without altering systemic estrogen levels affects both insulin sensitivity and fat distribution [70]. In support of the second mechanism, some of the venous drainage of visceral fat, particularly omental fat, is through the portal system. Therefore, metabolic, paracrine, and hormonal products released by visceral fat could affect the liver and pancreas directly. For example, high fatty acid concentrations impede hepatic insulin signaling [71] and increase hepatic gluconeogenesis [6], contributing to insulin resistance. Visceral obesity is associated with increased intra-abdominal pressure that may provoke sympathetic activity through effects on the kidneys [72]. In support of the third mechanism, many inherent differences in preadipocyte and fat cell function have been found among depots, as reviewed in Section 2.2. These three potential mechanisms likely combine to produce the metabolic effects of visceral obesity. Does selectively reducing visceral fat improve health? While the answer is not yet clear in humans, in rats surgical removal of visceral fat enhances hepatic insulin sensitivity and decreases hepatic glucose production (although venous drainage of epididymal and perinephric depots in rodents differs from that of omental fat in humans), while removing subcutaneous fat from hamsters has the opposite effect [73–75]. If this is also true in humans, substantial improvement in insulin

2.2 Physiology

sensitivity and metabolic risk proﬁle might be feasible if drugs that selectively reduce visceral fat could be developed. A number of acquired diseases and drugs affect fat distribution. Inﬂammatory bowel disease results in local accumulation of fat tissue [8]. Limb paralysis results in loss of muscle and increased fat in the affected limb [76]. TZDs result in subcutaneous fat growth, but have little effect on visceral fat [23–25]. HIV protease inhibitors are associated with loss of fat over the face and extremities, but accumulation of dorsal subcutaneous and visceral fat [21, 22]. Sex steroids are associated with fat redistribution. Subcutaneous and visceral adipocytes differ in extent of their responses to glucocorticoids. Subcutaneous fat cells express fewer glucocorticoid receptors and exhibit less lipoprotein lipase activation in response to glucocorticoids [77, 78]. Activity of the enzyme that converts cortisone to the more actively adipogenic cortisol, 11b-hydroxysteroid dehydrogenase (HSD)-1, is higher in visceral fat [79, 80]. Increased circulating cortisone would therefore be predicted to cause more growth of visceral than subcutaneous fat, as occurs in Cushings syndrome. Fat tissue-speciﬁc overexpression of 11b-HSD-1 increases 11b-HSD-1 in all fat depots. This results in visceral obesity in transgenic mice, but has little effect on subcutaneous fat abundance [81]. Furthermore, glucocorticoids that bypass 11b-HSD-1 and inhibit endogenous glucocorticoid production, such as dexamethasone, still cause fat redistribution. Therefore, regional differences in responses to glucocorticoids downstream of 11b-HSD-1 likely contribute to interdepot variation in fat growth.

2.2 Physiology

For many years, it has been recognized that physiological characteristics of fat tissue isolated from various depots are remarkably different [43, 82–85]. Fat cells isolated from different depots vary in size, responses to insulin and lipolytic agents, lipoprotein binding, fatty acid transfer, production of secreted proteins, and other features [1–4, 6, 51, 77, 86–110]. Visceral fat cells undergo more fatty acid turnover and lipolysis than subcutaneous cells [102, 103]. This ﬁts with clinical studies indicating increased fatty acid and triglyceride turnover in abdominal obesity [111–114]. Greater capacity of omental cells for hormone-stimulated lipolysis is related to presence of increased numbers of b-receptors and blunted a-receptor-mediated decline in cAMP levels compared to subcutaneous cells [97, 104, 106, 115]. Visceral adipocytes are also less responsive to antilipolytic effects of insulin [86, 103, 104, 107, 116]. Higher abundance of the exon 11 minus insulin receptor isoform in visceral than subcutaneous fat is associated with reduced insulin receptor afﬁnity [1]. Reduced insulin receptor substrate-1 responsiveness also contributes to reduced insulin sensitivity and increased lipolysis in omental adipocytes [77, 78]. Increased lipolysis is balanced by increased fatty acid transfer into these cells, maintaining stability in cell size [117]. Increased visceral fat cell fatty acid ﬂux may contribute to insulin resistance and hyperinsulinemia, since

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these fatty acids travel directly to the liver where they stimulate gluconeogenesis and reduce insulin removal [86, 118–121]. Acylation-stimulating protein (ASP) promotes human fat cell triglyceride synthesis [87]. Post-translational modiﬁcation of complement factor C3a, which is secreted by fat cells, leads to ASP generation. C3a mRNA abundance is higher in visceral than abdominal subcutaneous fat [88]. Higher expression of C3a and ASP in visceral than subcutaneous fat may act to counterbalance the greater lipolytic capacity of visceral fat by promoting triglyceride synthesis, contributing to higher fatty acid ﬂux in visceral fat. Leptin mRNA levels and secretion are higher in abdominal subcutaneous than omental fat tissue [1–3], even after 12 days in culture [4, 5], and circulating leptin levels are correlated more closely with subcutaneous than visceral fat mass [89]. Since leptin signals the state of fat stores to the brain, changes in visceral fat content may exert less effect on feeding regulatory responses mediated by leptin than subcutaneous fat. In this respect, visceral fat mass may be subject to less central nervous system regulation than peripheral fat. Leptin also has an insulin-sensitizing effect on muscle [90–92]. Thus, greater secretion of leptin by subcutaneous than visceral fat, coupled with the proximity of subcutaneous fat to muscle, may contribute to low insulin sensitivity in central obesity. Cholesterol ester transfer protein (CETP), which mediates cholesterol metabolism, is produced by fat tissue [93]. CETP mRNA levels are higher in abdominal subcutaneous than omental fat [88]. While CETP mRNA levels may not predict protein secretion, subcutaneous fat cells have higher cholesterol content and highdensity lipoprotein-binding capacities than omental cells [122], suggesting a role of CETP release in regional differences in cholesterol metabolism. Angiotensinogen is produced by fat cells. Fat tissue contains all the machinery needed to convert angiotensinogen into angiotensin II [94–96]. Angiotensin II is involved in preadipocyte differentiation into fat cells, but may also contribute to the hypertension frequently associated with obesity. More angiotensinogen mRNA is present in omental than subcutaneous fat tissue and increased waist-to-hip ratios are associated with increased angiotensinogen mRNA [88, 97]. Visceral fat angiotensinogen production may contribute to the hypertension common in the metabolic syndrome. Thus, fat isolated from various regions differs in metabolic pathways that impact morbidity. An understanding of mechanisms responsible for these differences could be invaluable in developing interventions to reduce this morbidity. 2.2.1 Contribution of Inherent Cell Dynamic Mechanisms to Regional Differences

Is regional variation solely a result of inﬂuences extrinsic to adipose cells (e.g., hormonal and paracrine microenvironment, local nutrient availability, innervation, anatomic constraints, or regional variation in abundance of nonadipose cell types (macrophages, endothelial cells) or do differences in inherent characteristics of adipose cells also contribute? As evident from the foregoing, extrinsic factors are

2.2 Physiology

clearly important. There is mounting evidence that inherent properties of fat cells and their progenitors, preadipocytes, also contribute (Figure 2.1). New fat cells continue to appear from preadipocytes in adulthood [98–100]. Preadipocytes cultured under identical conditions, originating from different depots from the same individuals, retain distinct, fat depot-dependent characteristics after weeks in culture [47, 64–66, 101–104, 115, 116]. A problem with studies of effects of fat depot origin on preadipocyte function is that interdepot differences in extent of contamination of primary cultures with nonadipose cell types, such as mesothelial cells or macrophages, are a potential cause of apparent differences. However, interdepot differences in fatty acid uptake have been found at the single-cell level in cultured preadipocytes matched for lipid content [117]. Furthermore, regional variation in developmental gene expression proﬁles and capacities for adipogenesis, replication, and susceptibility to apoptosis persist for many cell generations in strains derived from single human preadipocytes, as will be reviewed below [64–66]. 2.2.2 Preadipocyte Function

The main job of preadipocytes is to differentiate into fat cells when required. Fat tissue has to be able to expand in times of nutrient excess to sequester cytotoxic fatty acids as less toxic triglycerides. Fat tissue storage capacity can increase acutely through increases in fat cell size. Once fat cells get too large, they may become susceptible to breakage and necrosis with ensuing macrophage inﬁltration of fat tissue [106]. Fat tissue expansion in times of sustained nutrient availability is most efﬁciently achieved through increases in fat cell number. This is an especially prominent mechanism of fat tissue expansion in subcutaneous depots. To allow such fat tissue expansion, the preadipocyte pool is large: preadipocytes account for 15–50% of cells in fat tissue, one of the largest progenitor pools in the body [99]. Fat tissue also turns over regularly, with fat cells having a half life estimated to be approximately 3 months [100] – another reason the preadipocyte pool needs to be large. Beyond giving rise to new fat cells, fat cell progenitors are an important cell type in their own right, actively producing paracrine factors, hormones, and metabolic signals in a manner distinct from that of differentiated fat cells [99, 107]. For example, most tissue-type plasminogen activator appears to be produced by preadipocytes rather than fat cells [108]. Furthermore, much of the immune function of fat tissue is attributable to preadipocytes, which react to bacterial antigens and can recruit macrophages [109]. Indeed, preadipocytes are so closely related to macrophages that their expression proﬁles are more like macrophages than fat cells. Preadipocytes even appear to be able to develop into macrophages, among other cell types [110, 111]. 2.2.3 Preadipocyte Replication

Preadipocytes from different fat depots differ in replicative potential [47, 66, 101–103, 112–114, 116, 123]. Preadipocytes from rat perirenal depots are capable of more

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extensive replication than from epididymal depots [47, 101, 103, 104, 116, 123]. These differences are evident in colonies arising from single perirenal or epididymal preadipocytes after 3 weeks in identical cell culture conditions [103]. This indicates these differences are inherent. Higher replicative potential of perirenal preadipocytes is reﬂected in a greater increase in perirenal than epididymal fat cell number during depot growth in vivo [47]. Human abdominal subcutaneous preadipocytes are capable of more extensive replication than omental cells, with mesenteric preadipocytes being intermediate [66, 113, 114]. As in the rat, these regional differences in replicative potential are retained in colonies derived from single preadipocytes after several weeks in culture, indicating that regional variation in inherent preadipocyte replicative potential occurs across mammals. Consistent with greater replicative potential of subcutaneous than omental preadipocytes, telomeres are shorter in subcutaneous than omental cells [66]. This increased replicative history in subcutaneous cells may be related to the long-term lipid storage function of subcutaneous fat and the expansion of subcutaneous fat cell numbers that occurs in obesity. To address whether preadipocytes from different depots are distinct, preadipocyte strains were produced from single abdominal subcutaneous, mesenteric, and omental human preadipocytes by stably expressing human telomere reverse transcriptase (hTERT) [66]. These strains can be subcultured repeatedly and retain capacity for differentiation, while primary preadipocyte adipogenesis and replication decline with subculturing. Single subcutaneous, mesenteric, and omental hTERT preadipocytes that had undergone 35 population doublings were cultured for 21 days. The number of subcutaneous clones that achieved an additional 14 or more population doublings during the 21 days was 2.5-fold higher than omental clones. Thus, regional differences in preadipocyte replicative potential are heritable for many cell generations, do not depend on presence of other cell types, and are independent of telomerase activity. Heritable, possibly epigenetic processes, likely contribute to regional variation in fat tissue function. 2.2.4 Differences in Adipogenesis among Depots

Human preadipocyte capacity for adipogenesis may vary among fat depots, as indicated in several studies [65, 66, 102, 112, 115, 124–126]. In other studies, differences were not apparent [114, 127, 128], perhaps because of variability among subjects, the time point at which extent of differentiation was assessed, or culture media. In the studies in which regional differences were found, human abdominal subcutaneous preadipocytes had greater capacity for adipogenesis than omental cells. If left long enough in differentiation-promoting medium, omental preadipocytes eventually catch up to subcutaneous cells, possibly obscuring regional differences evident earlier during differentiation and contributing to discrepancies among studies [65]. In the studies in which differences among depots were found, human abdominal subcutaneous preadipocytes differentiated to a greater extent in response to TZDs than omental cells [65, 115, 124–126, 129]. TZDs bind to and activate the adipogenic transcription factor, peroxisome proliferator-activated receptor (PPAR)-c.

2.2 Physiology

Consistent with this, TZDs promote accumulation of fat in subcutaneous, but not visceral depots [23, 24]. Indeed, regional differences in TZD responsiveness could be a mechanism contributing to the antidiabetic effects of TZDs, since excess visceral relative to subcutaneous fat is associated with glucose intolerance [85]. Markers of adipogenesis (lipid accumulation, expression of the fatty acid-binding protein, aP2, and glycerol-3-phosphate dehydrogenase (G3PD) activity) are highest in human abdominal subcutaneous preadipocytes, intermediate in mesenteric cells, and lowest in omental cells following induction of differentiation. This pattern was also observed with respect to expression of PPAR-c and the CCAAT/enhancerbinding protein C/EBP-a – transcription factors necessary for adipogenesis to proceed. The aP2 and G3PD promoters respond to PPAR- and C/EBP-a [122, 130–132]. The magnitudes of the regional differences in C/EBP-a and PPAR-c expression are consistent with regional differences in aP2 and G3PD. Furthermore, ectopic expression of C/EBP-a in omental preadipocytes enhances capacity to accumulate lipid, suggesting genes downstream of C/EBP-a in omental preadipocytes are capable of responding to overexpressed levels of this adipogenic regulator [65]. Thus, mechanisms upstream of the level of adipogenic transcription factors appear to contribute to regional variation in function. Regional variation in adipogenesis is also found in cultured rat preadipocytes, indicating this occurs across mammals and does not result from particular features of subjects selected for the human studies. Under various conditions that promote differentiation, rat perirenal preadipocytes accumulate more lipid and develop higher levels of expression of differentiation-dependent genes than epididymal preadipocytes [47, 103, 104, 116]. Interdepot variation in cultured rat preadipocyte differentiation-dependent gene expression is reﬂected in patterns of fat tissue expression of the same genes among depots [104]. Despite exposure to hormonal manipulations in vivo, such as estrogen treatment, hypophysectomy, or castration in rats, preadipocytes cultured from various depots retain differences in adipogenesis, further implicating inherent differences among depots [133, 134]. To exclude effects of admixture with other cell types in primary preadipocytes in cultures isolated from different regions and to test effects of depot origin on adipogenesis deﬁnitively, preadipocytes cloned from different depots have been studied. The propensities of preadipocytes from different fat depots in primary cultures to switch on the differentiation program are reﬂected in colonies derived from single cells. In response to differentiation-inducing media, the proportion of differentiated cells in colonies derived from single preadipocytes is highest in abdominal subcutaneous clones, intermediate in mesenteric clones, and lowest in omental clones [65]. This was also found after 40 population doublings in strains made from single subcutaneous, mesenteric, and omental preadipocytes by stably expressing hTERT [66]. As in primary cultures, lipid accumulation, aP2 expression, G3PD activity, and C/EBP-a and PPAR-c are highest in subcutaneous, intermediate in mesenteric, and lowest in omental hTERT strains. Not every cell within clones differentiates at the same time. Differentiated cells appear later in visceral than subcutaneous clones. Although adipogenesis is delayed in omental preadipocytes, it does eventually proceed [65]. This is consistent with the

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ﬁnding that PPAR-c mRNA abundance is similar in abdominal subcutaneous and omental whole fat tissue samples [115]. Those cells that acquire lipid inclusions within clones also exhibit C/EBP-a upregulation, irrespective of depot origin. This indicates that regional variation in adipogenesis depends on mechanisms upstream of transcription factor expression, otherwise cells with increased C/EBP-a but no lipid inclusions should be evident. As in human studies, regional differences are evident in colonies arising from single rat cells after weeks in culture [47, 103, 104]. Thus, capacity for adipogenesis depends on depot origin in preadipocytes cloned from both rats and humans, indicating preadipocytes from different depots are inherently distinct. 2.2.5 Regional Variation in Susceptibility to Apoptosis

More apoptotic fat cells are found in human omental than abdominal subcutaneous fat [112, 135, 136]. Consistent with this, preadipocytes from different depots vary in susceptibility to apoptosis induced by serum deprivation or TNF-a [65, 66, 112, 136]. As in the cases of regional differences in replication and adipogenesis, mesenteric preadipocytes are intermediate between subcutaneous and omental cells. Furthermore, regional differences in susceptibility to TNF-a-induced apoptosis are retained in hTERT-expressing strains after 40 population doublings. Thus, differences in preadipocyte replication, adipogenesis, and apoptosis among depots appear to be inherent. 2.2.6 Differences in Preadipocyte Subpopulations among Fat Depots

Two preadipocyte subtypes, one capable of more extensive replication, differentiation, and adipogenic transcription factor expression and less apoptosis in response to TNF-a than the other, are present in fat tissue of humans as well as rodents [110, 113, 137]. Both subtypes are found in hTERT strains derived from single preadipocytes, conﬁrming both are of preadipocyte lineage. After subcloning cells of either subtype, both subtypes are found, indicating that switching can occur between them. The subtype with the lower capacities for replication, lipid accumulation, and C/EBPa expression, and greatest susceptibility to TNF-a-induced apoptosis is most abundant in omental preadipocyte populations. Thus, differences among depots in relative abundance of these subtypes could contribute to regional variation in replication, differentiation, and apoptosis. Characteristics of the subtypes also vary among depots. The omental rapidly and slowly replicating subtypes both accumulate less lipid than their subcutaneous counterparts. This implies that the subtypes from different depots are themselves distinct, that developmental imprinting of the cells occurs related to their microenvironment in vivo, or a combination of these mechanisms. Having a preadipocyte subtype that responds poorly to inducers of adipogenesis, yet can switch into a subtype that responds well, may ensure continued presence of

2.2 Physiology

the progenitor pool. If such a mechanism did not exist, all preadipocytes could be induced to differentiate into fat cells at the same time if insulin/insulin-like growth factor (IGF)-1 levels were high and other proadipogenic conditions existed. The presence of both subtypes could also serve to permit plasticity of the progenitor pool over time through selection for clones with particular properties. For example, inﬂammatory cytokine exposure could favor selection of the rapidly over the slowly replicating preadipocyte subtype. This could have long-term consequences for fat depot cellular composition and function. Additionally, the preadipocyte subtypes may develop into differentiated fat cells with distinct properties. In this regard, two populations of fat cells, one large and the other small, have been found in human fat and in fat-speciﬁc insulin receptor knockout mice [138–142]. 2.2.7 Differences in Preadipocyte Gene Expression Profiles among Depots

The large differences in preadipocyte cell dynamic properties (replication, adipogenesis, apoptosis, and subtype abundance) suggest that expression proﬁles of undifferentiated preadipocytes from different fat depots should be distinct. Indeed, genome-wide expression proﬁles of primary preadipocytes cultured in parallel from different depots of rats and humans are distinct [64, 143]. Over 500 genes vary signiﬁcantly among human abdominal subcutaneous, mesenteric, and omental fat depots by more than 2-fold. Interestingly, mesenteric preadipocytes have expression patterns distinct from omental cells, indicating that visceral fat is not homogeneous. Genes involved in developmental processes constitute approximately 25% of the transcripts whose expression varies among depots in both humans and rodents [64, 143, 144], with this gene category being the one that differs most among depots. Mesenteric undifferentiated preadipocytes are distinct from omental cells with respect to developmental gene expression [64]. Depot-speciﬁc developmental gene expression proﬁles persist for 40 population doublings in hTERT strains, indicating developmental gene signatures are inherent. An example of a developmental gene distinctly expressed in omental and subcutaneous preadipocytes is pregnancy associated plasma protein A (PAPPA) [145]. PAPPA is a potent inhibitor of IGF-binding protein 4, which negatively regulates IGF-I activity [146]. Since omental cells are resistant to insulin and IGF-1, they may require more IGF-1 to survive than subcutaneous cells, and PAPPA may facilitate increased local IGF-1 bioavailability. Preadipocytes isolated from different fat depots also differ with respect to developmental genes belonging to the homeobox superfamily [145]. In particular, engrailed-1 (En-1) and members of the homeobox (Hox) clusters are differentially expressed in preadipocytes [145] as well as abdominal subcutaneous and visceral whole fat tissue [147, 148]. En-1 might be expressed within and up to the anatomic boundaries of a fat depot effectively demarcating it, much as this gene demarcates the limits of metameres in Drosophila melanogaster [149]. Hox genes are master regulators of developmental processes controlling the diversiﬁcation of segments along the anteroposterior axis of animals [150]. During

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embryogenesis, groups of cells become committed to form body structures, including limbs and organs [151]. Evidence is accumulating that the combination of Hox genes expressed within functional domains along the anteroposterior axis speciﬁes development of structures within these domains [152]. In mammals, Hox genes are organized into four genomic clusters (A, B, C and D), each containing nine to 11 genes. Adjacent Hox genes are redundant in terms of function, as are paralogous Hox genes (i.e., HoxA9 and HoxC9) [153]. Hox genes that are more 30 in the genomic layout control more anterior structures [154]. Although Hox genes are believed to play a pivotal role in embryogenesis, their expression persists in adult tissue. A particular combination of Hox genes is expressed in every cell. This combination is referred to as the Hox code and is believed to be tightly associated with the positional identity of structures [155]. This cellular identity expressed through the Hox code is subject to regulation by an elaborate cellular memory mechanism [156]. The cellular memory that maintains inherent Hox gene expression is related to polycomb group (PcG) proteins. The later are subdivided into 2 groups: (1) polycomb (PcG), which facilitates stable repression of Hox gene expression, and (2) trithorax G (TrxG), which is responsible for continued activation of Hox genes through many cell generations [157]. PcG proteins are believed to control Hox gene expression by epigenetic processes, such as DNA and histone methylation [158]. Hox gene activation or repression coincides with a speciﬁc pattern of histone H3 lysine di- and trimethylation [159–161]. Those histone modiﬁcations are unique in that they span through large areas rather than being punctate, encompassing several adjacent Hox genes [159]. In particular, lysine 4 dimethylation of histone 3 (H3 K4) is associated with transcriptional activation, possibly by recruiting TrxG activating complexes [162]. H3 K27 trimethylation is believed to be repressive, possibly by recruiting PcG repressive complexes [163]. In addition, histone methylating is also regulated through the action of the histone de-methylation enzymes that ﬁne-tune histone methylation [164]. Thus, Hox gene expression seems to be tightly regulated by histone methylation, histone demethylation, and recruitment of effector proteins (PcG) to the promoters of Hox genes. The function of Hox genes in fat tissue is still unknown. Several studies have suggested that Hox genes might be involved in differentiation and replication of other types of adult cells [165, 166]. For example, in hematopoietic progenitor cells, with which preadipocytes share a common ancestor, Hox genes are believed to regulate progression through progenitor stages by effects on replication and differentiation [165]. These two processes are of pivotal importance in fat tissue function and maintenance. Hox genes can respond to sex steroids [167] and retinoids [168], which are important for fat tissue function and distribution. Hox genes comprise a network of genes whose expression is maintained epigenetically through many cell generations and is pivotal for coordinated development. Whether these genes represent the master developmental regulators in fat tissue that shape distinct fat depots, are a constellation of molecular markers that identify depot-speciﬁc adipocyte lineages, or are imprinted during early development or later by the fat tissue microenvironment are issues that need scrutiny.

2.3 Conclusions

2.3 Conclusions

Fat is not a homogeneous organ. Depot-dependent differences among the preadipocytes from which new fat cells arise appear to be inherent. Regional variations in capacities for replication, adipogenesis, apoptosis, subtype abundance, and expression proﬁles persist for many cell generations in identical culture conditions in strains derived from single preadipocytes. This effectively excludes the possibilities that contamination of cultures with other cell types or effects of hormones, circulation, innervation, or other inﬂuences extrinsic to preadipocytes fully account for the regional differences in preadipocyte characteristics. Differences are found in rats as well as humans, suggesting that particular characteristics of subjects from whom preadipocytes were isolated do not account for regional differences. Differences in transcription factor expression appear to contribute to regional differences in adipogenesis. Together with the ﬁnding that multiple cell dynamic processes vary among depots (replication, adipogenesis, and apoptosis), as does developmental gene expression, it appears that upstream, possibly epigenetic mechanisms contribute to fat cell progenitors from different regions being effectively distinct cell subtypes. These inherent mechanisms, combined with local variation in fat depot cellular composition, circulation, neural, and other factors, probably account for regional differences in fat tissue function. Gender, obesity, or other factors likely further impact the cellular composition and paracrine microenvironment of adipose tissue [169–173] and inﬂuence variation in function among depots. Thus, human fat cell progenitors from different regions are distinct, consistent with different fat depots being separate mini-organs. Perhaps regional heterogeneity in other mesenchymal tissues, such as bone or muscle, is also related to local differences in the inherent properties of the progenitors from which they arise. Interestingly, visceral fat is not functionally homogeneous. Preadipocytes from one of the two main visceral depots, mesenteric fat, have preadipocyte cell dynamic characteristics and expression proﬁles distinct from omental preadipocytes, the other main visceral depot. Differences in lipid synthesis and lipolysis between omental and mesenteric fat have been found, at least in obese women [170, 171]. Mesenteric cannot be readily distinguished from omental fat by computed tomography, magnetic resonance imaging, or other clinical imaging methods. These ﬁndings raise the possibility that the distribution of visceral fat between the mesenteric and omental depots may have as yet unexplained clinical implications. Remarkably, developmental gene products are prominent among transcripts that exhibit regional variation among undifferentiated preadipocytes. Developmental genes may contribute to or be a marker of regional differences in fat tissue function. Differences in developmental gene proﬁles could be set during early development. Alternatively, expression of some of these developmental genes might be modiﬁed in adulthood or imprinted by the local microenvironment. What could set the chain of events in motion that results in interdepot variation in preadipocyte replication, adipogenesis, and susceptibility to apoptosis, subtype

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abundance, and developmental gene expression proﬁles? Ultimately, epigenetic mechanisms, such as cytosine methylation, histone modiﬁcations (e.g., binding patterns, acetylation, and methylation), gene-silencing proteins, or other mechanisms that can be passed on to daughter cells, may be responsible for the differing characteristics of fat cells despite sharing the same genomic DNA. These fundamental processes remain to be explored. The interaction between subject characteristics, particularly BMI and aging, with regional differences in preadipocyte function needs to be examined. Since preadipocyte capacities for replication, adipogenesis, apoptosis, and gene expression proﬁles are affected by aging and obesity [38, 103, 174–181], such interactions are a strong possibility. Another interesting question is whether the pattern of regional variation in adipogenesis or replication of preadipocytes from subjects with peripheral obesity is distinct from that in subjects with visceral obesity. Few interventions are currently available for targeting speciﬁc fat depots. Liposuction and cosmetic surgery can be used to reduce areas of subcutaneous fat but might cause an increase in visceral fat [48–51]. Surgical approaches are not routinely available to reduce visceral fat. It may be possible to develop antiobesity drugs with region-speciﬁc effects since existing agents can alter fat distribution. For example, androgens reduce visceral fat in hypogonadal males, but increase it in premenopausal females. Reducing circulating glucocorticoids decreases visceral fat. Serotoninergic antidepressants, which appear from preliminary data to decrease abdominal fat mass, may do so by affecting hypothalamic–pituitary–adrenal axis activity [182]. Drugs that alter fat tissue cytokine production or sympathetic nervous system effects on fat tissue might improve regional fat distribution. Another potential approach may be to alter activity of adipogenic regulators and so change expression of their many downstream targets. Indeed, TZDs induce a relative increase in subcutaneous fat. If these approaches can be used to convert visceral fat cells into cells with properties more closely resembling those of subcutaneous cells, risks associated with central obesity may be reduced. It could even be possible to decrease visceral fat mass selectively. To aid in the rational development of such drugs, mechanisms responsible for regional obesity need to be understood more completely. A better understanding of these mechanisms may also point to new ways to treat cardiovascular and metabolic disease. Acknowledgments

The authors are grateful for administrative support by J. Armstrong. This work was funded by NIH grants AG13925, DK56891, and AG23960.

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Erdjument-Bromage, H., Tempst, P., Jones, R.S., and Zhang, Y. (2002) Role of histone H3 lysine 27 methylation in Polycomb-group silencing. Science, 298, 1039–1043. Christensen, J., Agger, K., Cloos, P.A., Pasini, D., Rose, S., Sennels, L., Rappsilber, J., Hansen, K.H., Salcini, A.E., and Helin, K. (2007) RBP2 belongs to a family of demethylases, speciﬁc for tri-and dimethylated lysine 4 on histone 3. Cell, 128, 1063–1076. Beslu, N., Krosl, J., Laurin, M., Mayotte, N., Humphries, K.R., and Sauvageau, G. (2004) Molecular interactions involved in HOXB4-induced activation of HSC selfrenewal. Blood, 104, 2307–2314. Lessard, J., Schumacher, A., Thorsteinsdottir, U., van Lohuizen, M., Magnuson, T., and Sauvageau, G. (1999) Functional antagonism of the PolycombGroup genes eed and Bmi1 in hemopoietic cell proliferation. Genes Dev., 13, 2691–2703. Akbas, G.E. and Taylor, H.S. (2004) HOXC and HOXD gene expression in human endometrium: lack of redundancy with HOXA paralogs. Biol. Reprod, 70, 39–45. Kessel, M. and Gruss, P. (1991) Homeotic transformations of murine vertebrae and concomitant alteration of Hox codes induced by retinoic acid. Cell, 67, 89–104. Fried, S. and Kral, J. (1987) Sex differences in regional distribution of fat cell size and lipoprotein lipase activity in morbidly obese patients. Int. J. Obes., 11, 129–140. Fried, S.K., Leibel, R.L., Edens, N., and Kral, J.G. (1993) Lipolysis in intraabdominal adipose tissues of obese men and women. Obes. Res., 1, 443–448. Edens, N.K., Fried, S.K., Kral, J.G., Hirsch, J., and Leibel, R.L. (1993) In vitro lipid synthesis in human adipose tissue from three abdominal sites. Am. J. Physiol. Endocrinol. Metabol., 265, E374–E379. Weisberg, S.P., McCann, D., Desai, M., Rosenbaum, M., Leibel, R.L., and Ferrante, A.W. (2003) Obesity is associated with macrophage

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accumulation in adipose tissue. J. Clin. Invest., 112, 1796–1808. Xu, H., Barnes, G.T., Yang, Q., Tna, G., Yang, D., Chou, C.J., Sole, J., Nichols, A., Ross, J.S., Tartaglia, L.A., and Chen, H. (2003) Chronic inﬂammation in fat plays a crucial role in the development of obesity-related insulin resistance. J. Clin. Invest., 112, 1821–1830. Cartwright, M.J., Tchkonia, T., and Kirkland, J.L. (2007) Aging in adipocytes: potential impact of inherent, depotspeciﬁc mechanisms. Exp. Gerontol., 42, 463–471. Nair, S., Lee, Y.H., Rousseau, E., Cam, M., Tataranni, P.A., Baier, L.J., Bogardus, C., and Permana, P.A. (2005) Increased expression of inﬂammation-related genes in cultured preadipocytes/stromal vascular cells from obese compared with non-obese Pima Indians. Diabetologia, 48, 1784–1788. Dubois, S.G., Heilbronn, L.K., Smith, S.R., Albu, J.B., Kelley, D.E., and Ravussin, E. (2006) Decreased expression of adipogenic genes in obese subjects with type 2 diabetes. Obesity, 14, 1543–1552. Shillabeer, G., Forden, J.M., Russell, J.C., and Lau, D.C. (1990) Paradoxically slow preadipocyte replication and differentiation in corpulent rats. Am. J. Physiol. Endocrinol. Metab., 258, E368–E376. Permana, P.A., Nair, S., Lee, YH., LuczyBachman, G., Vozarova de Courten, B., and Tataranni, P.A. (2004) Subcutaneous abdominal preadipocyte differentiation in vitro inversely correlates with central obesity. Am. J. Physiol. Endocrinol. Metab., 286, E958–E962. Turkenkopf, I.J., Chow, G., East-Palmer, J., Greenwood, M.R., and Johnson, P.R. (1988) Regional and genotypic differences in stromal-vascular cells from obese and lean Zucker rats. Int. J. Obes. Relat. Metab. Disord., 12, 515–524. Karagiannides, I., Tchkonia, T., Dobson, D.E., Steppan, C.M., Cummins, P., Chan, G., Salvatori, K., HadzopoulouCladaras, M., and Kirkland, J.L. (2001) Altered expression of C/EBP family members results in decreased

References adipogenesis with aging. Am. J. Physiol. Regul. Integr. Comp. Physiol., 280, R1772–R1780. 181 Karagiannides, I., Thomou, T., Tchkonia, T., Pirtskhalava, T., Kypreos, K.E., Cartwright, A., Dalagiorgou, G., Lash, T.L., Farmer, S.R., Timchenko, N.A., and Kirkland, J.L. (2006) Increased CUG

triplet repeat binding protein-1 predisposes to impaired adipogenesis with aging. J. Biol. Chem., 281, 23025–23033. 182 Rosmond, R. and Bj€ orntorp, P. (2000) The role of antidepressants in the treatment of abdominal obesity. Med. Hypotheses, 54, 990–994.

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3 Plasticity of the Adipose Organ Saverio Cinti 3.1 Introduction

For many years our group has been developing a new anatomic-functional concept –: the adipose organ. The innovative part of this concept tries to give an explanation to the fact that the two well-known and distinct adipose tissues (white adipose tissue (WAT) and brown adipose tissue (BAT)) are contained mixed together into a dissectible organ formed by several subcutaneous and visceral depots. It was generally believed that the two tissues are separated entities, with different functional activities, contained in different anatomical sites, but our studies demonstrate that anatomical sites exclusive for WAT or BAT do not exist. In fact, many subcutaneous and visceral depots in small mammals and some depots in humans are composed of a mixed tissue made up of white and brown adipocytes. The reason why these two different tissues are mixed together is not clear, but the problem is underestimated because it is easy to accepted that cells both denominated as adipocytes, in spite the fact that their morphology and physiology is completely different, are mixed together in the same organ. We offer an explanation for this anatomical organization – the plasticity of the adipose organ. In other words, the well-known phenomenon of reversible transformation of the adipose tissues due to physiological stimuli such as exposure to cold (the organ phenotype change from white to brown) or to the hormonal environment during pregnancy and lactation (part of the organ phenotype change from WAT to milk-secreting glands) induce a transformation of the phenotype of the adipose organ based on the intrinsic biologic properties of adipocytes that are able to undertake a reversible transdifferentiation. This concept opens a new perspective in the ﬁeld of cell biology – the ability of differentiated cells of mammals to undergo a physiologic reversible process that drives into a new phenotype (with new morphology and physiology). In this chapter, we describe the most important data obtained in our and others laboratories to support this concept of the adipose organ with particular emphasis on data supporting the transdifferentiation as the main process at the base of its property of plasticity.

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3.2 Gross Anatomy Demonstrates that WAT and BAT are Mixed Together in the Adipose Organ

The adipose organ is a multidepot organ (Figure 3.1) [1–3]. In rodents, there are two main subcutaneous depots (anterior and posterior) and several visceral depots (mediastinic, omental, mesenteric, perirenal, retroperitoneal, perigonadic, and perivescical). Discrete subcutaneous depots are also dissectable at the level of the major joints in the limbs. The color of the organ is white and brown – the white parts are made mainly by white adipocytes; the brown parts are made mainly by brown adipocytes. The relative amounts of white and brown parts are genetically determined and depend on several factors (mainly age, sex, environmental temperature, and nutritional status). In most small rodents brown areas are visually evident in the interscapular, axillary, and cervical parts of the anterior subcutaneous depots, and in the mediastinic and perirenal visceral depots. In other words, it is anatomically evident that some subcutaneous (anterior) and visceral (mediastinic and perirenal) depots are composed in part by WAT and in part by BAT.

Figure 3.1 Gross anatomy of the adipose organ of adult female 129Sv mice. The subcutaneous and visceral depots were dissected and positioned on a template of the mice to show their location in the organism. The mouse on the left was maintained under warm conditions (28 C for 10 days) and the mouse on the right under cold conditions (6 C for 10 days). Note the visually evident transformation of the color of the organ due to the increase of BAT and decrease of WAT contained in the

organ. The organ is made up of two subcutaneous depots: A ¼ anterior (deep cervical, superficial cervical, interscapular, subscapular, axillothoracic) and F ¼ posterior (dorso-lumbar, inguinal, gluteal) and of several visceral depots: B ¼ mediastinal, C ¼ mesenteric, D ¼ retroperitoneal, and E ¼ abdomino-pelvic (perirenal, periovarian, parametrial, perivesical). Bar ¼ 1 cm. Reproduced from [4] with permission.

3.3 Light and Electron Microscopy show that White and Brown Adipocytes

In a recent paper, we described quantitative data of the anatomy of the adipose organ of Sv129 adult female mice. We calculated the total number of white and brown adipocytes contained in most depots (anterior subcutaneous, posterior subcutaneous, mediastinic, perirenal, perigonadic, perivescical, retroperitoneal, and mesenteric). Of note, we observed that perirenal, perigonadic, and perivescical depots that are usually described as distinct depots have a unique anatomy, and therefore we denominated this depot abdomino-pelvic. Our data show that, in this strain, all subcutaneous and all visceral depots contain both white and brown adipocytes mixed together. In some depots white adipocytes are more numerous than brown (posterior subcutaneous, mesenteric, and retroperitoneal); in other depots brown adipocytes are more numerous than white (anterior subcutaneous, mediastinic, and abdominopelvic). Interestingly, in these mice, 60% of the total amount of adipocytes of the adipose organ are brown [4].

3.3 Light and Electron Microscopy show that White and Brown Adipocytes have a Well-Defined and Distinct Morphology 3.3.1 WAT

These spherical cells (Figure 3.2) have a variable diameter depending on the size of the cytoplasmic lipid droplet. White adipocytes with a complete set of organelles, considered to be characteristic for differentiated cells, can be very small (less than 10 mm in diameter) in comparison with the average diameter of the adipocytes found in the different depots of adult mammals. Small adipocytes are often found in the adipose organ of young animals. In physiological conditions, the largest adipocytes

Figure 3.2 Scanning electron microscopy of BAT (a) and WAT (b). Brown adipocytes are polyhedral with a multilocular lipid depot. White adipocytes are spherical with a unilocular lipid depot. Bar ¼ left 40 mm; right 80 mm. Used with permission of The American Physiological Society.

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Figure 3.3 Mouse subcutaneous WAT. Part of a white adipocyte is visible. Mitochondria (M) are small, elongated with randomly oriented cristae. BM ¼ basal membrane; L ¼ lipid droplet. Transmission electron microscopy. Bar ¼ 1.4 mm.

(diameter of about 90–100 mm) are found in epididymal fat of old animals. Human adipocytes are identical to those of rodents, but the size is 20–30% larger. The lipid vacuole is in direct contact with the cytosol (i.e., without any interposed membranous structure). The nucleus is compressed by the lipid droplet, but the cytoplasm near it is usually thicker than in the rest of the cell. Here, elongated mitochondria (Figure 3.3), Golgi complex, rough and smooth endoplasmic reticulum, vesicles, and other organelles are usually visible by transmission electron microscopy. Many pinocytotic vesicles are present in the proximity of the plasma membrane and an external lamina surrounds the cell. In general, small adipocytes have more numerous mitochondria than large adipocytes. 3.3.2 BAT

Brown adipocytes store triglycerides as numerous small vacuoles (multilocular cell) (Figure 3.2). The shape of brown adipocytes is polygonal or ellipsoid with a maximum diameter between a minimum of 15–20 mm and a maximum of 40–50 mm. The nucleus is usually central and the cytoplasm is rich in organelles. The most characteristic organelles are the mitochondria. They are numerous, big, and rich in transverse cristae (Figure 3.4). Peroxisomes, Golgi complex, rough and smooth endoplasmic reticulum, vesicles, and other organelles are also visible by transmission electron microscopy. Pinocytotic vesicles and external lamina are also present in this cell. Brown adipocytes are joined by gap junctions [5]. Over the last 27 years in which we have studied the adipose tissues, when an adipocyte with the light microscopic appearance of a multilocular cell was examined under the electron microscope, it always exhibited mitochondria that were characteristic and distinctive of brown adipocytes [3, 6]. This was independent of its

3.4 WAT and BAT have a Different Vascular and Nerve Supply

Figure 3.4 Rat interscapular BAT. Numerous brown adipocyte mitochondria packed with transverse cristae are visible. Three small lipid droplets (L) are also visible. Transmission electron microscopy. Bar ¼ 1.1 mm.

uncoupling protein (UCP)-1 expression. At variance with the researchers who hold that brown adipocytes are cells expressing UCP-1, we believe that expression of UCP1 merely reﬂects the cells thermogenic activity, and that a brown adipocyte is a cell with a distinctive morphology, a multilocular lipid content, and characteristic mitochondria. Moreover, the brown adipocytes of UCP-1 knockout mice are multilocular, while the morphology of their mitochondria is indistinguishable from that of white adipocytes (our unpublished data in collaboration with Barbara Cannon and Jan Nedergaard). In addition, the white adipocytes of aP2-UCP-1 transgenic mice, which express UCP-1 also in white adipocytes, are unilocular and their mitochondria are similar to those of brown adipocytes [7]. Multilocularity is widely considered to be a characteristic feature of developing white adipocytes (preadipocytes). In fact, this is true only in the ﬁrst weeks of postnatal life, when the adipose organ is developing. In this speciﬁc situation there are some multilocular preadipocytes (committed to become white unilocular adipocytes), but they are small and easily distinguished from mature adipocytes [8–10]. For all these reasons, we believe that the multilocular adipocytes found in the adipose organ of adult animals must be considered thermogenically hypo functioning brown adipocytes when they are UCP-1-negative and thermogenically active when they are UCP-1-positive. In this chapter we use a nomenclature in accordance with this deﬁnition of brown adipocyte. 3.4 WAT and BAT have a Different Vascular and Nerve Supply

The adipose organ is diffuse into the organism and most of its depots receive vascular supply by regional visceral or parietal nerve-vascular bundles. Speciﬁc bundles are

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present in the two main subcutaneous depots (murine adipose organ). The extension of the capillary network is quite different in the white and brown parts of the organ. In the brown areas the density of the capillaries is much higher than in the white areas. This may be due to the fact that since brown adipocytes are highly metabolic, they need rapid oxygen supply. Furthermore, a rapid heat dissipation is needed in order to avoid any heat-dependent cellular damage. The nerve supply to the adipose organ is different in the brown and white areas. The former are more innervated than the latter. In brown areas, numerous noradrenergic ﬁbers are also found in fat lobules (parenchymal nerves), running with blood vessels (until the level of capillaries) and directly in contact with adipocytes. Adrenergic receptors (a1,2 and b1–3) are present in the adipose organ and all badrenoceptors are present on adipocytes. The density of parenchymal ﬁbers varies according to the functional status of the organ. During cold exposure the noradrenergic parenchymal ﬁbers increase their density in the brown part of the organ [11]. During fasting, the noradrenergic parenchymal ﬁbers increase their density in the white part of the organ [12]. Vascular noradrenergic ﬁbers are also immunoreactive for neuropeptide Y. The vast majority of these nerves also contain noradrenaline [12, 13], suggesting that they belong to the sympathetic nerve supply to WAT blood vessels. Brown and white areas also have a provision of sensory nerves [14] that are capsaicin-sensitive, and are immunoreactive for calcitonin gene-related peptide and substance P. The functional signiﬁcance of these sensory nerves is not precisely known, although in the rat periovarian adipose depot they affect the recruitment of visceral brown adipocytes during cold acclimation [15]. Recently, a parasympathetic innervation of white fat with possible functional implications has been described, but the matter is still debated [16, 17].

3.5 WAT and BAT have a Different Physiology

White adipocytes have the main purpose to accumulate and release highly energetic molecules (fatty acids) that can supply fuel to the organism during intervals between meals. When the interval prolongs over the weeks, WAT represents the survival tissue. Brown adipocytes use the same molecules to produce heat (nonshivering thermogenesis). This function is due to a protein, UCP-1, exclusively present in mitochondria of brown adipocytes [18–23]. The signal for brown adipocyte activation is a temperature below thermoneutrality (34 C for mice, 28 C for rats, and 20–22 C for humans) that induces activation of the sympathetic nervous system. These neurons of the sympathetic chain directly reach brown adipocytes in the adipose organ [19]. The total volume of the adipose organ is dependent on the equilibrium between WAT and BAT activities because the main cytoplasmic structure (lipid droplets) can be dissipated (BAT) or accumulated (WAT). Of note, genetic ablation of BAT or of all b-adrenergic receptors induces obesity in mice [24, 25], although mice lacking UCP-1 are cold-sensitive, but not

3.6 Phenotype of the Adipose Organ is Variable: Plasticity of the Adipose Organ

obese [26]. On the other hand, ectopic expression of UCP-1 in WAT results in obesity resistance [27]. Accordingly, it has been shown recently that obesity-prone mice have less BAT than obesity-resistant mice [28]. In 1994, another primary function of white adipocytes was discovered – production of a hormone able to inﬂuence animal behavior concerning food intake [29]. This hormone, denominated leptin, also induces energy dispersion (via BAT and locomotor activation) and has gonadotrophic properties. Brown adipocytes in their classic multilocular conﬁguration (i.e., during thermogenic activity) are not immunoreactive for leptin [30, 31]. A growing body of evidence suggests that the adipose organ produces several factors known as adipokines, controlling several important functions such as glucose and lipid metabolism, blood coagulation, blood pressure, and steroid hormones modulation (see other chapters of this book). The production of all these adipokines raised the recent concept of the adipose organ as an endocrine organ [32, 33]. It must be outlined that adipocytes are not the only cell type present in the adipose organ. It has been calculated that only about 50% of its cell content are adipocytes [34]. Vascular elements, preadipocytes, ﬁbroblasts, mast cells, macrophages, nervous elements, and mesenchymal cells with unknown functions are usually found in all depots of the adipose organ.

3.6 Phenotype of the Adipose Organ is Variable: Plasticity of the Adipose Organ

It is well known that the anatomy of the adipose organ is variable. There are two physiologic conditions in which this phenomenon is quite evident – exposure (or acclimatation) to different temperatures and pregnancy/lactation. 3.6.1 Transformation of the Phenotype: Cold and Warm Exposure and Acclimatization

If we observe the adipose organ of cold-exposed mice we see that the color of the organ is changed, suggesting a change into a more brown phenotype (Figure 3.1). This wellknown reversible phenomenon is due to an increase in the number of brown adipocytes, capillaries, and noradrenergic parenchymal ﬁbers of the adipose organ. The same phenomenon can be achieved by the administration of b3-adrenoceptor agonists and is suppressed in mice lacking b3-adrenoceptors [35, 36], suggesting that noradrenergic ﬁbers play a central role mainly acting on b3-adrenoceptors of adipocytes. Accordingly, after cold exposure, the density of noradrenergic ﬁbers increase in all parts of the adipose organ [11, 37]. Capillary density increases in close association with the increase in number of brown adipocytes that produce potent vasculotrophic factors such as vascular endothelial growth factor. The newly formed brown adipocytes could, in theory, derive from the development of pre-existing stem cells resident in the tissue, from migrating stem cells, or from the direct transformation of differentiated white adipocytes (transdifferentiation), or from a combination of the three phenomena. We think that reversible transdifferentiation is the

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most important phenomenon underling this plasticity of the adipose organ for the following reasons: 1)

After cold acclimation a number of brown adipocytes appear in WAT that almost correspond to that of disappeared white adipocytes. From our calculations the total number of adipocytes of the adipose organ after 10 days of cold acclimation (Sv129 adult female mice) increases by 11 million (176 versus 187 million). Of note, the total number of brown adipocytes increases by 42 million and the total number of white adipocytes decreases by 31 million. These data suggest that, in the absence of histological signs of apoptosis or other forms of degeneration of white adipocytes, most of the newly formed brown adipocytes derive from direct transformation of white into brown adipocytes. The absence of necrosis or apoptosis is in line with the fact that the cold-induced adrenergic stimulus is protective of brown adipocytes [38], which are more subject to apoptosis than white adipocytes [39]. It has been recently reported that apoptosis occurs in white adipocytes after intracerebroventricular injection of leptin [40]; however, cold acclimation reduces leptin expression and leptinemia [41], and therefore leptininduced apoptosis can probably be ruled out in our study. 2) Between 80 and 95% of the newly formed brown adipocytes appearing in WAT after b3-adrenoceptor agonist treatment are bromodeoxyuridine (BrdU)-negative. The retroperitoneum depot of 20-week-old rats (Sprague-Dawley) is almost entirely composed by white adipocytes. After 7 days of treatment with the b3adrenoceptor agonist CL 316,243 about 17% of the adipocytes of this depot are brown adipocytes. Electron microscopy and biochemical studies together with immunohistochemistry demonstrate that these newly formed brown adipocytes are prevalently not thermogenically active (only 8% of them are UCP-1-positive) and show evidence for intense mitochondria biogenesis. Importantly, many adipocytes with intermediate morphology between white and brown adipocytes were found by electron microscopy. BrdU experiments showed that the mitotic index of these adipocytes was very low (5%; i.e., 95% of these adipocytes were BrdU-negative, implying that they have not had a mitotic process) [35]. Of note, recently, Granneman et al. obtained similar results: 80% of the multilocular cells (we do not use the term brown here in respect to the authors deﬁnition) were BrdU-negative [42]. 3) After cold acclimation adipocyte precursors do not increase in WAT in spite of the striking increase of brown adipocytes. Electron microscopy can easily detect adipocyte precursors in the developing adipose organ or in the adipose organ of adrenergic stimulated animals in both WAT and BAT [1]. We found rare adipocyte precursors in WAT of cold-exposed and acclimated animals, but quantitative evaluation demonstrates that their number was similar to that found in room temperature control animals (our unpublished observations) and we were not able to ﬁnd adipocyte precursors in WATof rats treated with the b3-adrenoceptor agonist CL 316,243 [35]. 4) After cold acclimation and b3-adrenoceptor agonist treatment a new form of UCP-1-positive brown adipocytes appear in WAT with intermediate aspects

3.6 Phenotype of the Adipose Organ is Variable: Plasticity of the Adipose Organ

Figure 3.5 Adipocytes with intermediate features between white and brown adipocytes (paucilocular transdifferentiating adipocytes). After cold exposure transdifferentiating paucilocular adipocytes appear in WAT with lipid accumulation similar to white adipocytes

(L) and mitochondria similar to those of brown adipocytes. Note that together with large brown mitochondria (B), some small white mitochondria are also visible (W) in the same cell. CAP ¼ capillary. Transmission electron microscopy. Bar ¼ 1 mm.

between white and brown adipocytes. In a recent study we found that cold acclimatization or treatment with b3 agonist induces the appearance in WAT of numerous UCP-1-positive multilocular adipocytes with intermediate morphology between white and brown adipocytes by electron microscopy. The two main aspects of these cells are the presence of a predominant lipid vacuole in the cytoplasm, and a mitochondria population with different intermediate forms between white and brown mitochondria (Figure 3.5). 5) Isolated brown adipocytes have intermediate aspects between white and brown adipocytes. Early in vitro studies from our laboratory showed that adipocytes developed from the stromal vascular fraction obtained from the interscapular BAT of young rats and from humans have intermediate morphology between white and brown adipocytes, including mitochondria with intermediate features [43, 44]. The addition of noradrenaline to the culture medium induces a more typical morphology of the mitochondria of those adipocytes (rats) [45]. In line with these studies, the genetic removal of all types of b-adrenoceptors in mice induces a transformation of the morphology of interscapular BATadipocytes that became unilocular and express leptin [25]. Again, the mitochondria of those cells have intermediate features between white and brown adipocytes. 6) The morphology and protein expression of brown adipocytes is variable in relationship to the level of noradrenaline in the tissue. In murine interscapular BAT of warm-acclimated animals the noradrenaline level is lower than in the interscapular BAT of cold-acclimated animals and the morphology of brown

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adipocytes varies in these conditions as follows: in cold-acclimated animals brown adipocytes have the classic morphology with multilocular lipid droplets and big mitochondria reach of cristae. These cells express UCP-1 and do not express leptin nor S-100B (a protein expressed in white adipocytes and preadipocytes that could be implicated in the organization of the cytoskeleton). Warm acclimated animals show brown adipocytes with unilocular lipid droplet and mitochondria with intermediate features between those of white and brown adipocytes. These adipocytes express leptin and S-100B, but not UCP-1 [31, 46]. Genetically obese ob/ob and db/db mice have a reduced noradrenaline ﬂux to interscapular BAT and brown adipocytes have the same characteristic of the above-described adipocytes of warm-acclimated mice. The above reported data came mainly from our laboratories, but ﬁt with data obtained by other laboratories showing that after cold acclimation of rodents the DNA does not increase in WAT nor does the number of adipocytes. Furthermore, an antiproliferative effect of the sympathetic nervous system on WAT has been recently shown [47]. White into brown transdifferentiation is also suggested by in vitro studies using primary cultures from human subcutaneous adipose tissue: in these papers, UCP-1 expression was induced by treatment with peroxisome proliferator activated receptor-c (PPAR)-c agonists [48] or PPAR-c coactivator-1 transfection [49]. Altogether these data suggest a possible role for b3-adrenoceptors agonists in the treatment of human obesity and diabetes. Human WAT is immunoreactive to b3-adrenoceptors monoclonal highly speciﬁc antibodies [50], but a b3-adrenoceptors agonist producing curative effects for human obesity has not yet been identiﬁed [51]. 3.6.2 Transformation of the Phenotype: Pregnancy and Lactation

The mammary gland is composed of branched epithelial ducts inﬁltrating subcutaneous adipose tissue and connected to a nipple. In adult female mice three bilateral nipples are connected to epithelial ducts inﬁltrating the whole anterior subcutaneous fat depot of the adipose organ. Two bilateral nipples are connected to epithelial ducts inﬁltrating the whole posterior subcutaneous fat depot of the adipose organ. Therefore, virgin adult (postpuberal) female mice are provided by ﬁve bilateral incomplete mammary glands that are ready to became milk-secreting glands during pregnancy and lactation. The two subcutaneous depots containing the glands differ from that of male mice only for the presence of the above-described branched epithelial ducts. The adipose component of these depots follows the general rules described above for the adipose organ – a mixed composition of white and brown adipocytes (with the relative amount depending mainly on age, strain, and environmental conditions). Of note, adipocytes of the mammary glands express the prolactin receptor [52]. During pregnancy and lactation the anatomy of the mammary glands becomes complete with progressive reduction of adipocytes and the formation of

3.6 Phenotype of the Adipose Organ is Variable: Plasticity of the Adipose Organ

milk-secreting lobuloalveolar epithelial glands. This plastic phenomenon is reversible and at the end of lactation the milk-secreting components of the gland disappear to give room for the reappearing adipocytes with a complete reconstruction of the pregravidic anatomy of the gland. This phenomenon was previously viewed as being due to a hiding among the glands of adipocytes that undergo a delipidation process during pregnancy and a lipid reﬁlling process of adipocytes in the postlactation period. Our recent morphological studies combined with the Crelox fate mapping technique on the mammary gland suggested that adipocytes undergo a reversible adipoepithelial transdifferentiation process during pregnancy and lactation [53]. 3.6.3 Transformation of the Phenotype: Hypertrophy and Hyperplasia (Positive Energy Balance: Overweight and Obesity)

When the energy balance becomes positive, the adipose organ increases its white part. White adipocytes undergo hypertropy followed by hyperplasia. In fact, it has been proposed that adipocytes have a maximum volume and cannot be further expanded. This maximum volume, also referred to as critical cell size, is genetically determined and speciﬁc for each depot [54]. Adipocytes with the critical cell size trigger an increase in cell numbers [55, 56]. Not all depots have the same tendency to hypertrophy and hyperplasia – the former seems to be more characteristic of epididymal and mesenteric depots, the latter of inguinal and perirenal depots [54]. Adipose tissue expresses numerous factors that could be implicated in modulation of adipogenesis: insulin-like growth factor-1, transforming growth factor-b, tumor necrosis factor (TNF)-a, macrophage colony-stimulating factor, angiotensin II, autotaxin-lysophosphatidic acid, leptin, resistin, and so on [57]. Interestingly, it has been shown in mice that obesity induced by a high-fat diet is hypertrophic, while obesity induced by hypothalamic lesions due to administration of monosodium glutamate is hyperplastic [58]. It has been suggested recently that adipocyte precursors can derive from bone marrow [59]. It has been shown that the WAT of obese mice and humans is inﬁltrated by macrophages, and that the level of inﬁltration correlates with body mass index (BMI) and mean size of adipocytes [60–62]. This inﬁltration seems to be an important cause for the insulin resistance associated with obesity. We observed recently that macrophages are mainly located at the level of dead adipocytes in WAT of obese mice, obese humans, and in transgenic mice that are lean but with hypertrophic adipocytes (hormone-sensitive lipase knockout mice) [63]. Also, the brown part of the organ is modiﬁed under this condition of positive energy balance. In obese mice, the rate of apoptosis of brown adipocytes increases and this is strongly attenuated in mice lacking TNF-a receptors [64]. In obese animals the morphology of brown adipocytes gradually changes into a morphology similar to that of white adipocytes, including transformation of the multilocular lipid depot into a unilocular one. This is accompanied by activation of the leptin gene and these cells are leptin-immunoreactive [30, 31].

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3.6.4 Transformation of the Phenotype: Hypoplasia (Negative Energy Balance: Caloric Restriction and Fasting)

The morphology of the adipose organ during fasting is quite characteristic, because a variable amount of slimmed cells are present in WAT. The slimmed cells are barely visible by light microscopy, but they are easily recognized by electron microscopy – they have a speciﬁc ultrastructural morphology: cytoplasmic irregular and thin projections with numerous invaginations rich of pinocytotic vesicles (Figure 3.6). These projections enlarge in the proximity of the nucleus and of the residual lipid droplet. In acute fasting, completely delipidized adipocytes can be found near apparently unaffected unilocular cells. Vasculogenesis and neurogenesis is also found in WAT of fasted animals. Capillaries are often surrounded by the thin cytoplasmic projections of the slimmed adipocytes. Neurogenesis is mainly supported by an increase of noradrenergic ﬁbers [12]. In chronic caloric restriction, the reduction in size of adipocytes is homogeneously distributed [65].

Figure 3.6 White adipocytes of retroperitoneal WAT of an adult rat fasted for 3 days. The adipocyte on the right part of the figure (L) shows a tail of slimmed cytoplasm (arrow). The adipocyte on the bottom part of the figure

exhibits few, small lipid droplets (L) and numerous cytoplasmic introflections (arrowheads). CAP ¼ capillary. Transmission electron microscopy. Bar ¼ 2.3 mm. Reproduced from [1] with permission.

3.7 Adipose Organ of Humans

3.7 Adipose Organ of Humans

The basic concepts of the adipose organ of small mammals reported above are probably applicable also to the adipose organ of humans. In fact, WAT, BAT, and mixed adipose tissues are also present in the adipose organ of humans, with most of the morphological and physiological characteristics described for the murine adipose organ. Although a detailed description of the gross anatomy of the human adipose organ has never been performed, it is well known that it is composed of subcutaneous and visceral depots. In humans, the subcutaneous adipose tissue is in continuity with the dermal adipose tissue (in rodents, dermal adipose tissue is separated from subcutaneous adipose tissue by a smooth muscle layer) and it is not limited to deﬁned areas, but is present as a continuous layer beneath the skin. Mammary and gluteofemoral subcutaneous adipose tissues are more developed in females than in males. Visceral depots correspond to those described above for the rodent adipose organ, but the omental depot is particularly well developed in humans. The weight of the human adipose organ of lean adults is about 8–18% of the body weight in males and 14–28% in females (about 5% in monkeys) [66]. Light and electron microscopy of human adipose tissues is identical to that of murine adipose tissues. Development of the human adipose organ extends for a long period, until puberty, mainly through proliferation [67]. During the ﬁrst year of age there is mainly an increase in size. In line with these data, the number of adipocytes, total fat mass, and the percentage of body fat correlate positively with age in both sexes. Instead, adipocyte size does not seem to positively correlate with age, but it seems to be correlated to the amount of fat mass and percentage in both sexes [67]. In massively obese humans, the adipose organ can increase 4 times and reach 60–70% of body weight [48]. In the case of negative energy balance, the adipose organ reduces its volume and adipocytes reduce their size. The reduction in size of adipocytes is important because the size of adipocytes correlates with insulin sensitivity [68]. Completely delipidized adipocytes can be found in the adipose tissue of subjects with negative energy balance. The morphology of delipidized adipocytes is quite similar to that described above for the equivalent cells of mice or rats. The fate of these delipidized adipocytes is still debated; some authors suggest that they undergo apoptosis. Not all depots react in the same way to negative energy balance. Subcutaneous adipose tissue from the gluteofemoral region of premenopausal women is more resistant to slimming than the subcutaneous abdominal adipose tissue, but after menopause the slimming process is similar. This seems to be due to a combination of increased lipoprotein lipase activity and reduced lipolytic activity in the gluteofemoral adipose tissue. The reduced lipolytic activity seems to be due to a relative preponderance of antilipolytic activity of a2-adrenoceptors over the lipolytic b-adrenoceptors [69]. In general, a2-adrenoceptors are more abundant in human adipose tissue than in murine adipose tissue. In genetically modiﬁed mice lacking b3- and expressing human a2-adrenoceptors, a high-fat diet induces hyperplasia (but not

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hypertrophy) of adipose tissue and mice are insulin-sensitive. These experiments are in line with the importance of a2-adrenoceptors for adipocyte hyperplasia and with the relationship of their size with insulin sensitivity [70]. Like the murine adipose organ, the human adipose organ contains BAT. It is easy to understand that the surface/volume relationship of the human body is quite different from that of small mammals and therefore human thermodispersion is much lower than in rodents. This alone justiﬁes a reduced need for BAT in adult humans. Newborns have a different surface/volume relationship and a considerable amount of BAT is present at that age. Nevertheless, brown adipocytes dispersed among white adipocytes have been described in several histological studies (including studies showing the presence of UCP-1) [71, 72]. BAT in human newborns has been described in almost all the same sites described for rodents, and UCP-1 gene expression was found in biopsies from visceral adipose tissue of adult lean and obese patients. In the same paper, the authors calculated the presence of one brown adipocyte for every 100–200 white adipocytes in the visceral adipose tissue of adult lean humans [73]. BAT has also been described to be increased in outdoor workers in northern Europe [74] and in patients with pheochromocytoma (a noradrenaline-secreting tumor). Furthermore, rare cases of hibernoma (BAT tumors occurring in several anatomical sites, including subcutaneous and visceral fat) have been described (about 100 cases have been described in the literature [75], and we recently observed a case (Figure 3.7) in which brown adipocytes expressed UCP-1 and had the classic electron microscopy with typical mitochondria). Brown adipocytes are able to incorporate high levels of glucose and ﬂuorodeoxyglucose is used in radiology (positron emission tomography (PET)) to identify tissues with a high rate of glucose incorporation such as tumor metastasis. Through

Figure 3.7 Human subcutaneous WAT of the neck. UCP-1 immunohistochemistry. UCP-1immunoreactive multilocular brown adipocytes (brown) are mixed with unstained unilocular

white adipocytes. Light microscopy. (UCP-1 antibody kindly provided by Dr. Daniel Ricquier, Paris.) Bar ¼ 50 mm. Reproduced from FASEB Journal with permission.

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4 Biology of Adipose Tissue Stem Cells Jeffrey M. Gimble, Bruce A. Bunnell, and Farshid Guilak 4.1 Introduction

The broad deﬁnition of a stem cell is a cell that has the ability to self-renew and differentiate into one or more specialized terminally differentiated cell types. It has become evident that stem cells persist in and can be isolated from many adult tissues. Adipose tissue represents an abundant and accessible source of adult stem cells with the ability to differentiate along multiple lineage pathways. This chapter summarizes the origins, isolation, characterization, and differentiation potential of adiposederived stem cells (ASCs) in both in vitro and in vivo models.

4.2 In Situ Localization and Embryology

The topic of adipose tissue embryology has been the subject of a number of excellent and thorough reviews [1, 2]. The embryonic mesoderm gives rise to most, if not all, adipose tissues. The adipose depots ﬁrst appear as condensations of mesenchymal cells that are spindle-like ﬁbroblasts characterized by an abundant endoplasmic reticulum and a high nuclear-to-cytoplasmic ratio [3]. By the second trimester in humans, the fat depots contain preadipocytes distinguished by the paranuclear localization of lipid vacuoles, mitochondria, and other organelles [1, 2]. Eventually, the lipid vacuoles within the cells become spherical and/or multilocular, and are associated with glycogen granules and increased numbers of mitochondria [3]. The ﬁrst mature adipocytes have been reported to lie close to capillaries, and this suggests that a possible relationship exists between pericytes and adipogenic progenitors or stem cells. Isotopic labeling and in vitro studies in rodent models have demonstrated that adipocyte progenitors continue to proliferate throughout life [4–6]. In addition, the adipocytes increase their size when animals are placed on a high-fat diet [7, 8]. Radioisotope turnover studies indicate that rat adipose tissues contain two cell populations – one with a rapid turnover rate measured in days and one with a

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longer half-life of months to years [6]. Consistent with this, in vitro studies using human adipose tissue conﬁrm the presence of at least two populations based on proliferative rates, although the relative abundance of each population varies between adipose depots [9]. Likewise, in vivo studies of human subjects using 2 H2 O mass isoptemer distribution analysis estimate that adipose tissue cells have a half-life of between 8 and 15 months [10], suggesting the need for a local stem cell population to support continual renewal of adipose cells. Nonetheless, there remains room for further investigation. For example, stem cell populations in the hair follicle have been identiﬁed base on their long-term retention of bromodeoxyuridine labeling [11] and such methods should be applied in studies of adipose tissue depots.

4.3 Isolation Methods

The isolation of cells from adipose tissue was pioneered by Rodbell et al. in the mid1960s using the rat as an experimental model [12–15]. These methods were further reﬁned and developed by Ailhaud, Arner, Bjorntorp, Hauner, Smith, and others for use on adipose tissue isolated from human and other species [16–22]. The advent of tumescent liposuction methods [23] further simpliﬁed the isolation procedure by providing a minced tissue homogenate starting material without signiﬁcant damage to the stem cell population [24, 25]. However, it should be noted that ultrasoundassisted liposuction has been found to reduce the yield and viability of the isolated adipose-derived cells [26]. Currently, most laboratories wash the tumescent lipoaspirate tissue or minced resected adipose tissue with a buffered salt solution, digest it in collagenase solution for periods of up to 1 h at 37 C, and collect the preadipocyte stromal cells in a stromal vascular fraction by differential centrifugation. The collagenase should be screened prior to use since this enzyme activity can vary between lots even when purchased from the same manufacturer [27]. At this stage, some laboratories plate the pellet directly to select for a plastic adherent cell population over a 24- to 72-h period [28]. Others laboratories ﬁrst ﬁlter the cell pellet through a 40- to 200-mm nylon mesh before plating. Alternatively, investigators are beginning to use antibody-based methods (ﬂow cytometry, magnetic immunobeads) to select for a stromal vascular fraction cell subpopulation based on surface immunophenotype. There is recent evidence that cells with characteristics similar to those found in the stromal vascular fraction can be recovered directly from lipoaspirate ﬂuid without collagenase digestion [29] and it is possible that more laboratories will employ this modiﬁed isolation method in the future. 4.3.1 Yield, Proliferation Rate, Depot, and Aging Influences

Following collagenase digestion, around 310 000 nucleated cells are recovered per milliliter of lipoaspirate tissue [30]. Of these, around 3% are adherent to plastic in

4.4 Characterization

the presence of 10% fetal bovine serum based on a colony-forming unit-ﬁbroblast assay and yield around 250 000 ASCs within a 6-day culture period [30]. The adherent ASCs proliferate with a doubling time between 3.2 and 4.7 days [30, 31]. The results obtained by others are comparable [26, 29, 32, 33]. According to one report, the cells recovered from the various subcutaneous adipose depots display similar proliferative and immunophenotypic features in vitro [26]. However, there is evidence that human adipose tissue cells from subcutaneous depots are distinct when compared to those from mesenteric or omental sites [9, 34–36]. There have been variable reports concerning the effect of age on the yield of ASCs per unit volume of subcutaneous adipose tissue. While initial reports described a decrease in ASC numbers with advancing age [20], more recent studies from the same investigators and others did not conﬁrm this ﬁnding [37–39]. Indeed, it has been suggested that adipogenesis may be the default differentiation pathway for aging mesenchymal stromal cells and this may account for the accumulation of ectopic adipose depots in older individuals [40].

4.4 Characterization

Multiple criteria have been used to characterize adipose-derived cells, including immunophenotype by cell surface antigens, immunogenicity, proteomic or genomic expression proﬁles, cytokine proﬁle, or clonality. 4.4.1 Immunophenotype

Due to the success in using surface antigens to distinguish between hematopoietic cell populations, a number of investigators have used ﬂow cytometry [30, 33, 41–46]. The immunophenotype of the adipose-derived cells changes as a function of plastic adherence and expansion [30, 45]. Overall, the immunophenotype of passaged ASCs is similar to that of bone marrow-derived mesenchymal stem cells (MSCs). 4.4.2 Immunogenicity

The immunogenic features of adipose-derived cells have been examined as a function of isolation and expansion. While freshly isolated stromal vascular fraction cells initiate a mixed lymphocyte reaction (MLR), plastic adherent ASCs do not [45, 47–50]. Indeed, the passaged ASCs suppress an active MLR between initiated between responder and stimulator peripheral blood monocytes [45, 48, 49]. These immunosuppressive effects appears to be mediated, in part, by secreted factors such as prostaglandin E2 [47]. The immunogenic features of the ASCs are similar to those of bone marrow-derived MSCs [48, 51].

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4.4.3 Proteomic and Transcriptomic Analysis

A limited number of laboratories have used global methodologies to characterize the adipose-derived cells. Mass spectrometry proteomic analyses have identiﬁed around 170 proteins in undifferentiated human ASC protein extracts [52] and around 80 proteins in conditioned medium from human ASCs [53]. Upon adipocyte differentiation, over 40 cellular proteins were induced more than 2-fold while over 10 proteins were reduced more than 3-fold [52]. Overall, the ASC proteome exhibited greater than 55% similarity or identity with the published proteome described of human bone marrow-derived MSCs [52, 54]. However, this may be an underestimate since a direct comparison of human ASC and MSC proteomes reveals that the proteins detected in the two cell populations share greater than 98% identity, with no novel cell-typespeciﬁc proteins being identiﬁed (Izadpanah et al., in preparation). Transcriptomic approaches have been used to characterize undifferentiated human ASCs in relation to their immunophenotype [44]. Comparison of the human ASC transcriptome to that of human MSCs in the undifferentiated [44] and lineage differentiated state (adipocyte, chondrocyte, osteoblast) [55] revealed a high degree of similarity. 4.4.4 Cytokine Profile

The ASCs secrete a number of cytokines based on analyses of conditioned medium of the undifferentiated and adipocyte differentiated cells. As expected, adipogenic conditions induce human ASC expression of adipokines, including leptin, adiponectin, and the serine protease inhibitor, plasminogen activation inhibitor-1 [28, 53]. It is of interest that adipocyte differentiated ASCs release multiple serine protease inhibitors, suggesting that this secreted protein family plays a role in adipocyte function [53]. Likewise, human ASCs express a number of angiogenic cytokines, including vascular endothelial growth factor and hepatocyte growth factor (HGF) [56– 58]. The ASCs induce their expression of HGF following exposure to exogenous basic ﬁbroblast growth factor (bFGF), epidermal growth factor, and ascorbate [59]. In addition, the human ASCs express inﬂammatory and hematopoietic cytokines at a constitutive level that is induced further following exposure to endotoxin or lipopolysaccharide [59]. Overall, the cytokine expression proﬁle of the human ASCs is similar to that reported for human bone marrow-derived MSCs [59]. 4.4.5 Clonality

The hallmarks of a stem cell are the ability to self-replicate and to differentiate along multiple lineage pathways. Several laboratories have used clonal assays to demonstrate that progeny isolated from a single adherent human adipose-derived cell have multipotentiality in vitro [33, 60]. Over 50% of the ring-cloned ASC lines from three different donors differentiated along two or more lineage pathways following

4.5 Differentiation and Potential Utility for Regenerative Medicine

expansion over 22–25 passages [60]; however, the percentage of multipotential clones isolated by limit dilution methods was lower [33]. Nevertheless, these ﬁndings support the stem cell identiﬁcation applied to the culture-expanded adherent adipose-derived cell populations. Further in vivo studies evaluating the ability to transplant multipotential ASCs successively between individual animals will be necessary to prove their stem cell nature.

4.5 Differentiation and Potential Utility for Regenerative Medicine

The abundance and accessibility of human adipose tissue makes ASCs an attractive source of adult stem cells for tissue engineering and regenerative medical applications. Transplanted adult stem cells may improve recovery in damaged tissues through lineage-speciﬁc differentiation and repopulation of the damaged organ, and/or through the release of paracrine factors modulating the regenerative capacity of endogenous cells in the host tissue. In vitro and in vivo studies have shown that ASCs possess broad differentiation ability as summarized below. 4.5.1 Mesodermal Lineages

In response to speciﬁc inductive cocktails of cytokines and chemical agents, human ASCs display biochemical and morphological markers of adipocytes, cardiomyocytes, chondrocytes, endothelial cells, osteoblasts, hematopoietic supporting stromal cells, skeletal myocytes, and smooth muscle cells [28, 32, 33, 56–59, 61–76]. Multiple groups have compared the chondrogenic and osteogenic capacity of ASCs to bone marrow-derived MSCs in parallel assays [55, 77–81]. Some authors have concluded that the ASCs are less robust than MSCs in their ability to differentiate along these pathways [77, 78]; however, it should be noted that the culture conditions utilized in such studies often have been optimized for MSCs rather than ASCs and thus direct comparisons are not particularly meaningful. Indeed, recent studies have found that ASC chondrogenesis is improved markedly upon induction with bone morphogenetic proteins [79, 82, 83], whereas MSCs assume a hypertrophic phenotype under the same conditions. Likewise, the passage number of the ASCs also inﬂuences their chondrogenic potential [84]. Further studies will be necessary to deﬁne the appropriate ASC-speciﬁc growth medium for each lineage pathway. 4.5.2 Endodermal and Ectodermal Lineages

Although adipose tissue has mesodermal origins in the embryo, the ASCs can display biochemical and morphological characteristics of embryonic endodermal and ectodermal cells. In response to oncostatin M, HGF, bFGF, and other factors, ASCs

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express albumin, urea, and a-fetoprotein in a manner similar to hepatocytes [85–87]. Likewise, in response to a cocktail containing activin, extendin, and additional factors, the ASCs express mRNAs encoding insulin, glucagon, and other pancreatic islet cell-associated products [88]. In the presence of retinoids, the ASCs express cytokeratins consistent with epithelial cell differentiation [89]. Multiple groups have demonstrated that ASCs exposed to antioxidants in serum-free conditions assume the morphology and express the proteins (nestin, intermediate ﬁlament M, NeuN) associated with neuronal differentiation [90–97]. Altogether, these ﬁndings support the characterization of ASCs as a pluripotent stem cell population.

4.6 Conclusions

The ASCs meet many of the criteria of an ideal adult stem cell [98]. The source tissue is accessible and yields millions to billions of cells. The ASCs themselves are multipotent, can be transplanted in autologous manner, and can be manufactured according to deﬁned methods consistent with current Good Manufacturing Practices. Nevertheless, several questions remain to be answered. Can allogeneic ASCs be transplanted in a safe and effective manner? Do ASCs have the risk of causing tumors after long-term implantation in animal models? Do ASCs exert some of the regenerative potential at a site of injury through the release of growth factors or their metabolic function? These and other issues will engage investigators in the years to come.

Acknowledgments

The authors acknowledge support from the Pennington Biomedical Research Center Clinical Nutrition Research Unit (P30 DK072476) (J.M.G.), the Pennington Biomedical Research Foundation (J.M.G.), the National Center for Research Resources (B.A.B.), NIH grant RR00164 (B.A.B.), the State of Louisiana Millennium Health Excellence Fund (B.A.B.), the Louisiana Gene Therapy Research Consortium (B.A. B.), NIH grants AR50245 and AG157678 (F.G.), and editorial assistance from Ms. Laura Dallam and Ms. Lori Steib.

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Part Two Metabolic Functions of Adipose Tissue

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5 Molecular Mechanisms of Adipocyte Lipolysis James G. Granneman and Hsiao-Ping H. Moore 5.1 Introduction

The storage and mobilization of lipid are fundamental cellular processes and multicellular organisms, from insects to mammals, have evolved specialized cells that store lipid energy in times of excess for mobilization in times of need. In mammals, adipose tissue functions as a highly specialized lipid energy buffer that stores excess energy as triglyceride for systemic mobilization as free fatty acids (FFAs). Although it is well established that triglyceride stores are dynamically regulated, the cellular and molecular bases of this dynamic regulation are just now being revealed. Recent work indicates that storage and mobilization of intracellular lipid involves the assembly of specialized subcellular structures (i.e., lipid droplets), the targeting of unique protein scaffolds and enzymes, and the dynamic trafﬁcking of lipases and regulatory proteins. The signiﬁcance of these interactions is not well understood. However, adipocytes are a key source of fatty acids during fasting, exercise, and in insulin-resistant states, and alterations in adipose tissue lipolysis can have an important impact on energy balance and insulin sensitivity. It is likely a better understanding of the new lipid droplet biology and the molecular mechanisms of lipolysis will lead to new therapeutic approaches to obesity and diabetes.

5.2 Key Players in Adipocyte Lipolysis

Adipocyte lipolysis is a multifaceted phenomenon that is subject to distinct temporal controls, many of which are poorly understood. This chapter focuses on protein trafﬁcking during rapid, hormone-stimulated lipolysis (for recent reviews on the general regulation of lipolysis, see [1, 2]). Interest in lipolytic protein trafﬁcking stems from the seminal work of Londos et al. (reviewed in [3, 4]). These investigators demonstrated that hormone-stimulated lipolysis is directly related to the level of protein kinase A (PKA) activation [5, 6]. Thus, major signals that acutely inhibit

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Figure 5.1 Rapid regulation of adipocyte lipolysis of PKA. Adipocyte lipolysis is regulated by diverse stimulatory and inhibitory ligands acting through G-protein coupled receptors (GPCR) that transverse the plasma membrane bilayer. GPCR-generated signals are integrated by adenylyl cyclase, which generates cAMP. In addition, insulin suppresses lipolysis through protein kinase B (PKB)-dependent activation of

phosphodiesterase 3 (PDE3), which degrades cAMP to AMP. Signals controlling lipolysis converge at PKA, which directly phosphorylates HSL and PLINA, the major PKA substrate in fat cells. PLINA is targeted to the surface of lipid droplets that are covered by a phospholipid monolayer. ATGL and its coactivator, CGI-58, are key elements PKA-regulated lipolysis, yet neither appears to be direct targets of PKA.

(e.g., insulin and adenosine) or stimulate (e.g., catecholamines) lipolysis converge at the level of PKA (Figure 5.1). In addition, they discovered perilipin (PLIN) as a major lipid coat protein in adipocytes and demonstrated that this protein is the dominant target of PKA activation [7]. Lastly, they showed that hormone-sensitive lipase (HSL), itself a target of PKA, translocates to lipid droplet surfaces during PKA activation [8]. Together, these observations provided a conceptual framework whereby lipolysis is regulated by the PKA-dependent trafﬁcking of proteins to lipid droplet surfaces. 5.2.1 Lipid Droplets and Droplet Scaffold Proteins

Virtually all cells can accumulate small amounts of neutral lipid in structures termed lipid droplets or lipid bodies. Until relatively recently, these structures were

5.2 Key Players in Adipocyte Lipolysis

considered to be inert repositories of lipid; however, recent series of proteomic and lipidomic experiments have clearly established that lipid droplets are heterogeneous, dynamic organelles involved in the synthesis, movement and degradation of lipid [3, 4, 9–15]. Lipid droplets of adipocytes are thought to form in a specialized subdomain of the endoplasmic reticulum [16], and are sequentially bound by several proteins containing conserved PAT (for PLIN/adipose differentiation-related protein (ADRP)/ Tip-47) domains as they migrate and enlarge [17–19]. The biology of PAT proteins is an active area of research and accumulating evidence suggests that speciﬁc PAT proteins play specialized roles in lipid droplet biology [4, 18–20]. PAT proteins have functionally divided in to those that translocate between the cytoplasm and lipid droplet surface, and those that are constitutively bound to the lipid droplet surface. Exchangeable PATs, or ePATs [18], appear to be involved in the assembly and enlargement of lipid droplets, and include S3-12, Tip-47 and to some extent lipid storage droplet protein-5 (also known as myocardial lipid droplet protein and oxidative tissues-enriched PAT protein). PAT proteins that are constitutively bound to lipid droplets (i.e., cPATs) include ADRP and PLIN. ADRP expression is upregulated during lipid droplet formation and is induced by conditions that expose adipocytes elevated levels of fatty acid [21, 22]; however, mature white adipocytes normally express little if any ADRP in vivo. Both PLIN and ADRP are unstable proteins that are stabilized when targeted to lipid droplets [23, 24]. Recently, two members of the cell death-inducing DNA fragmentation factor Alike effector (CIDE) protein family, CIDEA and CIDEC (also known as fat-speciﬁc protein-27), have been shown to be targeted to adipocyte lipid droplets. These proteins have some sequence homology to PLIN and ADRP, and appear to play a role in the formation and structure of adipocyte lipid droplets [25, 26]. CIDEA is expressed in human white adipose tissue, and CIDEA levels in obesity are correlated with insulin sensitivity and decreased lipolysis [26, 27]. In mice, CIDEA is expressed in brown, not white, fat, and knockout of the CIDEA elevates thermogenesis and brown fat metabolism [28–30]. CIDEC protein is found in white fat cells where its expression promotes triglyceride storage [25, 28, 29]. CIDEC appears to be critical for maintaining the unilocular phenotype of typical white adipocytes [31]. Interestingly, mice lacking Cidec have elevated thermogenesis, expanded mitochondrial biogenesis in white fat, and are resistant to dietinduced obesity [31]. The neutral lipid core of adipocyte lipid droplets is surrounded by a monolayer of phospholipid [32], and it has been proposed that the monolayer phospholipid composition plays a role in the activity of lipolytic effectors [33]. Although the phospholipid composition of the adipocyte lipid droplet monolayer is not known, that of HepG2 cells is unique among intracellular membranes [34]. Recently, Guo et al. [35] conducted a functional genomic screen of genes involved in lipid droplet formation in Drosophila S2 cells. Interestingly, numerous genes of phospholipid metabolism had strong effects on lipid droplet number and size, indicating that the phospholipid composition of the monolayer strongly affects formation and maintenance of lipid droplet structures.

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Of the PAT proteins expressed in adipocytes, there is compelling evidence that PLINA, a PLIN splice variant, plays a central role in orchestrating hormonestimulated lipolysis in adipocytes (discussed in detail below). PLINA is the major lipid droplet PAT protein of fully differentiated adipocytes [17, 19]. PLINA can be phosphorylated at up to six sites by PKA and is the major target of PKA phosphorylation in adipocytes [7]. Recent evidence indicates that PLIN is a multifunctional protein, capable of lowering basal lipolysis, promoting lipolysis after PKA activation, and controlling lipid droplet fragmentation through mechanisms that are lipasedependent and -independent. Basal lipolytic rate is inversely related to PLIN expression [36]. Given its location and effect on basal lipolysis, PLIN was initially hypothesized to form a protective barrier that shielded triglyceride from cellular lipases, and that PKA phosphorylation led to the dissociation of PLIN, loss of the protective barrier, and subsequent attack by intracellular lipases. However, recent data suggest that Plin does not form a continuous barrier at the lipid droplet surface, but rather may provide a scaffold for the targeting and trafﬁcking of lipolytic effectors. For example, proteomic studies demonstrate that lipid droplets contain numerous proteins, including lipases, in addition to PLIN [11, 37]. Furthermore, the abundance of PLIN protein is largely unrelated to fat cell size [38, 39] and high-resolution immunoﬂuorescence analysis indicate a discontinuous pattern of PLIN on the surface of large droplets of mature fat cells [32, 40, 41]. Genetic deletion of Plin in mice largely abrogates hormone-stimulated lipolysis [42–44]. Furthermore, expression of wild-type PlinA, but not phosphorylationdefective mutants, restores PKA-dependent lipolysis in adipocytes differentiated from Plin null mice [43]. Reconstitution experiments in heterologous (non-fat cell) systems demonstrate that PLINA confers the ability of PKA to increase HSLdependent and -independent lipolysis [45–48]. It is important to note that PLIN regulation of lipolysis is complex and involves phosphorylation-dependent and independent interactions among different functional domains of the protein [48]. Recent work has identiﬁed a single PKA site in PLINA, Ser517, that appears to globally regulates PKA-dependent lipolysis in fat cells [49]. 5.2.2 Lipases 5.2.2.1 HSL Until recently, HSL was considered to be the major, if not exclusive, lipase mediating hormone-stimulated lipolysis (reviewed in [50–52]). HSL exhibits strong diglyceride lipase hydrolase activity, and 10- and 5- fold weaker activity against triglyceride and monoglyceride substrates, respectively [53]. Interestingly, phosphorylation of HSL by PKA increases its activity toward triglyceride, but not aqueous esterase substrates [54]. Hormone-stimulated lipolysis (measured by cellular FFA release) is signiﬁcantly impaired, but not eliminated, in HSL knockout mice. Adipocytes of HSL null mice are largely incapable of releasing glycerol and have massive accumulation of cellular

5.2 Key Players in Adipocyte Lipolysis

diacylglycerol – clearly demonstrating the importance of HSL as a diglyceride lipase [55]. 5.2.2.2 Adipose Triglyceride Lipase The fact that loss of HSL does not abolish hormone stimulated lipolysis indicated the existence of another triglyceride lipase in adipose tissue. The identiﬁcation of a second major lipase, adipose triglyceride lipase (ATGL, also known as desnutrin, TTS-2.1; UniGene name PNPLA2 (patatin-like phospholipase domain-containing2)) [56–58] and the discovery that comparative gene identiﬁcation (UniGene name ABHD5 (a/b-hydrolase domain-containing protein-5), a PLIN-interacting protein [59, 60], is a key coactivator of ATGL [61] greatly advanced our understanding of the biochemical basis of adipose tissue lipolysis. ATGL is a member of the patatin-domain containing family of proteins. While most abundant in fat, ATGL is found in numerous tissues [57, 58]. ATGL has strong triglyceride hydrolase activity, but no activity against diglyceride or monoglyceride substrates [58]. Interestingly, ATGL has low, but detectable transacylase and phospholipase activities [56, 62]. It is unclear how these minor activities of ATGL are regulated in adipocytes or if they are involved in hormone-stimulated lipolysis. ATGL is a phosphoprotein; however, it is not a direct PKA target [9, 58]. Genetic disruption of Atgl in mice results in massive lipid accumulation in muscle (cardiac and skeletal), liver, kidney and testes [63]. Atgl deletion reduces isoproterenol-stimulated lipolysis in white fat explants by more than 70%, corresponding to the loss in total triglyceride hydrolase activity. Similar results have been observed using small hairpin RNA knockdown in cultured cells [64, 65]. Atgl null mice have a profound defect in lipid mobilization during fasting and are cold-intolerant. Human mutations of ATGL have been discovered that result in ectopic lipid accumulation and myopathy – demonstrating the importance of this lipase in humans [66–68]. Interestingly, a C-terminal truncation mutation of ATGL that abrogates activity in vivo does not inhibit activity in vitro; rather, the mutation results in mistargeting of the protein away from lipid droplets in vivo [69]. These observations further reinforce the importance of lipase targeting and trafﬁcking in the regulation of lipolysis. 5.2.2.3 CGI-58 CGI-58 (ofﬁcially, a/b-hydrolase domain-containing protein-5) belongs to the esterase/lipase subfamily of proteins containing a/b-hydrolase folds. However, unlike bona ﬁde lipases, the predicted catalytic serine within the consensus GXSXG motif contains asparagine [70], and thus CGI-58 exhibits no lipase activity [61, 71]. Rare homozygous mutations of CGI-58 results in Chanarin–Dorfman syndrome (Mendelian Inheritance in Man #27630) that is characterized by ectopic lipid accumulation in numerous tissues [70, 72–75]. A major advance in understanding the function of CGI-58 came with the discovery that CGI-58 dramatically increases the triglyceride hydrolase activity of ATGL owing to a direct interaction between these proteins [61]. A second key feature of CGI-58, discussed below, is its ability to associate with PAT proteins, in particular PLIN [59, 60, 71]. Very recently, CGI-58 was found to mediate the acylation of lysophosphatidic acid [76]. The signiﬁcance of this activity is presently

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unclear; however, mutations of CGI-58 that result in Chanarin–Dorfman syndrome do not affect this acyltransferase activity. 5.2.2.4 Other Lipases and the Biological Significance of HSL versus ATGL A recent proteomic survey using activity-based probes concluded that ATGL, HSL, and monoglyceride lipase are the major lipases in mouse adipose tissue [77], while combined pharmacological, immunological, and genetic approaches strongly indicate that ATGL and HSL constitute greater than 90% of triglyceride hydrolase activity in fat cell extracts [78–80]. It is important to remember, however, that triglyceride hydrolase activities against artiﬁcial triglyceride emulsions may not reﬂect activity against biological substrates within the cell [69]. ATGL and HSL appear to work in concert during PKA activation because of their complementary enzymatic activities. Moreover, the relative contribution of each of these lipases is unlikely to be ﬁxed, but rather will depend on factors like nutritional status (fasting selectively upregulates ATGL [57]), subcellular targeting of ATGL, and the relative abundance of CGI-58.

5.3 Lipolytic Protein Trafficking 5.3.1 PLIN Subcellular Targeting

As mentioned above, there is strong evidence that lipolysis occurs at lipid droplets, which have a unique protein composition, and that PLIN is indispensable in regulating the access and activity of lipases at these lipid droplet surfaces. Immunoelectron microscopic analysis of adipocytes demonstrated that PLIN is on the surface of lipid droplets [32]. Original immunoﬂuorescence analysis in cultured model adipocytes found PLIN to be targeted to numerous small lipid droplets, while staining of larger lipid droplets was described as discontinuous [32]. When 3T3-L1 adipocytes are grown under conditions that allow development of both large (more than 10 mm) and small droplets, PLIN appears to accumulate preferentially on small lipid droplets [41, 81] (see also [45]). As mentioned above, fat cells appear to express variable amounts of PLIN that is unrelated to the lipid droplet surface area. To the extent that PLIN deﬁnes where lipolysis occurs, these observations suggest there is heterogeneity with respect to amount and location of droplet structures that are competent for hormone-stimulated lipolysis [41]. 5.3.2 Interactions with CGI-58

CGI-58 was independently discovered by two groups as a PLIN-interacting protein [59, 60]. In preadipocytes, CGI-58 is cytosolic and endogenous or ectopic expression of PLIN results in targeting of CGI-58 to lipid droplets [59]. Double-label immunoﬂuorescence analysis indicates that CGI-58 and PLIN are highly colocalized

5.3 Lipolytic Protein Trafficking

in unstimulated adipocytes [37, 59]. CGI-58 rapidly (within minutes) dissociates from PLIN following forskolin stimulation and becomes progressively cytosolic, while PLIN remains on lipid droplet surfaces [37, 71]. In vitro experiments demonstrated that CGI-58 binds directly to PLIN, but not phosphorylated PLIN [59, 71]. Imaging the molecular interaction of PLIN and CGI-58 in live cells by ﬂuorescence resonance energy transfer (FRET) indicates that PKA activation rapidly triggers dissociation of CGI-58 and PLIN in a manner that is entirely dependent upon PLIN phosphorylation [37]. Double-label immunoﬂuorescence analysis of 3T3-L1 adipocytes demonstrates that stimulation simultaneously decreases the colocalization of CGI-58 with PLIN while increasing it with ATGL [37]. 5.3.3 Interactions with HSL

It is well established that HSL translocates to neutral lipid fractions following PKA activation [8, 82]. PKA-induced HSL translocation to lipid droplets does not occur in cells lacking PLIN and is conferred by ectopic PLIN expression [43, 46]. In cells expressing PLIN, forskolin triggers HSL translocation to droplets containing PLIN, but not to droplets lacking PLIN [37]. Interestingly, although PKA-induced HSL translocation in adipocytes requires PLIN, it does not require PLIN phosphorylation [43]. Experiments with ﬂuorescent reporters demonstrate that HSL translocation begins within 1 min following forskolin stimulation and is essentially complete by 5 min. Translocation results in very close molecular proximity between HSL and PLIN (less than 8 nm) as determined by FRET experiments [37]. Similarly, Miyoshi et al. [43] found that stimulation promotes the ability to chemically cross-link endogenous HSL and PLIN in differentiated embryonic ﬁbroblasts. Cross-linking, FRET, and protein complementation experiments indicate that PKA activation generates a complex containing PLIN and HSL [37, 43]; however, analysis of phosphorylation-defective mutants of PLIN in adipocytes strongly indicates that HSL translocation is not sufﬁcient to initiate lipolysis in the absence of PLIN phosphorylation [43]. Thus, PLIN phosphorylation affects HSL activity beyond its ability to promote translocation, but it is not known whether this effect involves direct interactions with phosphorylated PLIN, or is brought about indirectly by such factors as ATGL activation (and generation of diglyceride substrate) or droplet remodeling. 5.3.4 Interactions with ATGL

In the basal state, ATGL is found in the cytoplasm [37, 57, 62] and on lipid droplets [9, 58], including those containing PLIN [37]. Stimulation elicits minor translocation of ATGL to lipid droplets [37, 83], but does not improve the colocalization of ATGL with PLIN. Unlike HSL, ATGL does not interact with PLIN, as judged by FRET and bimolecular ﬂuorescence complementation assays [37]. The mechanism of ATGL translocation is likely to be indirect, since ATGL is not a substrate for PKA [58].

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Possible mechanisms include PLIN-induced remodeling of the lipid droplet surface and/or interactions with released CGI-58. It should be noted that a considerable amount of ATGL resides on PLIN-containing lipid droplets in the basal state and its activation might be triggered by local release of CGI-58 from PLIN. In this regard, although stimulation provokes release of CGI-58 from PLIN, some CGI-58 is retained on these lipid droplets during PKA activation and could represent sites of ATGL activation [37, 59]. As mentioned above, immunoﬂuorescence studies indicate that forskolin increases the colocalization of CGI-58 and ATGL. Since stimulation does not alter the overall colocalization of ATGL and PLIN, the stimulation-induced increase in colocalization of CGI-58 and ATGL may involve subcellular structures that lack PLIN. 5.3.5 Disruption and Dispersion of Lipid Droplets Following PKA Activation

Recently, Yamaguchi et al. [71] observed the formation of microdroplets within 10 min of PKA activation using coherent anti-Stokes Raman microscopy. As these microdroplets ﬁrst appeared some distance from larger lipid droplets, and seemed to grow in size and number over the course of stimulation, it was proposed that the droplets were not directly derived from fragmentation of larger lipid droplets. It is possible that these microdroplets are formed from the re-esteriﬁcation of mobilized fatty acids; however, the exact source of triglyceride contained in the microdroplets and its relation to lipolysis are presently unclear. Sustained PKA activation produced by forskolin dramatically fragments lipid droplets in cultured adipocytes. In contrast to the microdroplet formation mentioned above, this fragmentation ﬁrst appears after 30 min, with extensive fragmentation seen by 8 h [84]. Given that the appearance of the microdroplets coincides with loss of larger droplets, and the microdroplets contain both PLIN and lipid, it seems highly likely that these structures are derived from larger, PLIN-containing lipid droplets. Signiﬁcantly, this massive lipid droplet fragmentation occurs in the presence of pharmacological inhibition of lipolysis [84]. Further analysis of this phenomenon demonstrated that lipid droplet fragmentation requires phosphorylation of PLIN on Ser492. Fragmentation greatly increases the lipid droplet surface area that is exposed to cytoplasm, as well as dramatically increasing the lipid droplet curvature – either of which could increase lipase activity. Indeed, it was recently reported that mutation of Ser492 reduces PKA-stimulated lipolysis in differentiated embryonic ﬁbroblasts [65]. On the other hand, mutation of Ser517 nearly eliminates PKA-induced lipolysis, yet has little effect on fragmentation. Thus, while fragmentation likely promotes lipolysis, fragmentation per se is not sufﬁcient to initiate it [65]. 5.3.6 Additional Interactions

Caveolin1 (Cav1) is a structural protein of caveolae – membrane invaginations involved in cholesterol metabolism, lipid transport and signal transduction [85].

5.4 Working Model and Unresolved Issues

Adipocytes are among the cell types that express the greatest amounts of Cav1. Cav1 translocates to lipid droplets in response to elevated fatty acids or cholesterol [86, 87] and during sustained lipolytic activation [11]. Importantly, mice lacking Cav1 have dramatically impaired ability to stimulate lipolysis in brown and white adipocytes [88, 89]. The architecture of the lipid droplet surface of adipocytes is disturbed in Cav1deﬁcient mice and these mice are unable to phosphorylate PLIN during b-adrenergic stimulation [88]. Thus, Cav1 is important in organizing the signaling pathway leading from receptors to lipid droplet activation. Recent proteomic analysis has revealed a surprisingly large number of vesicular trafﬁcking proteins – those involved in membrane budding, targeting and fusion – in lipid droplets puriﬁed from a variety of cell lines [9, 11, 13, 90, 91]. Rab5, Rab7, Rab11, Rab14, and Rab18 have been consistently found on lipid droplets in different cell types, implying that they play fundamental roles in lipid droplet functions. Immunoelectron microscopic examination clearly conﬁrmed the speciﬁc localization of Rab18 to lipid droplet surfaces [90, 92, 93], and only the GTP-bound form of Rab18 was targeted to lipid droplets [90]. Moreover, the amount of Rab18 on lipid droplets was positively correlated with the lipolytic activity of the cell [11, 93]. Overexpression of active Rab18 reduces the amount of ADRP on lipid droplets, and concomitantly induces close apposition of endoplasmic reticulum membranes to lipid droplets [90]. Since a major function of Rab-GTPases is to recruit speciﬁc tethering factors that bring transport vesicle and target membrane into close proximity [94], these emerging data indicate that Rabs are bona ﬁde lipid droplet constituents that might be involved in tethering lipid droplets to speciﬁc membranes during lipolytic activation.

5.4 Working Model and Unresolved Issues

A model of PKA-induced lipolytic protein trafﬁcking is depicted in Figure 5.2 and focuses on the central role of PLIN. In the basal state, PLIN and CGI-58 are tightly bound. Some ATGL is localized to lipid droplets containing PLIN; however, ATGL is likely to be inactive, since its coactivator is sequestered by PLIN. PKA activation leads to PLIN phosphorylation, which has two parallel effects. (i) PLIN phosphorylation frees CGI-58 to activate ATGL on lipid droplets surfaces with or without PLIN. (ii) PKA activation promotes the rapid translocation of HSL to PLIN-containing lipid droplets. ATGL activity is directed toward triglyceride substrate and its activation by CGI-58 may be the event that initiates lipolysis. Although HSL has triglyceride hydrolase activity, its role as a DG lipase seems to be rate-limiting [58, 95]. Nonetheless, the coordinated regulation of these lipases at PLIN-containing lipid droplets appears to be essential for full, regulated lipolysis. The model presented above leaves many important questions unresolved. PLIN clearly regulates the activity and accessibility of cellular lipases during PKA activation; however, the biochemical and biophysical bases are uncertain. Targeting of ATGL and HSL to lipid droplet surfaces does not appear to be sufﬁcient to trigger lipolysis in the

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Figure 5.2 Protein trafficking during initiation of lipolysis. Under basal conditions, PLIN and CGI-58 form a complex on the surface of lipid droplets. HSL is mainly cytosolic, while ATGL is localized to lipid droplets, including those containing PLIN. Stimulation by PKA activation leads to trafficking of HSL and CGI-58 (green arrows). Phosphorylation of PLIN releases CGI-58, which then activates ATGL. As ATGL acts exclusively on triglyceride (TG), it is likely that ATGL initiates generation of FFA.

Phosphorylation of HSL promotes its translocation and tight association with Plin. A major component of HSL activity likely depends on generation of diglyceride (DG) substrate from the action of ATGL, while monoglyceride (MG) lipase acts to liberate glycerol and the final FFA. Not illustrated are potential effects of Plin phosphorylation on the biophysical properties of the lipid droplet surface, and the effects of sustained activity on lipid droplet fragmentation and movement.

absence of PLIN phosphorylation [43]. Thus, initiation of lipolysis likely involves PLIN phosphorylation-dependent events beyond simple lipase recruitment. It is unclear whether PKA-dependent release of CGI-58 from PLIN is sufﬁcient to activate ATGL or whether PLIN-induced remodeling of the lipid droplet surface plays a role. PLIN appears to interact with multiple proteins and it will be important to clarify how these interacts are organized. For example, CGI-58 interacts with both ATGL and PLIN, but it is unclear whether these interactions are mutually exclusive within cells. Bimolecular ﬂuorescence complementation experiments suggest that PLIN assembles into higher-order structures with itself [37]. The structure role that PLIN might play in the formation, maintenance and dispersion of lipid droplets is an important topic of future research. The demonstration of PKA-dependent, lipase-independent droplet fragmentation and the identiﬁcation of multiple Rab subtypes on lipid droplets raise a host of questions. Rab-mediated membrane tethering may facilitate lipolytic signaling and/ or shutting of lipolytic products to individual organelles or to the cell surface. The latter could occur through transient intercompartmental contact sites without membrane fusion [96]. It is interesting to note that lipolytic stimulation increases pinocytic activity that shifts a fraction of cell surface G-proteins to endocytic

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The authors wish to thank E. Mottillo for comments and R. Granneman for illustrations. Supported by NIH grants DK 062292 and DK 076629, ADA grant RA50, and the Department of Veterans Affairs.

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Jedrychowski, M.P., Berg, E.A., Costello, C.E., and Gygi, S.P. (2007) J. Lipid Res., 48, 2103–2111. Le Lay, S., Hajduch, E., Lindsay, M.R., Le Liepvre, X., Thiele, C., Ferre, P., Parton, R.G., Kurzchalia, T., Simons, K., and Dugail, I. (2006) Trafﬁc, 7, 549–561. Pol, A., Martin, S., Fernandez, M.A., Ferguson, C., Carozzi, A., Luetterforst, R., Enrich, C., and Parton, R.G. (2004) Mol. Biol. Cell, 15, 99–110. Cohen, A.W., Razani, B., Schubert, W., Williams, T.M., Wang, X.B., Iyengar, P., Brasaemle, D.L., Scherer, P.E., and Lisanti, M.P. (2004) Diabetes, 53, 1261–1270. Cohen, A.W., Schubert, W., Brasaemle, D.L., Scherer, P.E., and Lisanti, M.P. (2005) Diabetes, 54, 679–686. Ozeki, S., Cheng, J., Tauchi-Sato, K., Hatano, N., Taniguchi, H., and Fujimoto, T. (2005) J. Cell Sci., 118, 2601–2611. Turro, S., Ingelmo-Torres, M., Estanyol, J.M., Tebar, F., Fernandez, M.A., Albor, C.V., Gaus, K., Grewal, T., Enrich, C., and Pol, A. (2006) Trafﬁc, 7, 1254–1269. Liu, P., Bartz, R., Zehmer, J.K., Ying, Y.S., Zhu, M., Serrero, G., and Anderson, R.G. (2007) Biochim. Biophys. Acta, 1773, 784–793. Martin, S., Driessen, K., Nixon, S.J., Zerial, M., and Parton, R.G. (2005) J. Biol. Chem., 280, 42325–42335. Grosshans, B.L., Ortiz, D., and Novick, P. (2006) Proc. Natl. Acad. Sci. USA, 103, 11821–11827. Zechner, R., Strauss, J.G., Haemmerle, G., Lass, A., and Zimmermann, R. (2005) Curr. Opin. Lipidol., 16, 333–340. Levine, T. and Loewen, C. (2006) Curr. Opin. Cell. Biol., 18, 371–378. Haraguchi, K. and Rodbell, M. (1990) Proc. Natl. Acad. Sci. USA, 87, 1208–1212.

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6 New Developments in the Lipolytic Processing of Triglyceride-Rich Lipoproteins Andre Bensadoun, Anne P. Beigneux, Loren G. Fong, and Stephen G. Young 6.1 Introduction

Most of the triglycerides in plasma are contained in chylomicrons and very-lowdensity lipoproteins (VLDL). The transfer of this lipid fuel from lipoprotein particles to tissues requires that triglycerides undergo hydrolysis. The hydrolysis of lipoprotein triglycerides is an enzymatic process that occurs along the surface of the capillary endothelium. By far the most important enzyme for triglyceride hydrolysis is lipoprotein lipase (LPL). Inhibition of LPL with an LPL-speciﬁc antiserum results in a complete and nearly instantaneous block of triglyceride hydrolysis [1]. LPL is expressed at signiﬁcant levels in adipose tissue, heart, and skeletal muscle. Although LPL expression is essentially absent in the adult liver, LPL is expressed highly in the rodent liver during the suckling period [2]. The rationale for high hepatic LPL activity in suckling rodents is not fully understood, but this adaptation presumably allows the liver to accumulate lipids for use as fuel. In tissues expressing LPL, the enzyme is synthesized exclusively in parenchymal cells (e.g., adipocytes, cardiomyocytes, and skeletal myocytes) and then transported to the luminal surface of the capillary endothelium. LPL binds avidly to heparan sulfate chains on cultured endothelial cells in vitro [3] and most have presumed that the same type of binding also occurs in vivo. Very recent data indicate that LPL also binds to a newly discovered membrane-tethered protein on the luminal surface of endothelial cells, glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 (GPIHBP1) [4]. LPL expression is regulated differentially in different tissues, providing a mechanism for regulating the delivery of triglyceride fatty acids in a tissue-speciﬁc manner. LPL may have other functions that are independent of its catalytic activity. LPL binds to members of the LDL receptor family and to plasma lipoproteins, providing a potential mechanism for enhancing the uptake of remnant lipoproteins – a function that has been referred to as a bridging function [5]. There is also some evidence that

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LPL can mediate the selective uptake of cholesterol esters from low-density lipoproteinLDL particles [6]. Owing to the central role of triglyceride hydrolysis in plasma lipid metabolism, the molecular mechanisms for the regulation of LPL activity have been studied intensively. These studies have consistently pointed to posttranslational mechanisms as being most important for the short-term regulation of LPL. Important progress in understanding this post-translational regulation was reported recently by the Olivecrona group – they found that the angiopoietin-like proteins play a major role in regulating the enzymatic activity of LPL [7]. LPL is also regulated at the transcriptional level, and this topic has been extensively and expertly presented in several recent articles [8–10]. Chylomicrons and VLDL contain a hydrophobic triglyceride and cholesterol ester core stabilized in the aqueous plasma environment by a polar surface coat of phospholipids, cholesterol, and apolipoproteins. Several of the apolipoproteins on triglyceride-rich lipoproteins modulate the rate of triglyceride hydrolysis, while others are important for mediating the cellular uptake of remnant lipoproteins. A recently identiﬁed apolipoprotein, apo-AV, plays a very important role in enhancing the rate of triglyceride lipolysis in vivo [11]. This chapter focuses on newer aspects of LPL biology, including the regulation of LPL activity by angiopoietin-like proteins, by apo-AV, and by the newly discovered LPL-binding protein, GPIHBP1.

6.2 LPL

Human LPL is a 448-amino-acid glycoprotein. The amino acid sequence of human LPL is highly conserved; 87–94% identical to other mammalian LPLs and 76% identical to chicken LPL [12]. Based on analyses of amino acid sequence and gene structure, LPL is a member of a larger lipase gene family that includes hepatic lipase, endothelial lipase, pancreatic lipase, pancreatic lipase-related protein-1 and -2, and phosphatidylserine phospholipase A1 [13]. Human LPL is N-glycosylated at Asn43 and Asn359. N-linked glycosylation at Asn43 is necessary for both secretion and maturation of active LPL [14]. LPL is catalytically active as a dimer [15]. Dimerization of the enzyme and processing of the oligosaccharides are likely interrelated. Glucose trimming of the high mannose oligosaccharides by glucosidases (but not mannose removal) is essential for dimerization and full enzyme activation [16]. Translocation to the cis-Golgi is not required for the maturation of enzymatically active LPL [16]. In chicken LPL, the oligosaccharide at Asn45 is sulfated [17]. By sequential exoglycosidase digestion, thin-layer chromatography of the released monosaccharides, and the use of glycosylation inhibitors, the sulfated sugar was shown to be a core N-acetylglucosamine. The kinetics of acid hydrolysis of the sulfated sugar indicated the presence of a single class of primary sulfate groups in an ester linkage to the oxygen on C6 of the core N-acetylglucosamine. The relevance of the sulfation of the

6.3 Functional Domains of LPL

carbohydrate chain has not been fully elucidated; desulfation of the enzyme by culturing cells in the presence of chlorate, an inhibitor of sulfate adenyltransferase, has no effect on catalytic efﬁciency of LPL or on the ability of the enzyme to bind to heparan sulfate chains on adipocytes [18]. The tertiary folding pattern of LPL has been inferred from the crystal structure of another member of the same lipase family, pancreatic lipase [19]. The proposed molecular model for LPL structure contains two domains, a large N-terminal domain with the a/b-fold characteristic of the mammalian lipase gene family and a smaller C-terminal domain with a b-sandwich structure. The N-terminal domain contains the catalytic triad of the enzyme (Ser132, Asp156, and His241). By analogy with pancreatic lipase, the active Ser132 is covered with a lid structure (residues 237–261), which is located between two conserved cysteine residues. Upon addition of lipid substrate, the orientation of the lid and of two surface loops changes, so that the lipid substrate is accessible to the catalytic site of the enzyme. LPL activity requires the presence of apo-CII, an apolipoprotein found in both high-density lipoprotein (HDL) and VLDL, for maximal catalytic activity [20, 21]. In the absence of lipid substrates, LPL forms a stable surface ﬁlm with apo-CII [21]. This tight interaction suggests that, in vivo, triglyceride-rich lipoproteins may bind, at least in part, to the surface of the endothelium via the binding of apo-CII on lipoproteins to LPL bound to a speciﬁc endothelial cell receptor.

6.3 Functional Domains of LPL

The availability of the crystal structure for pancreatic lipase [22] and a predicted structure for LPL [19] has made it feasible to identify functional domains of LPL by domain-exchange methods. LPL and hepatic lipase are structurally similar but have signiﬁcant differences in substrate speciﬁcity and binding interactions. Schotz, Wong, and coworkers [23–25] constructed chimeras with the N-terminal domain of LPL and the C-terminal domain of hepatic lipase (LPL/HL chimeras), as well as the reverse chimeras (HL/LPL chimeras). The properties of these chimeras were characterized, with the goal of identifying important functional domains. The regions of LPL necessary for maximal apo-CII activation was localized to both the N-terminal domain and a C-terminal LPL domain. The ﬂuorescent hydrophobic probe 1,10 -bis(aniline)-4,40 -bis(naphthalene)-8,80 -disulfonate (bis-ANS) binds tightly to LPL and inhibits its enzymatic activity [26]. Apo-CII restored LPL activity in a competitive fashion. The bis-ANS binding site was located in a peptide in the center of the N-terminal domain, suggesting that the same sequence may be important for the LPL–apo-CII interaction. The catalytic site of LPL – as predicted from the structure of pancreatic lipase and later conﬁrmed by site-directed mutagenesis [27] – was localized to the N-terminus of LPL by the domain-exchange strategy. The characteristic inhibition of LPL by 1 M NaCl was also dependent on sequences in the N-terminal domain. The major heparin-binding domain and the substrate-binding domains were assigned to the

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C-terminal domain. Extensive studies involving both site-directed mutagenesis and deletion analysis demonstrated unequivocally that the lid sequence covering the catalytic site interacts with lipid substrates and governs enzyme–substrate speciﬁcity [28]. Identiﬁcation of speciﬁc residues essential for heparin binding were guided by site-directed mutagenesis [29] in the four positively charged clusters (clusters 1, 2, 3, and 4) predicted by van Tilbeurgh from the models of LPL structure [19]. In agreement with domain-exchange studies [13], the most signiﬁcant residues for heparin interaction are located in the C-terminal domain [29]. Mutation of Lys321, Arg405, Arg407, Lys409, and Lys416 (cluster 4) in chicken LPL results in signiﬁcantly reduced afﬁnity for heparin. A triple mutant (R405N, R407N, K409N) displayed almost no high-afﬁnity binding to heparin [29]. Amino acid residues that make a lesser contribution to the overall heparin-binding afﬁnity were identiﬁed within the N-terminal domain in clusters 1 and 2 [30], whereas cluster 3 was not found to be a functional heparin-binding site [30]. In the dimeric holoenzyme, there are a total of six functional heparin-binding regions (clusters 1, 2, and 4 on each monomeric subunit). In the monomer, these three functional clusters are predicted to lie on one face of the protein, all within 10–16 A of one another [19]. Studies of a recombinant lipase molecule have suggested that the two LPL monomers are positioned within the dimer in a head-to-tail conformation [31]. The speciﬁc molecular orientation of the heparin-binding domains in the dimeric holoenzyme enzyme awaits clariﬁcation and will likely be deﬁned by crystallization studies of heparin–LPL complexes.

6.4 Regulation of LPL Activity by Angiopoietin-Like Proteins

Changes in adipose tissue LPL activity with fasting and refeeding have been studied extensively as a model of post-transcriptional LPL regulation. Recently, the Olivecrona laboratory showed that decreased LPL activity in adipose tissue after a fast was due mainly to a change in LPL-speciﬁc activity, with little change in total enzyme mass [32]. Interestingly, all of the decreased LPL mass and activity was due to a decrease in the speciﬁc activity of the extracellular pool [33]; there was no change in LPL mass or activity within the adipocyte pool, suggesting that the decreased LPL activity with fasting could be due to the inactivation of LPL within the extracellular pool [34]. The molecular basis for the decrease in LPL speciﬁc activity was recently shown to be due to the conversion, by angiopoietin-like protein 4 (ANGPTL4), of active dimeric LPL to inactive monomeric LPL [35]. A transient interaction of the coil-coiled domain of ANGPTL4 with active LPL leads to the formation of inactive monomeric LPL. The formation of inactive monomer is thought to occur as a result of a change in the conformation of the folded monomer. The amount of ANGPTL4, relative to active LPL, required for enzyme inactivation is quite low, suggesting that ANGPTL4 may act as a catalyst or as an extracellular chaperone. The ANGPTL4 mRNA turns over

6.5 Role of GPIHBP1 in the Lipolysis of Triglyceride-Rich Lipoproteins

rapidly, consistent with a role in minute-to-minute changes in LPL speciﬁc activity within adipose tissue. Interestingly, levels of ANGPTL4 mRNA are correlated with adipose tissue LPL activity, irrespective of nutritional state [35].

6.5 Role of GPIHBP1 in the Lipolysis of Triglyceride-Rich Lipoproteins

GPIHBP1 is a newly discovered endothelial cell protein required for optimal lipolytic processing of triglyceride-rich lipoproteins. GPIHBP1 was ﬁrst identiﬁed, by expression cloning, as a molecule that confers upon CHO cells the ability to bind to HDL and was initially proposed to have a role in cholesterol transport [36]. The structure of GPIHBP1 is intriguing. The molecule contains a hydrophobic region corresponding to a signal sequence. This domain is followed by a highly negatively charged domain, with 17 out of 25 residues in the mouse sequence and 21 out of 25 residues in the human sequence being glutamate and aspartate. Following this acidic domain, there is a cysteine-rich Ly-6 domain similar to that in the receptor for urokinase-type plasminogen activator. The C-terminal domain contains a hydrophobic sequence that speciﬁes the addition of a glycosylphosphatidylinositol anchor. The molecule contains a consensus sequence for N-glycosylation at Asn76. To characterize the function and physiologic importance of GPIHBP1 in vivo, Beigneux et al. [4] created Gpihbp1-deﬁcent mice (Gpihbp1/). Gpihbp1/ mice exhibited severe hypertriglyceridemia – a phenotype that was obvious in all mice by 6–10 weeks of age. On a low-fat chow diet, adult Gpihbp1/ mice have plasma triglyceride levels above 1000 mg/dl, with some having triglyceride levels above 5000 mg/dl. The plasma cholesterol levels in Gpihbp1/ mice are also quite elevated (above 300 mg/dl). The plasma lipid levels in heterozygous knockout mice (Gpihbp1 þ /) are not different from that of wild-type mice. Size-fractionation of plasma lipoproteins from Gpihbp1/ mice by gel ﬁltration revealed that the vast majority of the triglyceride and cholesterol in the plasma was contained in large triglyceride-rich lipoproteins. Sizing of lipoproteins from Gpihbp1/ mice revealed that many were quite immense (15.4% of the particles were greater than 122 nm in diameter). The triglyceride-rich lipoproteins in Gpihbp1/ mice were enriched in apo-B48. Both the Gpihbp1/ mice and wild-type control mice were given a retinyl palmitate clearance test – a test that assesses the clearance rate of chylomicrons from the plasma. In wild-type control mice, the retinyl palmitate levels peaked between 1 and 3 h, and were essentially undetectable by 10 h. In the Gpihbp1/ mice, the peak retinyl palmitate levels were more than 10-fold higher than in the wild-type mice and the high levels of the retinyl esters persisted for more than 24 h [4]. Thus, the clearance of chylomicrons from the plasma of Gpihbp1/ mice was dramatically impaired. Immunoﬂuorescence microscopy revealed that GPIHBP1 is expressed only in the capillary endothelial cells of tissues that express LPL (e.g., adipose tissue, skeletal

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muscle, and heart), colocalizing with CD31 – an endothelial cell marker. Confocal microscopy demonstrated that GPIHBP1 is expressed on the luminal surface of the capillary endothelium in these tissues. GPIHBP1 was not present in the capillary endothelium of the brain – a tissue that relies more on glucose uptake than triglyceride hydrolysis [4]. Owing to the gross hypertriglyceridemia in Gpihbp1/ mice and the localization of GPIHBP1 on the luminal surface of the capillary endothelium, Beigneux et al. [4] hypothesized that GPIHBP1 could be an endothelial cell receptor for triglyceride-rich lipoproteins, LPL, or both. Experiments with GPIHPB1-overexpressing CHOpgsA745 cells (a mutant CHO cell line deﬁcient in the ability to produce heparan sulfate chains) demonstrated speciﬁc and saturable high-afﬁnity binding of LPL to GPIHBP1. The LPL bound to GPIHBP1 was readily releasable by heparin. GPIHBP1 also binds chylomicrons. Fluorescently labeled chylomicrons (isolated from the plasma of Gpihbp1/ mice) bound avidly to GPIHBP1-expressing CHO cells, as judged by confocal ﬂuorescent microscopy. Pretreatment of GPIHBP1-expressing CHO cells with a phosphatidylinositol-speciﬁc phospholipase C eliminated the binding of LPL and chylomicrons to cells, strongly supporting the view that both the LPL and the chylomicrons were bound to a glycosylphosphatidylinositol-anchored molecule on the surface of the cell. Additional experiments revealed that apoAV, a newly discovered apolipoprotein with an important role in triglyceride metabolism, binds avidly to GPIHBP1 [4]. LPL in the vascular bed has been widely thought to be bound to heparan sulfate proteoglycans (HSPGs) on the surface of the endothelium [9]. The discovery of GPIHBP1, a new LPL-binding protein, raised the possibility that Gpihbp1/ mice might have lower-than-normal levels of LPL in the plasma following an injection of heparin. Indeed, we observed reduced amounts of heparin-releasable LPL in Gpihbp1/ mice, compared with wild-type mice, in four independent experiments, although this difference did not achieve statistical signiﬁcance. The average reduction in postheparin LPL activity in Gpihbp1/ mice was 41%. We had expected to observe a greater reduction in heparin-releasable LPL in Gpihbp1/ mice, given the severity of the chylomicronemia. However, it is important to point out that the levels of LPL mass and activity in postheparin plasma may not be a reliable measure of the amount of physiologically relevant, active LPL on the surface of the capillary endothelium. Northern blot studies have indicated that GPIHBP1 and LPL are expressed highly in adipose tissue, heart, and skeletal muscle [4]. GPIHBP1 expression is clearly regulated by fasting and refeeding [4]. It is conceivable that the regulation of GPIHBP1 expression could help controlling the amount of LPL within capillaries, thereby contributing to the regulation of fatty acid delivery to vital tissues. As GPIHBP1 binds both LPL and chylomicrons, and because this molecule is clearly essential for the efﬁcient lipolytic processing of chylomicrons, Beigneux et al. [4] proposed that GPIHBP1 functions as a platform for lipolysis, perhaps by bringing LPL and triglyceride-rich lipoproteins into close proximity on the surface of endothelial cells.

6.6 Role of Apo-AV in Lipolysis

6.6 Role of Apo-AV in Lipolysis

Apo-AV is a newly discovered apolipoprotein that has a major impact on plasma triglyceride metabolism. The gene encoding this protein was identiﬁed by comparative genomics [37] and by differential display in a screen for proteins involved in liver regeneration [38]. The APOAV gene encodes a mature protein with a molecular weight of 39 kDa and is expressed only in the liver. Overexpression of human APOAV in transgenic mice reduces plasma triglyceride level by 70% [39], while targeted inactivation of mouse Apoav increased plasma triglyceride levels by fourfold [37]. Commonly occurring polymorphisms in the human APOAV gene are strongly associated with plasma triglyceride levels [40], and truncating mutations in apo-AV have been reported to cause severe hypertriglyceridemia [41]. Although apo-AV clearly has a major impact on plasma triglyceride metabolism, the concentration of apo-AV in the plasma is extremely low, compared with the concentrations of other apolipoproteins. In one study of normolipidemic subjects, the plasma concentration of apo-AV varied between 24 and 406 mg/l [42]. The molecular mechanisms by which apo-AV modulates triglyceride metabolism are still somewhat unsettled. Owing to its high afﬁnity for lipids, it has been suggested that apo-AV could inhibit lipidation of apolipoprotein B during VLDL assembly [43] and thus decrease VLDL triglyceride secretion. Indeed, in one study involving adenoviral overexpression of apo-AV, a roughly 30% decrease in VLDL triglyceride production was observed [44] with no change in apo-B production rate. The decrease in triglyceride secretion was dependent on the amount apo-AV adenoviral vector that was administered. In transgenic mice overexpressing human apo-AV, however, the hepatic VLDL and chylomicron production rates were not different from control mice [45]. The reason for the apparent discrepancy is not known, but it is likely that the levels of apo-AV expression were quite different in the different studies. A complete deﬁciency of apo-AV in mice delays chylomicron clearance [46] and overproduction of human apo-AV increases the clearance of VLDL [47]. The possibility that apo-AV causes triglyceride lowering by stimulating LPL activity has also been examined, but the results of different studies have not been consistent. In one report [44], LPL activity was stimulated by recombinant apo-AV in a dose-dependent manner (by as much as 2.3-fold). A smaller stimulation in LPL activity, although highly signiﬁcant, was observed by Fruchart-Najib et al. [47] when comparing the hydrolysis of apo-AV–deﬁcient VLDL and the same VLDL sample that had been supplemented with recombinant apo-AV. In another study, however, activation of LPL activity was observed only when the LPL was bound to proteoglycans (HSPG) [45]. Lookene et al. [48] examined three different substrates – VLDL, triglyceride phospholipid emulsions, and dimyristoylphosphatidylcholine (DMPC) liposomes – and failed to show a stimulatory effect of apo-AV on LPL activity. In the same report, these authors reported that DMPC–apo-AV disks bind avidly to a heparin–Sepharose columns. Apo-AV could be eluted from these columns with 0.36 M NaCl. The apo-AV–heparin interaction was also demonstrated by surface plasmon resonance.

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Interaction of VLDL and chylomicrons with the immobilized HSPG or with immobilized HSPG and LPL was enhanced in the presence of apo-AV. These studies strongly suggest that apo-AV promotes the formation of HSPG/LPL/triglyceride-rich lipoprotein complexes. In vivo, the presence of apo-AV could enhance the binding of VLDL and chylomicrons to the surface of capillaries, thereby enhancing lipolysis.

6.7 Newly Discovered Regulators of LPL Activity and their Physiological Significance

Several observations [4] support the concept that GPIHBP1 acts as a platform for lipolysis, bringing LPL into close proximity with triglyceride-rich lipoproteins. (i) Gpihbp1/ mice develop severe chylomicronemia, underscoring the physiological importance of GPIHBP1 in chylomicron clearance. (ii) In vitro, GPIHBP1 binds both LPL and chylomicrons avidly. (iii) GPIHBP1 is expressed on the luminal surface of capillary endothelial cells – where lipolysis is known to occur – and exclusively in tissues that express high levels of LPL. (iv) Apo-AV–phospholipid disks bind avidly to GPIHBP1-expressing CHO cells [4], raising the possibility that a direct interaction between apo-AV and GPIHBP1 could be important for the lipolysis of triglyceriderich lipoproteins. We postulate that the same positively charged region of apo-AV that binds to heparin [48], residues 186–227, interacts with the N-terminal acidic domain of GPIHBP1. It is possible, of course, that other apolipoproteins with heparinbinding domains [49], such as apo-E and apo-B, also bind to GPIHBP1. However, the fact that apo-AV-deﬁcient mice develop hypertriglyceridemia [37] suggests that apoAV and GPIHBP1 are partners in the lipolytic processing pathway. One of the puzzles surrounding Gpihbp1/ mice is that the chylomicronemia is severe, yet signiﬁcant amounts of LPL can be released into the circulation by heparin. With a respectable amount of LPL being released by heparin, why is the chylomicronemia in Gpihbp1/ mice so severe? One possibility is that LPL bound to GPIHBP1 on the surface of capillaries is far more enzymatically active than LPL bound to other molecules, such as HSPGs. Another possibility is that, in vivo, GPIHPB1 is effectively the only site of attachment for LPL within the lumen of capillaries. LPL is likely released by heparin from other sites (e.g., HSPGs within the basement membranes of capillaries, HSPGs on the surface of myocytes and adipocytes), but that LPL is likely irrelevant to chylomicron processing. The LPL that is released by heparin in Gpihbp1/ mice could come exclusively from these other sites and therefore could be functionally irrelevant to triglyceride hydrolysis in the lumen of capillaries. There are other mysteries surrounding GPIHBP1. It will be important to determine if GPIHBP1, like the related urokinase-type plasminogen activator receptor, forms dimeric structures that preferentially partition in lipid rafts [50]. Dimerization of GPIHBP1, or clustering of GPIHBP1 in lipid rafts, could bring triglyceride-rich lipoproteins and LPL into close proximity. The localization of lipolysis complexes in lipid rafts might make physiological sense, as the fatty acid transporter CD36 has also been localized in lipid rafts [51]. Also, it will be interesting to further explore the

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G., Lee, N.S., and Olivecrona, T. (1985) Studies on inactivation of lipoprotein lipase: role of the dimer to monomer dissociation. Biochemistry, 24, 5606–5611. Ben-Zeev, O., Doolittle, M.H., Davis, R.C., Elovson, J., and Schotz, M.C. (1992) Maturation of lipoprotein lipase. Expression of full catalytic activity requires glucose trimming but not translocation to the cis-Golgi compartment. J. Biol. Chem., 267, 6219–6227. Hoogewerf, A. and Bensadoun, A. (1991) Occurrence of sulfate in the asparaginelinked complex oligosaccharide of lipoprotein lipase. J. Biol. Chem., 266, 1048–1057. Hoogewerf, A.J., Cisar, L.A., Evans, D.C., and Bensadoun, A. (1991) Effect of chlorate on the sulfation of lipoprotein lipase and heparan sulfate proteoglycans: sulfation of heparan sulfate proteoglycans affects lipoprotein lipase degradation. J. Biol. Chem., 266, 16564–16571. Van Tilbeurgh, H.V., Roussel, A., Lalouel, J.M., and Cambillau, C. (1994) Lipoprotein lipase. Molecular model based on the pancreatic lipase X-ray structure: consequences for heparin binding and catalysis. J. Biol. Chem., 269, 4626–4633. Scanu, A. (2006) Serum high-density lipoprotein: effect of change in structure on activity of chicken adipose tissue lipase. Science, 153, 640–641. Miller, A.L. and Smith, L.C. (1973) Activation of lipoprotein lipase by apolipoprotein glutamic acid. Formation of a stable surface ﬁlm. J. Biol. Chem., 248, 3359–3362. Winkler, F.K., DArcy, A., and Hunziker, W. (1990) Structure of human pancreatic lipase. Nature, 343, 771–774. Wong, H.R., Davis, R.C., Nikazy, J., Seebart, K.E., and Schotz, M.C. (1991) Domain exchange: characterization of a chimeric lipase of hepatic lipase and lipoprotein lipase. Proc. Natl. Acad. Sci. USA, 88, 11290–11294. Davis, R.C., Wong, H., Nikazy, J., Seebart, K.E., and Schotz, M.C. (1992) Chimeras of hepatic lipase and lipoprotein lipase: domain localization of enzyme-speciﬁc

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properties. J. Biol. Chem., 267, 21499–21504. Hill, J.S., Yang, D., Nikazy, J., Curtiss, L.K., Sparrow, J.T., and Wong, H. (1998) Subdomain chimeras of hepatic and lipoprotein lipase: localization of heparin and cofactor binding. J. Biol. Chem., 273, 30979–30984. Lookene, A., Zhang, L., Tougu, V., and Olivecrona, G. (2003) 1,10 -Bis(aniline)4,40 -bis(naphtalene)-8,80 -disulfonate acts as an inhibitor of lipoprotein lipase and competes for binding with apolipoprotein CII. J. Biol. Chem., 278, 37183–37194. Emmerich, J., Beg, O.U., Peterson, J., Previato, L., Brunzell, J.D., Brewer, H.B., and Santamarina-Fojo, S. (1992) Human lipoprotein lipase. Analysis of the catalytic triad by site-directed mutagenesis of Ser132, Asp-156, and His-241. J. Biol. Chem., 267, 4161–4165. Dugi, K.A., Dichek, H.L., Talley, G.D., Brewer, B. Jr., and Santamarina-Fojo, S. (1992) Human lipoprotein lipase: the loop covering the catalytic site is essential for interaction with lipid substrates. J. Biol. Chem., 267, 25086–25091. Sendak, R.A. and Bensadoun, A. (1998) Identiﬁcation of a heparin-binding domain in the distal carboxyl-terminal region of lipoprotein lipase by site directed mutagenesis. J. Lipid Res., 39, 1310–1315. Hata, A., Ridinger, D.N., Sutherland, S., Emi, M., Shuhua, Z., Myers, R.L., Ren, K., Cheng, T., Inoue, I., Wilson, D.E., Iverius, P.H., and Lalouel, J.M. (1993) Binding of lipoprotein lipase to heparin. Identiﬁcation of ﬁve critical residues in two distinct segments of the amino-terminal domain. J. Biol. Chem., 268, 8447–8457. Wong, H.D., Yang, D., Hill, J.S., Davis, R.C., Nikazy, J., and Schotz, M.C. (1997) A molecular biology-based approach to resolve the subunit orientation of lipoprotein lipase. Proc. Natl. Acad. Sci. USA, 94, 5594–5598. Bergo, M., Olivecrona, G., and Olivecrona, T. (1996) Forms of lipoprotein lipase in rat tissues: in adipose tissue the proportion of inactive lipase increases on fasting. Biochem. J., 313, 893–898.

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(2003) The distribution of lipoprotein lipase in rat adipose tissue. Changes with nutritional state engage the extracellular enzyme. J. Biol. Chem., 278, 11925–11930. Bergo, M., Wu, G., Ruge, T., and Olivecrona, T. (2002) Down-regulation of adipose tissue lipoprotein lipase during fasting requires that a gene, separate from the lipase gene, is switched on. J. Biol. Chem., 277, 11927–11932. Sukonina, V., Lookene, A., Olivecrona, T., and Olivecrona, G. (2006) Angiopoietinlike protein4 converts lipoprotein lipase to inactive monomers and modulates lipase activity in adipose tissue. Proc. Natl. Acad. Sci. USA, 103, 17450–17455. Ioka, R.X., Kang, M.J., Kamiyama, S., Kim, D.H., Magouri, K., Kamataki, A., Ito, Y., Takei, Y.A., Sasaki, M., Suzuki, T., Sasano, H., Kahashi, S., Sakai, J., Fugino, T., and Yamamoto, T.Y. (2003) Expression cloning and characterization of a novel glycosylphosphatidylinositol-anchored high density lipoprotein-binding protein, GPI-HBP1. J. Biol. Chem., 278, 7344–7349. Pennacchio, L.A., Olivier, M., Hubacek, J.A., Cox, D.R., Fruchart, J.C., Krauss, R.M., and Rubin, E.M. (2001) An apolipoprotein inﬂuencing triglycerides in humans and mice revealed by comparative sequencing. Science, 294, 169–173. Van der Vliet, H.N., Sammels, M.G., Leegwater, A.C., Levels, J.H., Reitma, P.H., Boers, W., and Chalumeau, R.A. (2001) Apolipoprotein A-V: A novel apolipoprotein associated with an early phase of liver regeneration. J. Biol. Chem., 276, 44512–44520. Van der Vliet, H.N., Schaap, F.G., Levels, J.H., Ottenhoff, R., Looije, N., Wesseling, J.G., Groen, A.K., and Chalumeau, R.A. (2002) Adenoviral expression of apolipoprotein A-V reduces serum levels of triglycerides and cholesterol in mice. Biochem. Biophys. Res. Commun., 295, 1156–1159. Pennacchio, L.A., Olivier, M., Hubacek, J.A., Kraus, R.M., Rubin, E.M., and Cohen, J.C. (2002) Two independent apolipoprotein A5 haplotypes inﬂuence

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human plasma triglyceride levels. Hum. Mol. Genet., 11, 3031–3038. Marcais, C., Verges, B., Charriere, S., Pruneta, V., Merlin, M., Billon, S., Perrot, L., Drai, J., Sassolas, A., Pennachio, L.A., Fruchart-Najib, J., Fruchart, J.C., Durlach, V., and Moulin, P. (2005) ApoA5 Q139X truncation predisposes to late-onset hyperchylomicronemia due to lipoprotein lipase impairment. J. Clin. Invest., 115, 2862–2869. OBrien, P.J., Alborn, W.E., Sloan, J.H., Ulmer, M., Boodhoo, A., Knierman, M.D., Schultze, A.E., and Konrad, R.J. (2000) The novel apolipoprotein A5 is present in human serum, is associated with VLDL, HDL and chylomicrons, and circulates at very low concentrations compared with other apolipoproteins. Clin. Chem., 51, 351–359. Weinberg, R.B., Cook, V.R., Beckstead, J.A., Martin, D.D., Gallagher, J.W., Shelness, G.S., and Ryan, R.O. (2003) Structure and interfacial properties of human apolipoprotein A-V. J. Biol. Chem., 278, 34438–34444. Schaap, F.G., Rensen, P.C., Voshol, P.J., Vrins, C., van der Vliet, H.N., Chamuleau, R.A., Havekes, L.M., Groen, A.K., and van Dijk, K.W. (2004) ApoAV reduces plasma triglycerides by inhibiting very low density lipoprotein-triglyceride (VLDL-TG) production and stimulating lipoprotein lipase-mediated VLDL-TG hydrolysis. J. Biol. Chem., 279, 27941–27947. Merkel, M., Loefﬂer, B., Kluger, M., Fabig, N., Geppert, G., Pennacchio, L.A., Laatsch, A., and Heeren, J. (2005) Apolipoprotein AV accelerates plasma hydrolysis of triglyceride-rich lipoproteins by interaction with proteoglycan-bound lipoprotein lipase. J. Biol. Chem., 280, 21553–21560. Grosskopf, I., Baroukh, N., Lee, S.J., Kamari, Y., Harats, D., Rubin, E.M., Pennacchio, L.A., and Cooper, A.D. (2005) Apolipoprotein A-V deﬁciency results in marked hypertriglyceridemia attributable to decreased lipolysis of triglyceride-rich lipoproteins and removal of their remnants. Arterioscler. Throm. Vasc. Biol., 25, 2573–2579.

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47 Fruchart-Najib, J., Baug e, E., Niculescu,

L.S., Pham, T., Thomas, B., Rommens, C., Majd, Z., Brewer, B., Pennacchio, L.E., and Fruchart, J.C. (2004) Mechanism of triglyceride lowering in mice expressing human apolipoprotein A5. Biochem. Biophys. Res. Commun., 319, 397–404. 48 Lookene, A., Beckstead, J.A., Nilsson, S., Olivecrona, G., and Ryan, R.O. (2005) Apolipoprotein A-V–heparin interactions. Implications for plasma lipoprotein metabolism. J. Biol. Chem., 280, 25383–25387. 49 Cardin, A.D. and Weintraub, H.J. (1989) Molecular modeling of protein–

glycosaminoglycan interactions. Aterioscler. Thromb. Vasc. Biol., 9, 21–32. 50 Cunningham, O., Andolfo, A., Santovito, M.L., Luzzolino, L., Blasi, F., and Sidenius, N. (2003) Dimerization controls the lipid raft partitioning of uPAR/CD87 and regulates its biological functions. EMBO J., 22, 5994–6003. 51 Zeng, Y., Tao, N., Chung, K.N., Heuser, J.E., and Lublin, D.M. (2003) Endocytosis of oxidized low density lipoprotein through scavenger receptor CD36 utilizes a lipid raft pathway that does not require caveolin-1. J. Biol. Chem., 278, 45931–45936.

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7 Intracellular Fatty Acid Transport, Activation, and Trafficking Doug Mashek 7.1 Introduction

Adipose tissue, perhaps more than any other tissue, is characterized by its lipid content. By deﬁnition, lipids are hydrophobic molecules and, thus, are not soluble in cytosol. Since lipids cannot simply diffuse throughout the cell, numerous proteins are responsible for their movement. The focus of this chapter is to detail these intracellular lipid transport proteins, including their role in facilitating lipid metabolism and mediating the regulatory effects of lipids on physiological functions within the cell as well as their role in disease development. Although there are thousands of unique lipids present in adipocytes, this chapter will center upon proteins involved in the intracellular transport of fatty acids, their activation to acyl-CoAs, and the partitioning of these acyl-CoAs, which serve as important intermediates in many lipid metabolic pathways. Fatty acids are derived from numerous sources within cells, with the predominant sources being exogenous uptake, de novo synthesis or hydrolysis of intracellular lipids such as triacylglycerol or phospholipids. The ﬂux of fatty acids from these sources to their distinct metabolic fates has a large impact on energy metabolism and serves many regulatory functions within cells. As fatty acids and acyl-CoAs are potent signaling molecules, dysregulation of their transport and partitioning appears to play a signiﬁcant role in the development of prevalent diseases such as type 2 diabetes and obesity.

7.2 Fatty Acid-Binding Protein Family

In 1972, a group of researchers discovered that rat tissues contained a small, abundant cytosolic protein that bound fatty acids [1]. This original fatty acid-binding protein (FABP) is now recognized as a member of a multigene family of FABPs that serve a primary role in intracellular fatty acid trafﬁcking. Although the FABP family consists of at least 10 unique members, this chapter will only focus upon the two

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members that are expressed in adipose tissue. The ﬁrst member of this pair, FABP-4 (also known as aP2, adipocyte lipid-binding protein, and A-FABP), is highly expressed in macrophages and adipocytes where it comprises up to 6% of total cytosolic protein [2–4]. The second member, FABP-5 (also known as Mal-1, E-FABP and keratinocyte lipid-binding protein), is expressed in numerous tissues including adipose tissue where its expression is low relative to FABP-4 [5]. FABP-4 and -5 isoforms share considerable homology (52%), and the protein structures of all FABP isoforms are highly conserved and consist of a twisted-barrel comprised of antiparallel b-strands surrounding a hydrophobic core where fatty acids and other lipids bind [6]. FABP-4 and -5 bind fatty acids; however, unlike some other FABP members, they also bind different lipids with various afﬁnities. FABP-4 has the highest binding afﬁnity for saturated long-chain fatty acids although it also binds unsaturated fatty acids and retinoic acid [7–9]. In addition to binding fatty acids, FABP-5 also binds numerous eicosanoids, but unlike FABP-4 it does not bind retinoic acid [2, 10]. FABPs are thought to bind and transport fatty acids through one of two possible mechanisms [11]. The ﬁrst mechanism, which appears to be that used by liver FABP, involves aqueous diffusion of the fatty acid from a donor membrane followed by binding to FABP and diffusion to another acceptor membrane. Thus, fatty acids disassociate from membranes into the cytosol where they are rapidly bound by a particular FABP. Subsequently, fatty acids are released from a FABP and rapidly resorb into a nearby membrane. However, in-line with the second mechanism, most FABPs, including FABP-4 and -5, appear to physically interact with donor membranes and facilitate dissociation of the fatty acid from the lipid bilayer followed by transport through the cytosol to an acceptor membrane. This transfer of fatty acid between FABP and membranes involves transient collision of FABP with membranes. Additionally, the collision between FABPs and membranes appears to be enhanced by the availability of the ligand (i.e., fatty acid) and electrostatic interactions with anionic phospholipids in the membranes [12]. Therefore, fatty acid ﬂux and membrane composition may both contribute to the function of FABPs. 7.2.1 Function and Regulation of FABP-4

Most insights into the physiological role of FABP-4 have been derived from a series of studies utilizing knockout animal models. Although mice that lack FABP-4 appear normal under chow-fed conditions, challenging them with a high-fat diet invokes marked phenotypical changes. When fed a high-fat diet, FABP-4 null mice gain more weight including increased fat mass compared to wild-type C57Bl6/J mice [13]. Despite the increase in weight and fat mass, FABP-4 null mice fed a high-fat diet have lower serum concentrations of insulin, glucose, and triglycerides. The most surprising change is that FABP-4 null mice do not develop insulin resistance, which generally accompanies high-fat feeding induced obesity. Therefore, FABP-4 null mice represent a unique model that uncouples obesity from the common comorbidity of insulin resistance and diabetes. Although FABP-4 is the predominant isoform of FABP in adipose, it is interesting to note that FABP-5 is upregulated

7.2 Fatty Acid-Binding Protein Family

approximately 20-fold in FABP-4 null mice. Therefore, the true effect of FABP-4 knockdown is somewhat confounded by the upregulation of FABP-5. Regardless, it is clear that FABP-5 is unable to completely compensate for the lack of FABP-4. The unique phenotype exhibited in FABP-4 knockout mice raises the question of how the lack of FABP-4 expression speciﬁcally affects adipose tissue metabolism. Further characterization of adipose fatty acid metabolism in FABP-4 null mice has revealed a functional role of FABP-4 in triacylglycerol hydrolysis and fatty acid efﬂux. Under both basal conditions or during isoproterenol stimulation to induce lipolysis, glycerol release from FABP-4 null adipocytes is signiﬁcantly attenuated [14, 15]. These data are supported by in vivo studies using stable isotopes of fatty acids that show nearly a 60% decrease in serum free fatty acid (FFA) appearance rate in FABP-4 null mice [16]. Additionally, relative to wild-type mice, intracellular concentrations of fatty acids are also elevated in FABP-4 null mice [14]. Thus, fatty acid efﬂux is reduced in the absence of FABP-4 and, likely as a consequence, intracellular fatty acids accumulate. Despite decreases in fatty acid release from adipose tissue, FABP-4 null mice have increased or unchanged concentrations of serum FFAs [13, 16, 17]. These data suggest that the rate of whole-body fatty acid disappearance is greatly reduced in FABP-4 null mice. Indeed, fatty acid utilization in FABP-4 null mice is lower than wild-type mice, thus explaining the increased serum FFA pool [16]. The exact mechanism explaining the compensatory changes in fatty acid disposal in nonadipose tissue of FABP-4 null mice remains unknown. The important discovery that FABP-4 directly interacts with and enhances the activity of hormone-sensitive lipase (HSL) provides a key mechanism through which FABP-4 may inﬂuence fatty acid efﬂux [18]. Since this original ﬁnding, it has been shown that fatty acid binding is a prerequisite for the interaction between FABP-4 and HSL [19]. Thereby, only upon fatty acid binding to FABP-4 can it interact with HSL to facilitate its catalytic functions. How does FABP-4 enhance HSL activity? HSL is inhibited by one of its end-products, in this case fatty acids [20]. By binding to fatty acids, FABP-4 releases this inhibition on HSL [21]. Thus, FABP-4-mediated transport of fatty acid away from HSL may mediate triacylglycerol hydrolysis and fatty acid efﬂux by (i) increasing HSL activity through the removal of its inhibitor (i.e., fatty acids) and (ii) transporting fatty acids released from triacylglycerol hydrolysis to the plasma membrane where efﬂux occurs. It will be of interest to determine if acyl-CoAs or FABPs also inﬂuence the activity of the recently discovered adipose triacylglycerol lipase [22]. The increased fat mass in FABP-4 null mice could easily be attributed to decreased rates of lipolysis. However, adipocytes from FABP-4 null mice exhibit elevated rates of de novo fatty acid synthesis, which may also contribute to increased fat mass [17, 23]. The increase in de novo synthesis is reﬂected by increased glucose disposal in adipose tissue of FABP-4 null animals [23]. Despite these novel ﬁndings, the mechanism explaining how FABP-4 alters glucose uptake and conversion to fat has not been identiﬁed. Additionally, it is not known how adipose tissue contributes to improving insulin sensitivity in FABP-4 null mice. As would be expected of a protein so central to adipocyte metabolism, FABP-4 expression is highly regulated. FABP-4, measured at both the mRNA and protein

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level, is higher in subcutaneous adipose tissue compared with omental adipose tissue [24, 25]. This may explain why FABP-4 null mice show more pronounced increases in fat mass in subcutaneous rather than visceral adipose depots [17]. FABP4 expression increases following exposure to fatty acids under both in vitro and in vivo conditions [26–28]. Therefore, perhaps it is not surprising that FABP-4 expression increases in adipose tissue following weight loss in human subjects [29]. Fatty acids mediate these changes in FABP-4 levels primarily through their activation of transcription. Peroxisome proliferator-activated receptor (PPAR)-c is a major transcription factor involved in adipocyte differentiation and energy metabolism. The promoter region of FABP-4 contains the conserved DNA-binding domain for PPAR-c and consequently FABP-4 expression is responsive to PPAR-c activation [30]. Since fatty acids are ligands for PPAR-c, the increased expression of FABP-4 following increased exposure to fatty acids is likely due in large part to PPAR-c activation. As would be expected, FABP-4 expression is also highly upregulated, as is PPAR-c, during adipocyte differentiation [27]. Additionally, the upstream region of the FABP-4 gene contains an activator protein 1 site, a CCAAT/enhancer-binding protein (C/EBP) site and a fat-speciﬁc element, the latter of which is required for expression [31–33]. 7.2.2 Function and Regulation of FABP-5

Likely because of its lower expression, FABP-5 has not been as extensively studied as FABP-4 in adipose tissue. Yet, both transgenic and knockout models of FABP-5 have been developed to help elucidate the function of FABP-5. Compared to wild-type mice, FABP-5 null mice fed a chow diet consumed the same amount of chow, and had similar body weight and composition. Despite the lack of visible changes, FABP-5 null mice have reduced serum triacylglycerol and cholesterol, and increased serum FFAs [34]. However, there are no differences in the above-mentioned metabolites when animals are fed a high-fat diet despite similar changes in body weight and adiposity compared to control animals. FABP-5 null mice also have elevated blood glucose although their insulin sensitivity was improved, but not normalized, in response to high-fat feeding. Similarly, ablation of FABP-5 is able to partially alleviate the insulin resistance in ob/ob mice. Thus, like FABP-4 null mice, the lack of FABP-5 is able to somehow alleviate the development of insulin resistance induced through high-fat feeding or in a genetic rodent model of obesity and insulin resistance. Although FABP-5 null mice do not show altered FABP-4 expression to compensate for the lack of FABP-5, the same is not true in FABP-5 transgenic mice – the level of FABP-4 protein in the FABP-5 transgenic mice is reduced approximately 50% [34, 35]. Overexpression of FABP-5 exacerbates insulin resistance despite similar body weights and fat mass between the transgenic and wild-type mice [35]. As expected, mice transgenic for FABP-5 have increased rates of fatty acid release from adipocytes, thus supporting previous data highlighting an important role of FABPs in fatty acid efﬂux. Through the same mechanism as FABP-4, FABP-5 also interacts and regulates HSL to control fatty acid hydrolysis [19]. Despite the increase in FFA efﬂux from

7.3 Fatty Acid Activation and Channeling

adipocytes isolated from FABP-5 null mice, no changes in serum FFAs were noted, suggesting that whole-body fatty acid utilization increased to compensate for the increased rate of fatty acid appearance [17]. Owing to the compensatory changes in expression of FABP isoforms resulting from genetic manipulation of FABP-4 or -5 expression, a double-knockout model has been created [36]. These mice, which lack both FABP-4 and -5, are completely resistant to diet-induced obesity and insulin resistance. To further characterize these mice, they were crossed with obese leptin-deﬁcient ob/ob mice. The resulting offspring remained obese, but had improved insulin sensitivity. Furthermore, FABP-4 and -5 double-knockout ob/ob mice have decreased fatty liver inﬁltration, but unaltered muscle AMP-activated protein kinase (AMPK) activity. Interestingly, double knockouts on a wild-type background have increased muscle AMPK activity. Thus, these results suggest leptin may mediate some of the observed phenotypical effects of altering FABP expression. In support, other studies performed on FABP-4 and/or -5 knockout mice also show altered adipokine levels, especially those of leptin and adiponectin [17, 36]. Therefore, changes in adipokine expression and secretion may be partially responsible for the effects of FABP-4 and -5 on altering energy metabolism. Unlike FABP-4, expression of FABP-5 is higher in omental fat compared with subcutaneous adipose depots and is higher in lean compared with obese subjects [29, 37]. Additionally, FABP-5 expression is sharply increased upon weight loss even more so than FABP-4. In sharp contrast to FABP-4, the peroxisome proliferator response element is not functional on FABP-5; thus, this isoform is not responsive to PPAR activation [38]. FABP-5 also enhances the fatty acid-mediated activation of PPAR-d, but not other PPAR isoforms [39].

7.3 Fatty Acid Activation and Channeling: Role of Long-Chain Acyl-CoA Synthetases and Fatty Acid Transport Proteins

With the exception of fatty acids destined for efﬂux or eicosanoid synthesis, fatty acids must ﬁrst be activated to their acyl-CoA derivatives prior to entering either anabolic or catabolic pathways. This ATP-utilizing reaction is catalyzed by the acyl-CoA synthetase family of enzymes. The family of acyl-CoA synthetases, which show reactivity towards long chain fatty acids, was originally comprised of ﬁve long-chain acyl-CoA synthetases (ACSLs) [40]. However, it is now recognized that members of the fatty acid transport protein (FATP) family also possess activity to long- or very-long-chain fatty acids [41]. In white adipose tissue, ACSL-1 is the predominant isoform, although its expression varies among adipose depots; the highest expression is noted in subcutaneous adipose tissue [42]. Of the other isoforms, ACSL-3, -4, and -5 are expressed at lower levels in white adipose tissue, whereas in brown adipose tissue ACSL-5, along with ACSL-1, are highly expressed [42]. FATP-1 and -4 appear to be the only members of the FATP family to be expressed at signiﬁcant levels in adipose tissue [43].

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In addition to their unique tissue expression patterns, members of the ACSL and FATP families are membrane proteins that localize to different intracellular organelles. In adipocytes or 3T3-L1 cells, ACSL-1 localizes to the endoplasmic reticulum, plasma membrane, and glucose transporter 4 (GLUT4) vesicles [44–46]. Additionally, in 3T3-L1 cells, ACSL-1, -3 and -4 localize to lipid droplets during b-adrenergic stimulation, but only ACSL-1 is present under basal (i.e., unstimulated) conditions [47]. Therefore, despite being intrinsic membrane proteins, there is direct evidence of transport to and from lipid droplets under different physiological conditions. FATP-1 appears to be localized to many organelles in 3T3-L1 adipocytes including microsomes, nuclei, mitochondria, Golgi, and the plasma membrane, whereas the location of FATP-4 in adipocytes has not been characterized [44, 48]. Similar to the movement of ACSL isoforms to lipid droplets, the shuttling of FATP-1 and ACSL-1 to the plasma membrane is also regulated. Insulin stimulates FATP-1 movement to the plasma membrane although the proportion of total FATP-1 that is present on the plasma membrane remains relatively low (around 10%) [49]. Similarly, the stimulation of GLUT4 translocation to the plasma membrane is thought to be responsible for the movement of ACSL-1 in the presence of insulin [45]. This theory holds that ACSL-1, which is present on Golgi, migrates with GLUT4 vesicles as they bud off of the Golgi and translocate to the plasma membrane in response to insulin. As would be expected from their different intracellular locations, individual ACSL and FATP isoforms distinctly alter fatty acid channeling. These ﬁndings have been veriﬁed in a variety of tissues and cell lines, although the role of distinct ACSL and FATP isoforms, especially the former, in fatty acid partitioning in adipose tissue has been less characterized. It is known that tissue homogenate and organellespeciﬁc acyl-CoA synthetase activity changes in response to dietary or hormonal alterations. For example, total adipose (depot not speciﬁed) acyl-CoA synthetase activity towards palmitate is decreased following a 16 h fast; total activity is also decreased in diabetic rats [50]. Measurements of acyl-CoA synthetase activity in subcellular fractions reveals that although activity in mitochondria/nuclear fractions increases after a 16 h fast, concurrent decreases of activity in microsomes, cytosol, and the plasma membrane account for the net reduction in total cell acyl-CoA synthetase activity. Additionally, insulin treatment of adipocytes results in a small increase in acyl-CoA synthetase speciﬁc activity in low-density microsomes and nearly doubles the activity intrinsic to the plasma membrane. Yet, the amount of ACSL-1 that translocates to the plasma membrane only increases 22%. Thus, more than ACSL-1 translocation is responsible for the marked increase in plasma membrane ACS activity. Perhaps parallel FATP-1 translocation to the plasma membrane or post-translational modiﬁcations such as phosphorylation of either ACSL-1 or FATP-1 account for these differences. Thus, it is evident from these studies that measuring total acyl-CoA synthetase activity may not truly reﬂect actual changes in redistributing acyl-CoA synthetase activity within cells following nutritional or physiological alterations. Acyl-CoA synthetase activity and expression of individual isoforms is regulated differently between fat depots. For example, fasting decreases and refeeding in-

7.3 Fatty Acid Activation and Channeling

creases acyl-CoA synthetase activity towards palmitate in gonadal adipose homogenates, however, these nutritional manipulations have no effect on acyl-CoA synthetase activity in inguinal adipose tissue [42]. The same study showed that protein abundance of ACSL-1 in both adipose depots decreases with fasting and increases with refeeding although the changes are more pronounced in gonadal adipose tissue. Furthermore, exercise decreases acyl-CoA synthetase activity in mesenteric adipose tissue concurrent with decreases in ACSL-1 mRNA levels, whereas exercise had no effect on acyl-CoA synthetase activity and mRNA abundance in subcutaneous fat [51]. Based upon this information, it is tempting to speculate that alterations in speciﬁc ACSL or FATP isoforms may, at least in part, mediate the distinct differences in fatty acid metabolism between visceral and subcutaneous fat depots. During differentiation of 3T3-L1 cells, ACSL-1 mRNA abundance increases approximately 160-fold, whereas mRNA of other ACSL isoforms remains unchanged [52]. As this change in ACSL-1 mRNA coincides with the accumulation of triacylglycerol in differentiating 3T3-L1 cells, it is logical to speculate that ACSL-1 plays a major role in triacylglycerol synthesis. Yet, to date, no studies in adipose tissue have deﬁned the role of ACSL-1 in fatty acid partitioning and triacylglycerol synthesis. Heart-speciﬁc overexpression of ACSL-1 increases triacylglycerol content of cardiomyocytes, whereas overexpression of ACSL-1 in hepatocytes does not alter oleic acid incorporation into triacylglycerol synthesis, but decreases triacylglycerol turnover in addition to increasing phospholipid and cholesterol ester synthesis [53, 54]. It is possible that ACSL-1 serves a different physiological function in adipose tissue compared to liver, which has been suggested [55]. For example, ACSL-1 interacts with FATP-1 in adipocytes [56]. However, since FATP-1 is not highly expressed in liver, the role of ACSL-1 in fatty acid channeling may be different in liver relative to adipose tissue due to the expression of proteins it interacts with. Conversely, ACSL-1 may contribute to the increase in triacylglycerol accumulation in adipose tissue through altering triacylglycerol turnover. Regarding the latter, perhaps ACSL-1 enhances activation of fatty acids released from triacylglycerol hydrolysis and preferentially channels these acyl-CoAs back to re-esteriﬁcation pathways resulting in an increased triacylglycerol pool. Clearly, future studies that manipulate ACSL-1 expression or its post-translational processing will provide valuable information regarding its role in adipose lipid handling. Although ACSL-3, -4, and -5 are expressed in adipose tissue, albeit at relatively low levels, their function in fatty acid activation and channeling in adipose tissue has not been deﬁned [42]. Previous studies have pointed towards a role for ACSL-5 in partitioning exogenous fatty acids directly to triacylglycerol synthesis in hepatoma cells [57]. Additionally, ACSL-4 has a preference for long-chain unsaturated fatty acids, speciﬁcally arachidonic acid [58]. Therefore it has been linked to changes in steroid synthesis and may play an important role in phospholipid metabolism [59]. However, at the moment, we can only speculate about the speciﬁc roles of these isoforms in adipose tissue. In addition to their involvement in fatty acid uptake, FATP-1 and -4 are important enzymes in activating and partitioning fatty acids in adipose tissue. In

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order to study the distinct roles of FATP-1 and -4, Lobo et al. characterized fatty acid handling in stable small hairpin RNA knockdowns of 3T3-L1 cells [49]. Suppressing either FATP isoform results in decreased content of triacylglycerol, diacylglycerol, and monoacylglycerol. Yet, while knockdown of FATP-1 suppresses uptake of C16 : 0, C18 : 1, or C20 : 4, especially in response to insulin, blocking FATP-4 expression has no effect on fatty acid uptake. These data suggest that FATP-1 and -4 may regulate the concentration of these lipid pools through unique mechanisms. In support of these differences, further characterization of the cells shows that rates of palmitate incorporation into the above-mentioned glycerolipids decreases in FATP-1 knockdown cells, but is unaffected in FATP-4 knockdowns. Additionally, basal glycerol and fatty acid release from cells increases in FATP-4 knockdowns, whereas basal fatty acid release decreases in FATP-1 knockdowns. Therefore, it appears that FATP-1 may enhance fatty acid uptake, activation, and channeling to the triacylglycerol synthetic pathway. In contrast to altering synthesis rates, FATP-4 may decrease monoacylglycerol, diacylglycerol, and triacylglycerol content by enhancing the hydrolysis of these glycerolipids. The data elegantly show that FATP-1 and -4 have comparable effects on triacylglycerol content, but mediate these effects through two distinct mechanisms. Additionally, whole-body FATP-1 knockout mice have altered gene expression proﬁles, adipokine levels, and susceptibility to insulin resistance and obesity [60]. However, from these models it is difﬁcult to ascertain the role of the adipocyte relative to other tissues that express these isoforms and to differentiate the effects of intracellular fatty acid and acylCoA metabolism from the effects on fatty acid transport.

7.4 Role of Acyl-CoA-Binding Protein in Acyl-CoA Metabolism

Once formed in cells, acyl-CoAs can be metabolized through numerous pathways. Yet, like fatty acids, acyl-CoAs are also hydrophobic and require proteins to facilitate their intracellular movement. Acyl-CoA-binding protein (ACBP) serves this role (Figure 7.2 highlights the roles of ACBP and FABPs in adipocyte lipid metabolism). The predominant mammalian ACBP is ubiquitously expressed although its highest level of expression is in the liver [5]. Unlike FABPs, which bind numerous ligands, ACBP binds acyl-CoAs exclusively [61] and with high afﬁnity (KD ¼ 1–5 nM) [62]. Intracellular unbound acyl-CoA concentrations are low (below 10 nM) and acyl-CoAs readily incorporate into lipid bilayers such that the acyl chain is inserted laterally into membranes with the CoA headgroup remaining external to the membrane [63]. As a result, the majority of acyl-CoAs appear to be incorporated into membranes with a small fraction remaining in the cytosol. For example, mass spectrometric analysis of liver acyl-CoAs revealed that 98% of acyl-CoAs were bound in membranes [64]. It now appears that ACBP is able to desorb acyl-CoAs directly from membranes [63] in a manner that is dependent upon the charge and curvature of the membranes [65]. Similar to the FABPs, ACBP expression is regulated by PPAR-c [66]. ACBP expression increases dramatically during adipocyte differentiation although it

7.5 Regulation and Function of Distinct Fatty Acid and Acyl-CoA Pools

Figure 7.1 ACSLs and FATPs control fatty acid and acyl-CoA partitioning. FATP-1 appears to activate fatty acids that are destined for triacylglycerol synthesis, whereas FATP-4 may preferentially activate fatty acids released from triacylglycerol hydrolysis and promote their reesterification back into triacylglycerol. ACSL-1 has been suggested to serve a similar role in reesterification in hepatocytes although its role in adipocyte fatty acid metabolism remains

unknown. Both FATP-1 and ACSL-1 may be involved in the activation of fatty acids as they are transported across the plasma membrane. The role of other ACSL isoforms has not been studied. Solid lines represent sources of intracellular fatty acids and dotted lines reflect fates of fatty acids. Abbreviations not listed in text: FA, fatty acid; TAG, triacylglycerol; PL, phospholipid.

appears that other transcription factors in addition to PPAR-c are responsible for this increase [67]. The importance of ACBP upregulation during adipocyte differentiation is highlighted by the fact that antisense knockdown of ACBP blocks differentiation in 3T3-L1 cells [68]. Insights into the function of ACBP in liver are supportive for this dual role in controlling adipocyte differentiation and energy metabolism. Interestingly, liver ACBP expression is under the regulation of both PPAR-a and sterol regulatory element-binding protein (SREBP)-1c, which control catabolic and anabolic pathways of fatty acid metabolism, respectively [69]. Finally, an important function of ACBP in adipocytes might involve the regulation of several enzymes such as ACSLs and acetyl-CoA carboxylase, which produce acylCoAs as the end-product [70]. Similar to the effects of FABPs on HSL, ACBPs alleviate product inhibition and thereby enhance enzymatic activity. Thus, ACBPs can potentially regulate adipose tissue physiology at numerous steps, although few studies have characterized its function in depth.

7.5 Regulation and Function of Distinct Fatty Acid and Acyl-CoA Pools

Intracellular fatty acids and acyl-CoAs are bioactive signaling molecules that serve a multitude of functions within cells beyond being substrates for lipid metabolic

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Figure 7.2 FABPs and ACBP mediate intracellular fatty acid and acyl-CoA metabolism and signaling. Intracellular fatty acids, which can be derived from multiple sources, are activated to acyl-CoAs prior to their metabolism. ACBPs facilitate acyl-CoA synthetase activity by removal of acyl-CoA, which acts through end-product inhibition. Subsequently, ACBP facilitates acylCoA metabolism to triacylglycerol and other lipids or transports acyl-CoAs to the nucleus where the acyl-CoAs can serve as signaling molecules that regulate gene expression; the specific transcription factors regulated by acylCoAs in adipocytes is not known. FABPs bind to

intracellular fatty acids and appear to play a major role in triacylglycerol hydrolysis. FABPs directly interact with and enhance HSL activity by removing fatty acids, which inhibit HSL, and subsequently promote fatty acid efflux by transporting fatty acids to the plasma membrane. FABPs also can transport fatty acids to the nucleus to regulate transcription factors such as PPAR-c to control gene expression. Solid lines represent sources of intracellular fatty acids and dotted lines reflect fates of fatty acids. Abbreviations not listed in text: FA, fatty acid; TAG, triacylglycerol; PL, phospholipid.

pathways (Figure 7.3). Perhaps the most documented function of fatty acids and acylCoAs is their ability to regulate a host of transcription factors to control gene expression. For example, fatty acids and acyl-CoAs are ligands for PPAR-c, which is known to regulate numerous adipocyte functions including differentiation, metabolism and adipokine secretion. Thus, because FABPs, ACSLs, FATPs, and ACBP control the intracellular concentration and distribution of fatty acids and acylCoAs they mediate the effects of these signaling molecules on gene expression. Indeed, overexpression of either FABP-4 or ACBP attenuates the activation of the PPARs, especially PPAR-c [71]. Additionally, both FABP-4 and ACBP colocalize to the nucleus, thus suggesting that these proteins are responsible for the trafﬁcking of fatty acids and acyl-CoAs to the nucleus where they can carry out their regulatory effects on gene expression. Further studies reveal that FABP-4 directly interacts with

7.6 Contribution of Fatty Acid and Acyl-CoA Metabolism to Metabolic Diseases

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Allosteric regulators of metabolic enzymes: ACC, ACSL, ANT, G6PDH, HSL, PDH Protein kinases/phosphatases: AMPK, PKC, PP2A

Ligands for transcription factors (e.g. PPAR- γ ) Inhibitors of transcription factors (e.g. SREBP-1)

ATP-sensitive K+ channels

FAs/acyl-CoAs Monoacylglycerol

Kir channels Membrane trafficking

Diacylglycerol

Protein acylation

TAG β-oxidation Figure 7.3 Intracellular fatty acids and acylCoAs have diverse functions in cells. In addition to serving as substrates for glycerolipid synthesis or b-oxidation pathways, fatty acids and acyl-CoAs also play a large role in regulating energy metabolism, cell signaling and homeostasis in adipocytes. Abbreviations not listed in text: ACC, acetyl-CoA

ER stress

carboxylase; ANT, adenine nucleotide translocator; ER, endoplasmic reticulum; G6PDH, glucose-6-phosphate dehydrogenase; Kir, inwardly rectifying K þ channels; PDH, pyruvate dehydrogenase; PKC, protein kinase C; PP2A, protein phosphatase 2A; TAG, triacylglycerol.

PPAR-c and that its targeting to the nucleus is enhanced under stimulation with fatty acid [39, 72]. Although physical interaction between ACBP and transcription factors has not been shown in adipose tissue, ACBP is known to directly interact with and stimulate hepatocyte nuclear factor-4a activity in liver [73]. In addition to their regulation of gene expression, fatty acids and/or acyl-CoAs regulate other intracellular physiological processes. Acyl-CoAs act as allosteric inhibitors of numerous enzymes involved in pathways of energy metabolism such as HSL and acetyl-CoA carboxylase. Thus, accumulation of acyl-CoAs would signal the cell to block lipolysis or de novo fatty acid synthesis to prevent further production of intracellular fatty acids and acyl-CoAs. Moreover, acyl-CoAs regulate numerous processes involved in cell homeostasis including ATP-sensitive K þ channels [74], protein kinases [75, 76] and endoplasmic reticulum stress [77], the latter of which has been implicated in the etiology of type 2 diabetes [78]. Thus, altering fatty acid and/or acyl-CoA pools can have drastic consequences on cellular physiology well beyond simply altering lipid metabolism.

7.6 Contribution of Fatty Acid and Acyl-CoA Metabolism to Metabolic Diseases

Owing to the intimate role fatty acids and acyl-CoAs play in so many physiological processes, it is not surprising that disruption of fatty acid and acyl-CoA homeostasis

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is closely tied to the development of multiple diseases. For example, polymorphisms in FABP-4, FATP-1, FATP-4, and ACBP are linked to changes in blood lipids and/or risk for insulin resistance and type 2 diabetes (Table 7.1). The importance of FABP-4 is further highlighted with the recent discovery of a novel inhibitor to FABP-4 [79]. Administration of this inhibitor to an apolipoprotein E null mouse model of atherosclerosis decreases aortic lesions during both early and late stages of intervention. Similarly, the FABP-4 inhibitor improved insulin signaling and sensitivity in the ob/ob obese diabetic mouse model. Thus, altering FABP-4, through pharmaceutical therapy or natural allelic variation, may have a great impact on the development of diseases including both coronary heart disease and type 2 diabetes. Although the natural genetic variation in these genes is important in identifying individuals susceptible to disease, it will be of great interest to further identify how environmental factors such as diet interact with these speciﬁc genotypes. For example, if a speciﬁc ACSL or FATP isoform shows preference

Table 7.1 The effects of single nucleotide polymorphisms (SNPs) in genes involved in intracellular fatty acid and acyl-CoA trafficking with risk factors for disease and disease outcomes in humans.

Gene

Polymorphism

Phenotype

Reference

FABP-4

T-87C SNP in the C/EBPbinding site of FABP-4 promoter; decreases FABP-4 expression

[84]

FABP-4

A-376C SNP

FATP-1

A/G SNP in intron 8

FATP-1

A/G SNP in intron 8

FATP-4

Gly209Ser

ACSL-5

SNP in 5 0 -untranslated region; increases ACSL-5 expression in muscle SNP in promoter

decreased plasma triacylglycerol, decreased risk for type 2 diabetes (odds ratio ¼ 0.52 for obese subjects and 0.87 for lean subjects) and cardiovascular disease predicts insulin sensitivity and body composition in conjunction with PPAR-c polymorphism increased plasma triacylglycerol increased postprandial triacylglycerol, altered plasma FFA and LDL particle size decreased body mass index, plasma triacylglycerol, FFA, and glucose, and increased insulin increased responsiveness to weight loss

ACBP

decreased risk for type 2 diabetes

[85]

[86] [87]

[88]

[89]

[90]

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7.7 Conclusions

The intracellular trafﬁcking and partitioning of fatty acids and their acyl-CoA metabolites affect a multitude of intracellular processes. As a result, the regulation of FABPs, ACSLs, FATPs, and ACBP directly control energy metabolism and lipidmediated cell signaling. The consequences of these changes are evident by the striking phenotypes of knockout or transgenic animal models that show the importance of these proteins on whole-body energy metabolism and the development of insulin resistance and obesity. Additionally, genetic variation in these proteins is linked to numerous diseases and altered metabolic functions in humans. Further characterization of these proteins or identiﬁcation of novel proteins involved in fatty acid and acyl-CoA metabolism should signiﬁcantly contribute to our understanding of adipose tissue metabolism and its role in disease etiology. Taken as a whole, this group of proteins has great potential as targets for nutritional or pharmaceutical therapies to help prevent or treat metabolic diseases.

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8 Aquaporins and Adipose Tissue: Lesson from Discovery to Physiopathology and to the Clinic of Aquaporin Adipose (AQP7) Ken Kishida 8.1 Introduction

The metabolic syndrome comprises a cluster of insulin resistance, elevated blood pressure, and atherogenic dyslipidemia, and is a major cause of cardiovascular disease in industrial countries [1]. Accumulation of body fat, especially intraabdominal fat deposits, stands upstream of the metabolic syndrome [1]. In the early 1990s, our group pioneered research on the molecular mechanism of the metabolic syndrome [2–4]. We analyzed the gene expression proﬁle of human adipose tissue in collaboration with the Human Body Map Project team, and found that adipose tissue, especially visceral fat, expressed a variety of genes for secretory proteins, including growth factors and cytokines [5]. Traditionally, adipose tissue has been considered as a simple energy storage organ in which lipogenesis and lipolysis are observed in response to whole-body energy balance, but it is also currently understood as an endocrine organ that secretes a variety of bioactive molecules [5]. We named these adipocyte-derived molecules adipocytokines [1]. We and others have provided evidence for the potential role of dysregulation of adipocytokines in the development of the metabolic syndrome [1]. To ﬁnd a novel therapy for metabolic syndrome, it is necessary to focus on the biology and science of adipocyte that the authors have named adiposcience. Adipocytes hydrolyze triglyceride and rapidly liberate free fatty acids (FFAs) and glycerol into the circulation [6]. The underlying mechanism responsible for glycerol release from adipocytes remains unclear. Our team also discovered a highly expressed gene from the human adipose tissue cDNA library in 1997 [5]. This gene, encoded by a protein belonging to the aquaporin (AQP) family, was named AQP adipose (aquaporin adipose AQPap) [21, 22], which also had glycerol permeability and was later found to be a human homolog of AQP7 [9]. In this chapter, we review a series of studies on AQP7 to deﬁne the function and signiﬁcance of the glycerol gateway molecule, and discuss the relation between AQPs and glycerol metabolism.

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Figure 8.1 Schematic presentation of lipogenesis in adipocytes. Under lipogenic conditions, b-cells in the pancreas secret insulin in response to an increase in plasma glucose concentration. Insulin binds to the insulin receptor located on the surface of adipocytes, transfers GLUT4 to the plasma membrane, and takes glucose into the cell. Intracellular glucose

is converted to glycerol-3-phosphate (glycerol-3P). Insulin also activates LPL located on the cell surface of the vascular endothelium. ActivatedLPL removes fatty acids from intestine-derived CM and liver-derived VLDL, and then fatty acids are taken into adipocytes. Fatty acids and glycerol-3-phosphate are esterified into triglyceride (TG).

8.2 Characteristics of Adipocytes and Gycerol Metabolism in the Mammalian Body

Adipocytes continuously synthesize (lipogenesis) and hydrolyze (lipolysis) triglyceride in response to whole-body energy balance (Figures 8.1 and 8.2) [6]. Adipocytes have a unique feature – lipid droplets occupy a large part of the intracellular region, while the nucleus and cytosome are located in the periphery. In comparison, the nucleus is located in the center of cell in other tissue cells. Thus, adipocytes are morphologically characterized by triglyceride accumulation. The b-cells in the pancreas secret insulin in response to an increase in plasma glucose concentration during feeding. Insulin acts on adipose tissues as well as skeletal muscles, transfers glucose transporter 4 (GLUT4) to plasma membrane, and takes glucose into the cell [11]. In addition, insulin activates lipoprotein lipase (LPL) located on the cell surface of the vascular endothelium. LPL removes fatty acids from intestine-derived chylomicron (CM) and liver-derived very-low-density lipoprotein (VLDL), and then fatty acids are taken into adipocytes [12]. In adipocytes, glycerol-3-phosphates converted from glucose and fatty acids are esteriﬁed into triglyceride.

8.2 Characteristics of Adipocytes and Gycerol Metabolism in the Mammalian Body

Catecholamine

Insulin

Lipolysis HSL

Nucleus

translocation HSL

TG Glycerol transporter ?

FFA transporter

FFA

Glycerol

Portal vein Glycerol transporter ?

Liver

FFA oxidations Gluconeogenesis

Figure 8.2 Schematic presentation of lipolysis in adipocytes, and pathway of glycerol and FFA from adipocytes to liver. Under lipolysis or starvation, sympathetic nerves are activated and catecholamines are increased. Catecholamines stimulate adrenergic receptors located on the surface of adipocytes. Activation of adrenergic receptors results in activation of HSL. HSL

hydrolyzes triglyceride (TG) to FFA and glycerol, and both are released into the bloodstream. FFAs are partly re-esterified to triglyceride and incorporated to lipoprotein, and glycerol is used as a substrate of gluconeogenesis in liver. We hypothesized that AQP7 may function as a glycerol transporter in adipocytes [52].

Fatty acid-binding protein (FABP; aP2) [13], fatty acid translocase (FAT) [14], and fatty acid transport protein (FATP) [8, 15] are recognized as fatty acid transporters in adipocytes (Figure 8.1). In contrast to the feeding state, exercise and/or fasting induce lipolysis in adipocytes [16]. Fasting stimulates sympathetic nerves and elevates catecholamines, such as adrenaline and noradrenalin, which in turn stimulate adrenergic receptors located on the surface of adipocytes. Activation of adrenergic receptors results in converting ATP to cAMP. Elevation of intracellular cAMP activates hormone-sensitive lipase (HSL) by phosphorylation. Phosphorylated HSL hydrolyzes triglyceride to FFA and glycerol, and both are released into the bloodstream. FFA and glycerol are utilized for thermogenesis and gluconeogenesis, respectively (Figure 8.2). During the long history of mammals and humans, starvation has been a matter of life or death. Adipocytes play a crucial role in energy supply under starvation to maintain

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energy homeostasis and have contributed to the survival of animals during long fasting periods. During lipolysis, adipocytes liberate FFA and glycerol rapidly outside the cells. It had been considered that FFA and glycerol are transported outside the cell by simple diffusion. Interestingly, recent studies revealed the participation of several molecules in the transport of FFA, such as FATP [8, 15], FABP [13], and FAT [14]. However, the molecular mechanism involved in the transport of glycerol was poorly understood. The rapid increase in glycerol production during lipolysis results in an acute rise in intracellular osmotic pressure, which could damage the cell. We hypothesized that adipocytes may have a molecule for glycerol (Figure 8.2)

8.3 Adipose Glycerol Channel: AQP7 8.3.1 AQP7: A Putative Adipose-Specific Glycerol Channel

The AQPs are a group of channel-forming integral membrane proteins that selectively transport water and, in some cases, small neutral solutes, such as glycerol and urea [16–18]. The discovery of AQPs led to the ﬁnding that rapid movement of water molecules across the cell membrane is required in specialized cell types or tissues [16–18]. The AQPs are a family of homologous water channels widely distributed in plants, unicellular organisms, invertebrates, and vertebrates [16–19]. There are two subfamilies – the aquaporins, which transport only water, and the aquaglyceroporins, which transport glycerol in addition to water (Figure 8.3a) [19, 20]. Many investigators have demonstrated that AQPs play a crucial role in maintaining water homeostasis, but the physiological signiﬁcance of some AQPs as a glycerol channel is not fully understood. For example, the studies of AQP2, as the gene responsible for nephrogenic diabetes insipidus, highlighted the signiﬁcance of AQP in human diseases [7]. We analyzed the gene expression proﬁle of human visceral and subcutaneous fat to clarify the molecular mechanism of obesity-related diseases [5]. We identiﬁed a novel cDNA belonging to the AQP family during this analysis, and named it AQPap because its mRNA is expressed abundantly in adipose tissues and adipocytes [10, 21, 22]. AQPap is a human counterpart of AQP7 that was independently cloned from rat testes by another group at the same time [9]. In Figure 8.4, each AQP is speciﬁcally expressed in specialized tissues. The adipose tissue expresses two aquaporins, AQP1 and AQP7. AQP7 is highly expressed in white adipose tissue (WAT), brown adipose tissue (BAT), and testes [22]. The testes express a shorter size of AQP7 mRNA because of a different polyadenylation site [10]. A weak expression of AQP7 is also observed in the heart, skeletal muscles, and kidneys [22]. We analyzed the function of AQP7 using in Xenopus oocytes. AQP7-expressing Xenopus oocytes showed water and glycerol permeability [10, 21] (Figure 8.3b). This gain of function was inhibited by HgCl2 and recovered by 2-mercaptoethanol, similar

8.3 Adipose Glycerol Channel: AQP7

AQP2 (WCH-CD) AQP5

(a)

AQP0 (MIP) AQP6 AQP4 (MIWC) AQP1 (CHIP) AQP8 AQP11 AQP12

Mammalian aquaporins out NA P

P NA

in

(GlpF)

NH2

AQP7 (AQPap) AQP3 AQP9

COOH

AQP10

aquaglyceroporins (b)

Water Permeability (pmol/min/oocyte)

(x10-4cm/s)

400 300 200 100 0 H2O

6 5

Glycerol Uptake

4 3 2 1 0

AQP7

Figure 8.3 Phylogenic analysis of the mammalian AQPs and functional analysis of AQP7 in Xenopus oocytes. The phylogenic tree of the mammalian AQP gene family is shown (a). Water-permeable AQPs are AQP0, 1, 2, 4, 5, 6 and 8. Glycerol-permeable aquaglyceroporins are boxed (AQP3, 7, 9, 10 and GlpF). GlpF is the Escherichia coli homolog. AQP11 and AQP12

H2O

AQP7

belong to the unclassified subfamily. Osmotic water permeability (Pf ) and [14C]glycerol uptake of oocytes injected with water (open bar) or 50 ng of human AQPap cRNA (closed bar) are shown (b). The oocytes injected with AQP7 cRNA showed 4.5-fold stimulation of glycerol uptake compared with those of water-injected oocytes [10].

to the other AQPs [21, 22]. These results indicated that AQP7 could be subcategorized as an aquaglyceroporin. In mammals, AQP3, 7, 9, and 10 are considered to belong to the aquaglyceroporin subfamily at present (Figure 8.3a) [19, 20]. AQP1 is a simple water channel (Figure 8.3a). Therefore, AQP7 is the sole aquaglyceroporin in the adipose tissue (Figure 8.4). During the differentiation of 3T3-L1 adipocytes, the amount of glycerol released into the media increases and such an increase correlates with augmentation of the AQP7 mRNA expression level [22]. Many adipose-speciﬁc genes are controlled by the

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AQP0 AQP1 AQP2 AQP3 AQP4 AQP5 AQP6 AQP7 (AQPap) AQP8 AQP9 AQP11 AQP12 36B4 Figure 8.4 Tissue distribution of AQPs in mice. C57Black6 mice (12 weeks old) were sacrificed in the fed state and various tissues were removed. RNAs were extracted and subjected to Northern blot analyses. Shown is Northern blotting analysis of AQPs in the various tissues. From [22, 23].

nuclear factor peroxisome proliferator-activated receptor (PPAR)-c, which is a master regulator of adipocytes differentiation and regulates several adipose-speciﬁc genes at the transcriptional level [24]. PPAR-c forms a heterodimer with retinoic acid X receptor (RXR)-a and binds to peroxisome proliferator response element (PPRE) [24]. We identiﬁed the PPRE sequence in the AQP7 promoter [25]. The heterodimer of PPAR-c and RXR-a binds to the PPRE site of the AQP7 promoter and upregulates AQP7 mRNA expression in adipocytes (Figure 8.5a). Furthermore, transfection of PPAR/RXR and exogenous PPAR-c ligands, thiazolidinediones, stimulated the promoter activity of AQP7 [25]. AQP7 PPRE actually bound to PPAR/RXR complex (Figure 8.5) [25]. AQP7 is the adipose-speciﬁc novel PPAR-c target gene. 8.3.2 Function and Regulation of AQP7 in Adipocytes

Catecholamine activates lipolysis and may accelerate glycerol release from adipocytes through AQP7. Epinephrine does not affect AQP7 gene expression (Figure 8.6a). In 3T3-L1 adipocytes, AQP7 is localized at the periphery of the nucleus in the steady state [22]. However, AQP7 translocates to the plasma membrane following epinephrine stimulation, which induces lipolysis (Figure 8.6b) [22]. Epinephrine elevates

8.3 Adipose Glycerol Channel: AQP7

Figure 8.5 Specific binding of the PPAR-c/ RXR-a complex to the PPRE in the AQP7 promoter. The putative PPRE in the AQP7 promoter is underlined (a). The PPAR-c/RXR-a

complex directs and specifically binds to the PPRE in the AQP7 promoter as shown by electrophoretic mobility shift assay (b) [25].

intracellular cAMP levels through adrenergic receptor and then activates protein kinase A (PKA). Interestingly, similar results are observed in AQP2, which is a key water channel of kidney [26]. Brieﬂy, AQP2 exists in the principal cells of the renal collecting duct. Immunogold electron microscopy studies showed that very little AQP2 protein is found in the apical membrane of collecting duct principal cells, but most AQP2 protein exists in the membranes of intracellular vesicles. However, AQP2 protein relocates to the apical plasma membrane when collecting duct cells are stimulated by vasopressin – an antidiuretic hormone released from the brain [26]. Vasopressin binds to V2 receptor at the basolateral membrane of the renal collecting duct causing activation of a G-coupled adenylyl cyclase cascade that results in phosphorylation of AQP2 by PKA [27]. The phosphorylation site of AQP2 is located at residue 256 on the C-terminus of its protein. Phosphorylated AQP2 moves to the apical plasma membrane. These results suggest that AQP7, as well as AQP2, may be phosphorylated by PKA under the activation of adrenergic receptor. It is necessary to determine the phosphorylation site of AQP7 in the future. Moreover, fasting activates lipolysis. We investigated the regulation of AQP7 expression during fasting and refeeding in animals. In mice, fasting was associated with increased AQP7 mRNA level, whereas refeeding reduced the level in parallel with plasma glycerol levels [10]. These nutrition-related changes in AQP7 and plasma glycerol were in clear contrast to insulin levels (Figure 8.6c) [10]. The expression of

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Figure 8.6 Catecholamine-induced translocation and insulin-mediated repression of the AQP7 through negative IRE. We examined the effect of lipolytic hormone on AQP7 mRNA levels. Administration of epinephrine did not change AQP7 mRNA levels in 3T3-L1 adipocytes (a). When the 3T3-L1 adipocytes were stimulated by epinephrine for 20 min, the signal in the plasma membrane became robust in comparison to the intracellular regions, suggesting that AQP7 was translocated from intracellular regions to the plasma membrane (b) [22]. Next, we examined the effect of fasting on AQP7 mRNA expression in adipose tissues. Adipose AQP7 mRNA levels were increased by

fasting and suppressed by refeeding, similar to the change of glycerol. Plasma insulin levels were decreased in the fasted mice and restored in the refed mice (c). We found negative IRE in the human and mouse AQP7 promoter. Single point mutation analysis of the IRE sequence in the AQPap promoter using the promoter assay in 3T3-L1 adipocytes (d) [10]. The bases in the heptanucleotide IRE are numbered 1–7. The value for the wild-type construct without insulin was arbitrarily set at 1.0. In the lower panel, the percent inhibition of AQPap-mediated luciferase activity in mutant constructs by insulin is plotted below the corresponding mutated base pair.

AQP7 was upregulated in the adipose tissue of streptozotocin (STZ)-induced insulin deﬁcient mice [10]. In fact, in 3T3-L1 adipocytes, insulin suppresses AQP7 mRNA levels in a dose- and time-dependent manners. Many genes negatively controlled by insulin such as glucose-6-phosphatase and phosphoenolpyruvate carboxykinase (PEPCK), which are key enzymes of gluconeogenesis, have a negative insulin

8.3 Adipose Glycerol Channel: AQP7

response element (IRE) in their promoter. We found IRE in the promoter of AQP7 [10]. Single nucleotide substitution analysis revealed that four core nucleotides at the center of AQP7 IRE, GTTT, are necessary for the negative regulation by insulin (Figure 8.6d) [10]. These results indicated that AQP7 mRNA expression is closely regulated by insulin at the transcriptional level. Taken together, plasma adipocytederived glycerol levels through AQP7 are partly determined by insulin and catecholamine (Figure 8.7). 8.3.3 Human AQP7 Genetic Mutation

Next, we studied the mutations in the human AQP7 gene for further clariﬁcation of the signiﬁcance of AQP7. We screened human AQP7 gene mutation in 160 subjects including controls, diabetics, and obese individuals, and three types of polymorphism, which predict three types of missense mutation were identiﬁed: R12C (a C ! T substitution at nucleotide 206 in exon 3 led to substitution of arginine with cysteine at position 12, which resides in the N-terminal cytoplasmic domain),

Fasting (Lipolysis state) Insulin

Catecholamine

PI3K-pathway Nucleus HSL

AQP7

translocation

DNA

transcription

HSL

TG AQP7

FFA

Glycerol

Figure 8.7 Schematic presentation of the regulatory mechanisms that control adipose AQP7 under the fasting state. Catecholamine translocates HSL to the lipid droplets, while related stimuli move AQP7 to the plasma membrane. HSL hydrolyzes triglyceride (TG) to fatty acids and glycerol. Moreover, AQP7 mRNA levels are elevated by the decrease of the insulin

signaling cascade. Thus, long-term regulation of AQP7 is under the control of insulin, whereas short-term regulation is under the control of catecholamines. These two different regulatory pathways of AQP7 ensure the efficient release of glycerol from adipocytes under fasting conditions. PI3K, phosphoinositide-3-kinase.

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one subject; V59L (a G ! C substitution at nucleotide 347 in exon 4 caused substitution of valine with leucine at position 59, which resides in the ﬁrst bilayer-spanning domain), 13 subjects; and G264V (a G ! T substitution at nucleotide 963 in exon 8 led to substitution of glycine with valine at position 264, which resides in the sixth bilayer-spanning domain), six subjects (Figure 8.8a) [28]. When we injected cRNA carrying each mutation, oocytes normally expressed AQP7 protein [28]. V59L and R12C were functionally normal, but the G264V mutation lacked both water and glycerol permeability, while those expressing R12C or V59L mutant proteins retained water and glycerol permeability (Figure 8.8b) [28]. G264V mutation is located in the sixth bilayer-spanning domain. Structural analysis of AQP1 shows that the conserved GxxxG motif in the third and sixth bilayer-spanning domain is important for functional pore conformation of AQP family protein; glycine can be sometimes replaced by alanine in the motif (Figure 8.8c) [29]. In the sixth bilayerspanning domain of human AQP7, A260, G264, and G268 form this motif. The functional defect in the G264V mutant might be caused by a disturbance of this motif (Figure 8.8c) [28]. We found a homozygote carrying the G264V mutation. When

(a)

Glycerol Uptake

(b) 3

G264V

V59L

2

in

1

COOH

NH2

(c)

0

(d)

Transmembrane Helix-Helix Association GXXXG Motif Helix 3 and 6 AQP7 264 GAYLG AQP3

213

GTSMG

AQP9

208

GLNSG

Figure 8.8 Genetic mutations of the human AQP7 gene, and functional analysis of the AQP7 mutant proteins in both Xenopus oocytes and human. Three identified missense mutations, AQP7-R12C, V59L and G264V, in the topology of AQP7. NAA and NPS are amino acid residues composed of NPA motifs highly conserved among AQP family proteins (a). We note the functions of these mutant AQP7 proteins using

WT R12C V59L G264V

H2O Fold change (arbitrary unit)

R12C

out NS P

A NA

3.5

Plasma glycerol

3.0 2.5

WT

2.0 1.5

G264V

1.0 0

0 15 30 Exercise time (min)

glycerol permeability of oocytes expressing the G264V mutant is lower (WT, wild-type) (b). G264V is located in the GXXXG motif in the a- helix structures present in the third and sixth transmembrane region (c). This motif is well conserved in aquaglyceroporins (c). The plasma glycerol level is not increased in the subject homozygous for the G264V mutation during exercise (d) [28].

8.3 Adipose Glycerol Channel: AQP7

subjected to physical exercise after prolonged (20 h) fasting, the subjects with homozygous mutation of G264V showed a similar exercise-induced rise in plasma noradrenalin compared to healthy volunteers, whereas the increase in plasma glycerol was apparently disturbed during exercise (Figure 8.8d) [28]. This result indicates that AQP7 may be a crucial molecule for maintaining plasma glycerol levels in human. However, obesity and diabetes were not observed in subjects with homozygous mutation of G264V. Furthermore, it has been reported that single nucleotide polymorphisms (SNPs) in the AQP7 gene may modulate the risk of obesity and dysregulated glycerol release [30]. Further analysis of the human AQP7 gene and/or frequency of the AQP7 mutation or SNP in subjects with the metabolic syndrome should be performed in the future. 8.3.4 Adipose-Derived Glycerol and Gluconeogenesis through AQP7 – Lessons from AQP7-Deficient Mice and Cells

We identiﬁed only one subject with a loss-of-function mutation of AQP7 [28]. To further clarify the physiological function of AQP7 in vivo, we generated AQP7 knockout mice and analyzed their phenotype [31]. We removed exons 1, 2, and 3, and replaced them with a neo cassette. No difference was found in plasma FFA levels between wild-type and knockout mice, but AQP7 knockout mice exhibit lower portal glycerol concentrations under fasting state than wild-type mice under the same condition [31]. Administration of b3-adrenergic agonist, which speciﬁcally affects adipocytes and enhances lipolysis, results in impaired plasma glycerol elevation in AQP7 knockout mice, but does not modulate the normal increase of plasma FFA in both wild-type and AQP7 knockout mice [31]. Similar results are obtained in in vitro 3T3-L1 adipocytes introduced by RNA interference (RNAi) [31]. Brieﬂy, epinephrinemediated glycerol release is signiﬁcantly disturbed in AQP7 knockdown 3T3-L1 adipocytes, while epinephrine-mediated FFA release from AQP7 knockdown adipocytes is similar to that of 3T3-L1 adipocytes transfected with control RNAi. A longer starvation test demonstrated that AQP7 knockout mice exhibit impaired plasma glycerol elevation associated with severe hypoglycemia in comparison with wild-type mice [31]. The results of a series of studies indicate that AQP7 acts as an adipose glycerol channel in vivo and that adipose-derived glycerol is a signiﬁcant substrate for gluconeogenesis (Figure 8.9a). During the experiments with AQP7 knockout mice, we noticed an unexpected phenotype. There is no difference in body weight of wild-type and AQP7 knockout mice at a young age, but we found that AQP7 knockout mice developed mild obesity after 12 weeks of age [11]. Adipose tissue weights of AQP7 knockout mice are signiﬁcantly heavier than wild-type mice at 20 weeks of age. The mass of the WATwas larger in knockout mice compared to wild-type mice (Figure 8.9b) [32]. Histological analysis shows an increase in hypertrophic adipocytes in AQP7 knockout mice [32]. Recently, Hara-Chikuma et al. [33] also reported an increased number of hypertrophic adipocytes in AQP7 knockout mice, although the body weights of their AQP7 knockout mice were similar to those of wild-type mice. The difference of phenotypes

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Figure 8.9 Schematic presentation of the physiological role of AQP7 based on the analysis of AQP7 knockout mice. (a) Summary of younger mice. Under starvation, AQP7 knockout (AQP7-KO) mice exhibit impairment of plasma glycerol elevation that results in severe hypoglycemia in comparison with wildtype (WT) mice. There is evidence that AQP7 acts as an adipose glycerol channel in vivo and that adipose-derived glycerol is a significant substrate for gluconeogenesis [31]. (b) Summary and photo of older mice. AQP7

knockout mice develop obesity accompanied by adipocyte hypertrophy after 12 weeks of age. The high intracellular glycerol contents induce the enzymatic activity of glycerol kinase in adipose tissues of AQP7 knockout mice at a young age. Glycerol kinase promotes re-esterification of glycerol and accelerates triglyceride (TG) accumulation in adipocytes. Moreover, AQP7 knockout mice exhibit whole-body insulin resistance associated with obesity [32]. Glycerol3-P, glycerol-3-phosphate.

may be accounted for by the genetic background of mice. Why was AQP7 deﬁciency associated with the development of obesity? Intracellular glycerol contents of AQP7 knockout mice are signiﬁcantly higher than those of wild-type mice at a young age [32]. Under normal conditions, the adipose tissue exhibits only low glycerol kinase activity and it is believed that glycerol is not reused for lipogenesis in adipocytes. A recent study reported that glycerol induces conformational changes and enzymatic activity of glycerol kinase, which is a key enzyme in the conversion of glycerol to glycerol-3-phosphate [34–36]. Actually, the activity of adipose glycerol kinase of AQP7 knockout mice is elevated before development of obesity, relative to

8.4 Hepatic Glycerol Channel: AQP9

wild-type mice. A previous study indicated that overexpression of glycerol kinase promotes re-esteriﬁcation of glycerol and accelerates triglyceride accumulation in adipocytes [36]. Knockdown of AQP7 in 3T3-L1 adipocytes using RNAi increases intracellular glycerol content, elevates glycerol kinase activity, enhances oleic acid uptake, and ﬁnally results in triglyceride accumulation (Figure 8.9b) [32]. Thus, AQP7 knockout mice exhibit whole-body insulin resistance associated with obesity. In summary, a deﬁciency of adipose AQP7 inﬂuences not only glycerol metabolism, but also glucose metabolism in vivo.

8.4 Hepatic Glycerol Channel: AQP9 8.4.1 AQP9: A Putative Hepatic-Specific Glycerol Channel

AQP9 was independently identiﬁed in human leukocytes and liver [37], and rat liver [38, 39]. AQP9 is also highly expressed in mouse liver and testes (Figure 8.4). In hepatocytes, immunohistochemistry showed that AQP9 is localized at the sinusoidal plasma membrane [40]. Rat AQP9-expressing Xenopus oocytes exhibit water and glycerol permeability [37, 39]. A series of studies on Xenopus oocytes also demonstrated that rat AQP9 permeates urea, mannitol, sorbitol, and uracil. Consistent with this ﬁnding, another group found that rat AQP9 is permeable to water, glycerol, and urea [39]. These results indicate a broad selectivity of rat AQP9. However, another group found AQP9-induced permeability to be restricted to water and urea in humans [37]. Thus, there are conﬂicting results regarding AQP9-induced permeability between rats and humans. Intra-abdominal visceral fat accumulates mainly in the mesentery [2–4]. The anatomical distribution of intra-abdominal visceral fat indicates that substances released from the visceral fat directly ﬂow into the liver via the portal vein [2–4]. FFA derived from visceral fat during lipolysis elevates liver acyl-CoA synthetase and microsomal triglyceride transfer protein mRNA levels, and reduces degradation of apolipoprotein B. These changes induce the release of apolipoprotein B from the liver and increase plasma triglyceride concentrations. Hypertriglyceridemia, which is often observed in subjects with visceral fat accumulation, is partly accounted for by the increase in FFA derived from adipose tissues [41]. Glycerol, which is another product from adipose triglyceride during lipolysis, ﬂows directly into the liver via the portal vein and becomes a substrate for gluconeogenesis. The liver has glycerol kinase activity and can activate glycerol to use it for gluconeogenesis [34, 36]. Expression of AQP1, 8 and 9 was identiﬁed in the liver (Figure 8.4). The only aquaglyceroporin among these AQPs is AQP9 (Figure 8.3a). AQP9 is considered as the sole glycerol channel in liver and is localized at the sinusoidal plasma membrane facing the portal vein [40]. Taken together, AQP9 may act as a channel of glycerol uptake in the liver (Figure 8.2). AQP9 mRNA levels increase by fasting and decrease by feeding [42]. These changes in AQP9 mRNA are similar to those of glycerol kinase,

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which is a key enzyme involved in the conversion of glycerol to glycerol-3-phosphate, and PEPCK, which is a key enzyme for gluconeogenesis. Insulin suppresses AQP9 mRNA levels in a time- and dose-dependent manner in H4IIE rat hepatocytes. Administration of STZ results in increased AQP9 mRNA [42] and protein [43] levels in insulin-insufﬁcient mice. Promoter analysis demonstrates that insulin closely reduces AQP9 mRNA at the transcriptional level through IRE on AQP9 promoter, as in the regulation of AQP7 by insulin. 8.4.2 Gluconeogenesis through AQP9 – Lessons from AQP9-Deficient Mice

Another group generated AQP9 knockout mice and analyzed their phenotype [44]. In the absence of physiological stress, knockout mice did not display any visible behavioral or severe physical abnormalities. Compared with control mice, plasma levels of glycerol and triglycerides were markedly increased in AQP9 knockout mice, whereas glucose, urea, FFAs, and cholesterol were not signiﬁcantly different. Oral administration of glycerol to fasted mice resulted in an acute rise in blood glucose levels in AQP9 knockout mice, revealing no defect in utilization of exogenous glycerol as a gluconeogenic substrate and indicating a high gluconeogenic capacity in nonhepatic organs. In summary, AQP9 is important for hepatic glycerol metabolism, and may play a role in glycerol and glucose metabolism in diabetes mellitus.

(a)

Visceral Fat AQP7 mRNA

(b) 2

arbitrary unit

3 arbitrary unit

Hepatic AQP9 mRNA

2

1

1

0

0

Fast Fed Lean

Fast Fed Obese

Figure 8.10 Overexpression of adipose AQP7 and hepatic AQP9 in obesity. Total RNA (10 mg) of visceral fat and liver in obese db þ /db þ (n ¼ 8) and lean db þ /db þ (n ¼ 8) mice were analyzed by Northern blotting. Columns and bars represent the mean standard error of the results. Two aquaglyceroporins, adipose AQP7

Fast Fed Lean

Fast Fed Obese

(a) and hepatic AQP9 (b), are coordinately regulated. Both AQPs are upregulated under starvation and downregulated after feeding. However, downregulation of both AQP7 and AQP9 expression is blunted and remains high in obesity [42].

8.6 Dysregulation of AQP7 and AQP9 in Obesity with Insulin Resistance

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8.5 Coordination of Adipose Glycerol Channel, AQP7, and Hepatic Glycerol Channel, AQP9

In the feeding state, an increase in plasma insulin concentration results in suppression of lipolysis and the mRNA expression of adipose AQP7, and in reduced glycerol release from adipocytes (Figure 8.10) [10, 22, 42]. Feeding also reduces hepatic AQP9 mRNA and glycerol-based gluconeogenesis (Figure 8.11) [42]. In the physiological feeding state, high plasma insulin coordinately suppresses the AQP7 mRNA for glycerol release from adipocytes and AQP9 in mRNA for glycerol uptake into liver, through the IRE in the AQP7 and AQP9 gene promoter, respectively [10, 42].

8.6 Dysregulation of AQP7 and AQP9 in Obesity with Insulin Resistance

We also studied the regulation of AQP7 and AQP9 in obesity. Both adipose AQP7 and hepatic AQP9 were suppressed in the fed state of lean mice through insulin Glucose

Fasted state

Glucose

Fed state

Gluconeogenesis

Gluconeogenesis

Liver

Adipocyte

Liver

Adipocyte AQP9

AQP9

FFA

TG

Glucose

FFA

TG

FFA

Glycerol Glycerol AQP7 Portal vein

Glycerol AQP7

Portal vein Glucose

Obesity-Overnutrition (Fed state: Insulin resistance)

Gluconeogenesis Liver AQP9

TG Glycerol AQP7

Portal vein

Adipocyte

Figure 8.11 Schematic presentation of physiological and pathophysiological involvement of adipose AQP7 and hepatic AQP9. The coordinated regulation of AQP7 and AQP9 in the physiological fasted and fed states may lead to systemic glucose metabolism via

adipose-derived glycerol. In contrast, in the pathological fed state of insulin resistance, the coordinated augmentation of AQP7 and AQP9, in spite of hyperinsulinemia, augments the utilization of glycerol for hepatic glucose production [42]. TG, triglyceride.

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(Figure 8.10) [10, 42]. However, the suppression of the two AQPs is impaired in obese and insulin-resistant animals in spite of hyperinsulinemia, resulting in portal hyperglycerolemia, which at least partly participates in the systemic hyperglycemia (Figures 8.10 and 8.11) due to more utilization of adipose-derived glycerol for hepatic glucose production and release [42]. Considered collectively, physiological and pathological coordinated regulation of organ-speciﬁc glycerol channels, adipose AQP7 and hepatic AQP9, may contribute to glycerol and glucose metabolism (Figure 8.11) [42].

8.7 Conclusions

The discovery of AQP has made a great impact on life sciences. Structural and functional analyses of AQPs indicate that AQPs do not only permeate water. Novel metabolic mechanisms have been clariﬁed by the demonstration that some AQPs act as glycerol channels. Glycerol release is enhanced with increased expression of adipose AQP7 to maintain glucose levels during starvation. However, in AQP7 knockout mice glycerol release is disturbed and causes hypoglycemia [23, 31, 32, 45– 47]. Take together, we demonstrated that AQP7 serves as a glycerol channel in adipocytes [23]. Investigation of AQP-dependent glycerol metabolism should provide a pivotal insight for the design of novel therapeutic strategies to combat metabolic syndrome [48–51].

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(2002) A futile metabolic cycle activated in adipocytes by antidiabetic agents. Nat. Med., 8, 1122–1128. Lin, E.C. (1977) Glycerol utilization and its regulation in mammals. Annu. Rev. Biochem., 46, 765–795. Ishibashi, K., Kuwahara, M., Gu, Y., Tanaka, Y., Marumo, F., and Sasaki, S. (1998) Cloning and functional expression of a new aquaporin (AQP9) abundantly expressed in the peripheral leukocytes permeable to water and urea, but not to glycerol. Biochem. Biophys. Res. Commun., 244, 268–274. Tsukaguchi, H., Shayakul, C., Berger, U.V., Mackenzie, B., Devidas, S., Guggino, W.B., van Hoek, A.N., and Hediger, M.A. (1998) Molecular characterization of a broad selectivity neutral solute channel. J. Biol. Chem., 273, 24737–24743. Ko, S.B., Uchida, S., Naruse, S., Kuwahara, M., Ishibashi, K., Marumo, F., Hayakawa, T., and Sasaki, S. (1999) Cloning and functional expression of rAOP9L a new member of aquaporin family from rat liver. Biochem. Mol. Biol. Int., 47, 309–318. Elkjaer, M., Vajda, Z., Nejsum, L.N., Kwon, T., Jensen, U.B., AmiryMoghaddam, M., Frokiaer, J., and Nielsen, S. (2000) Immunolocalization of AQP9 in liver, epididymis, testis, spleen, and brain. Biochem. Biophys. Res. Commun., 276, 1118–1128. Kuriyama, H., Yamashita, S., Shimomura, I., Funahashi, T., Ishigami, M., Aragane, K., Miyaoka, K., Nakamura, T. et al. (1998) Enhanced expression of hepatic acylcoenzyme A synthetase and microsomal triglyceride transfer protein messenger RNAs in the obese and hypertriglyceridemic rat with visceral fat accumulation. Hepatology, 27, 557–562. Kuriyama, H., Shimomura, I., Kishida, K., Kondo, H., Furuyama, N., Nishizawa, H., Maeda, N., Matsuda, M. et al. (2002) Coordinated regulation of fat-speciﬁc and liver-speciﬁc glycerol channels, aquaporin adipose and aquaporin 9. Diabetes, 51, 2915–2921. Carbrey, J.M., Gorelick-Feldman, D.A., Kozono, D., Praetorius, J., Nielsen, S., and

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Agre, P. (2003) Aquaglyceroporin AQP9: solute permeation and metabolic control of expression in liver. Proc. Natl. Acad. Sci. USA, 100, 2945–2950. Rojek, A.M., Skowronski, M.T., Fuchtbauer, E.M., Fuchtbauer, A.C., Fenton, R.A., Agre, P., Frokiaer, J., and Nielsen, S. (2007) Defective glycerol metabolism in aquaporin 9 (AQP9) knockout mice. Proc. Natl. Acad. Sci. USA, 104, 3609–3614. Fruhbeck, G., Catalan, V., GomezAmbrosi, J., and Rodriguez, A. (2006) Aquaporin-7 and glycerol permeability as novel obesity drug-target pathways. Trends Pharmacol. Sci., 27, 345–347. Rodriguez, A., Catalan, V., GomezAmbrosi, J., and Fruhbeck, G. (2006) Role of aquaporin-7 in the pathophysiological control of fat accumulation in mice. FEBS Lett., 580, 4771–4776. Wintour, E.M. and Henry, B.A. (2006) Glycerol transport: an additional target for obesity therapy? Trends Endocrinol. Metab., 17, 77–78.

48 Fruhbeck, G. (2005) Obesity: aquaporin

enters the picture. Nature, 438, 436–437. 49 MacDougald, O.A. and Burant, C.F. (2005)

Obesity and metabolic perturbations after loss of aquaporin 7, the adipose glycerol transporter. Proc. Natl. Acad. Sci. USA, 102, 10759–10760. 50 Lee, D.H., Park, D.B., Lee, Y.K., An, C.S., Oh, Y.S., Kang, J.S., Kang, S.H., and Chung, M.Y. (2005) The effects of thiazolidinedione treatment on the regulations of aquaglyceroporins and glycerol kinase in OLETF rats. Metabolism, 54, 1282–1289. 51 Rojek, A.M., Skowronski, M.T., Fuchtbauer, E.M., Fuchtbauer, A.C., Fenton, R.A., Agre, P., Frokiaer, J., and Nielsen, S. (2007) Defective glycerol metabolism in aquaporin 9 (AQP9) knockout mice. Proc. Natl. Acad. Sci. USA, 104, 3609–3614. 52 Londos, C., Brasaemle, D.L., Schultz, C.J., Adler-Wailes, D.C., Levin, D.M., Kimmel, A.R., and Rondinone, C.M. (1999) On the control of lipolysis in adipocytes. Ann. NY Acad. Sci., 892, 155–168.

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9 Signaling Pathways Controlling Lipolysis and Lipid Mobilization in Humans Max Lafontan 9.1 Introduction

There are two types of adipose tissue, distinguished histologically and functionally – white adipose tissue (WAT) and brown adipose tissue (BAT). Both tissues have in common adipocytes that store lipid in the form of triacylglycerols and control the process of lipolysis where triacylglycerols are broken down to energy-rich nonesteriﬁed fatty acids (NEFAs) and glycerol. Although the control of lipolysis is very similar in both fat deposits, WAT and BAT functional roles differ noticeably. BAT is specialized in the production of heat and persists throughout life in rodents, but probably disappears soon after birth in larger mammals and humans [1]. In humans, BAT is not thought to contribute to a large part of thermogenesis in humans [2]. The physiological role of WAT is the storage of excess energy in the form of triacylglycerols and the supply of energy-rich fatty acids to other tissues, when needed. In fact, WAT exerts a buffering function by a control of the inﬂux of fatty acids into the circulation that occurs after meals and the delivery of NEFAs from the triacylglycerol stores into the circulation according to the bodys need for energy [3]. This is achieved by efﬁcient and highly regulated pathways whereby the triacylglycerols stored in the adipocyte are hydrolyzed and fatty acids delivered to the plasma. In general, fat mobilization and fat deposition are reciprocally regulated, so that fatty acids ﬂow in and out of the adipocyte according to the nutritional and physiological state of the organism. Lipolysis and fat mobilization are regulated by multiple mechanisms [4–8]. Lipolysis regulation depends on the acute interplay between various stimulating and inhibiting pathways. Insulin plays a major role in the control of lipolysis and NEFA disposal; it inhibits the rate of lipolysis and NEFA efﬂux by fat cells. Outside the antilipolytic action of insulin, the physiological role of other agents exhibiting antilipolytic effects (i.e., adenosine, prostaglandins, neuropeptide Y (NPY) and peptide YY (PYY), ligands of nicotinic acid receptor) remains to be elucidated. Fat mobilization is stimulated primarily (at least acutely) by catecholamines and

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natriuretic peptides. Catecholamines (epinephrine and norepinephrine) acting through b-adrenoceptors stimulate adenylyl cyclase, producing cAMP from ATP. Natriuretic peptides (atrial natriuretic peptide (ANP) and brain natriuretic peptide (BNP)), acting through natriuretic peptide receptor (NPR)-A that possesses guanylyl cyclase activity, generate cGMP; the two second messengers-dependent pathways are able to stimulate lipolysis (Figure 9.1). The lipolytic effect of other factors such as growth hormone and tumor necrosis factor (TNF)-a appears after long-lasting exposure of adipocytes to the cytokine [8].

Figure 9.1 Major pathways involved in the control of human fat cell lipolysis. Catecholamines stimulate both b1/b2- and a2adrenergic receptors in human fat cells. ANPs, acting via type A (NPR-A) and C (NPR-C) receptors, and insulin are also involved in the control of lipolysis. Protein kinases (PKA and PKG (cGK-I)) activated by intracellular increment of cAMP and cGMP, respectively, are involved in target proteins phosphorylation. Perilipin phosphorylation induces an important physical alteration of the droplet surface that facilitates the action of HSL and dissociation of the ATGL regulator CGI-58 from perilipin. CGI58 promotes the triacylglycerol lipase activity of ATGL. PKA-dependent HSL phosphorylation

promotes its translocation from the cytosol to the surface of the lipid droplet, and its activation. Both activated lipases contribute synergistically to the initiation of lipolysis. Docking of ALBP to HSL favors the fat cell outflow of NEFAs released by the hydrolysis of triglycerides. Glycerol outflow from fat cell is controlled by AQP7. PKA and PKG are also able to phosphorylate a number of other substrates and can also influence the secretion of various adipocyte productions (not shown in the diagram). AC, adenylyl cyclase; AQP7, aquaporin 7 GC, guanylyl cyclase; PI3-K, phosphatidylinositol-3-phosphate kinase; (P) phosphorylation of HSL and perilipin, ( þ ) stimulation; () inhibition.

9.2 Role of Lipases in the Regulation of Hydrolysis of Fat Cell Triacylglycerols

9.2 Role of Lipases in the Regulation of Hydrolysis of Fat Cell Triacylglycerols 9.2.1 Hormone-Sensitive Lipase

The rate-limiting step in lipolysis is the hydrolysis of triacylglycerols by adipocyte lipases (Figure 9.1). The hydrolysis of triacylglycerols involves lipases and lipid droplet-associated proteins, the activation/inhibition of which is under the tight control of signaling molecules. Triacylglycerols are broken down into diacylglycerols and then into monoacylglycerols. The ﬁnal step in the hydrolysis of triacylglycerols is catalyzed by monoacylglycerol lipase – an abundant lipase that is not regulated by hormonal signals [9]. At each step of hydrolysis, one molecule of free fatty acid appears and ﬁnal step generates fatty acid and glycerol, which are released by adipocytes. Hormone-sensitive lipase (HSL) is a multifunctional enzyme that possesses triacylglycerol, diacylglycerol, cholesterol ester, and retinyl ester hydrolase activities [8, 10]. The main metabolic role of HSL is hydrolysis of triacylglycerols and diacylglycerols. HSL activation is induced by its phosphorylation by protein kinase A (PKA) and protein kinase G (PKG) – an action leading to the activation of its catalytic activity. HSL is mainly distributed in the cytosol in the resting state; upon its phosphorylation by PKA and PKG, it is translocated to the surface of lipid droplets that are coated with perilipin. Under basal conditions, perilipin suppresses lipolysis by blocking access of the lipases to the lipid droplet; it is also phosphorylated by PKA and PKG [11]. Perilipin phosphorylation is essential for the fragmentation and dispersion of the lipid droplet, and full stimulation of lipolysis by catecholamines and other lipolytic hormones [12–14]. HSL translocation requires the phosphorylation of both HSL and perilipin [15]. Adipocyte lipid-binding protein (ALBP), also known as fatty acid binding protein 4 (FABP4), is another regulator of HSL; it interacts with the N-terminal region of HSL and forms an ALBP–HSL complex. The lipolytic activity of HSL is increased through its ability to bind and sequester NEFAs via such a speciﬁc protein–protein interaction. The ALBP–HSL complex translocates to the surface of the lipid droplets upon lipolytic stimulation and contributes to NEFA outﬂow [16]. Glycerol outﬂow from fat cells is critical for normal triglyceride/fatty acid metabolism; it is controlled by aquaporin 7 (AQP7) [17]. Glycerol transport defect due to AQP7 gene mutation in humans [18] or gene invalidation in mice results in obesity and insulin resistance in mice [19]. 9.2.2 Adipose Tissue Triglyceride Lipase

A novel lipase, adipose tissue triglyceride lipase (ATGL), belonging to a family of closely related proteins containing a patatin-like domain [20–22] has recently been discovered. ATGL has a triglyceride activity but does not show signiﬁcant diglyceride lipase activity. Together with HSL, ATGL participates in mouse adipose tissue lipolysis; these two enzymes are responsible for more than 95% of triacylglycerol

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hydrolysis activity in mouse fat cells [23, 24]. In mice, ATGL deﬁciency causes a drastic reduction in stimulated lipolysis and ATGL is required for PKA-stimulated lipolysis in adipocytes derived from mouse embryonic ﬁbroblasts [13, 22, 24]. In human fat cells, ATGL appears to be of less importance than HSL in regulating catecholamine-induced lipolysis and cannot replace HSL when the enzyme activity is blocked by pharmacological inhibitors [25, 26]. ATGL participates in basal lipolysis; HSL blockade completely inhibits glycerol release, reduces nonhormonal (basal) lipolysis by 50%, and partly suppresses NEFA release initiated by lipolytic agents. ATGL is activated by a key coactivator, comparative gene identiﬁcation-58 (CGI-58; also identiﬁed as a/b-hydrolase domain-containing-5) [27, 28]. It belongs to the esterase/thioesterase/lipase subfamily of proteins structurally characterized by the presence of a/b-hydrolase folds. Mutations in the CGI-58 gene are associated with a genetic disease characterized by neutral lipid accumulation in several tissues (i.e., Chanarin–Dorfman syndrome). As CGI-58 interacts with perilipins on the surface of the lipid droplets, a complex interplay has been proposed between ATGL, perilipins, and CGI-58 [27, 29, 30]. Trafﬁcking and interaction of these key proteins has recently been reassessed in 3T3-L1 adipocytes and the role of perilipin was clariﬁed. Perilipin mediates hormone-stimulated lipolysis via direct and indirect mechanisms. Perilipin directly regulates the access of HSL to substrate via close, if not direct, interactions, as previously observed [11]. It also indirectly controls ATGL activity by regulating its accessibility to its coactivator. CGI-58 strongly associated with perilipin in the unstimulated (i.e., basal state). PKA activation rapidly phosphorylates perilipin and promotes dissociation of CGI-58 from perilipin – an event followed by ATGL activation [28]. In vitro data suggests that CGI-58 is less efﬁcient in stimulating human than mouse ATGL [27]. In the absence of this factor, the lipase activity for ATGL is much lower than that of HSL in protein extracts of human adipose tissue [31].

9.3 Adrenergic Control of cAMP Production, Lipolysis and Lipid Mobilization

Catecholamines are important stimulators of NEFA release under conditions of stress and during exercise. Epinephrine and norepinephrine stimulate and/or inhibit lipolysis depending on their relative afﬁnity for the b1/b2- and a2-adrenergic receptor subtypes [32], the relative number adrenergic receptors in the fat cell membrane, and their coupling efﬁciency to heterotrimeric G-proteins involved in signal transduction (Gs- and Gi-protein, respectively). Human and rat epididymal fat cell membranes contain the short and long forms of Gas (largely the long isoform), and three forms of Gai protein (ai1, ai2 and ai3). Both b35 and b36 subunits have also been identiﬁed, whereas Gc subunits are not known [4]. The pathways are summarized in Figure 9.1. Human adipocyte responsiveness to catecholamines differs according to the anatomical location of adipose tissue [33–35]. Decreased catecholamine-induced lipolysis has been reported in the adipocytes of obese men and women [36–39]; it was attributed to decreased expression of b2-adrenergic receptor, and increased expression of a2-adrenergic receptors and a2-adrenergic responsiveness. Lipolysis

9.3 Adrenergic Control of cAMP Production, Lipolysis and Lipid Mobilization

dysfunction could also affect downstream elements of the lipolytic cascade (i.e., expression level of Gs/Gi-proteins, modiﬁcations of the catalytic and regulatory components of PKA complex or expression level of lipases (i.e., HSL and ATGL) and their regulators (i.e., perilipins and CGI-58)) [4, 40]. Decreased catecholamineinduced lipolysis and low HSL expression constitute a possibly primary defect in familial combined hyperlipidemia [41] and obesity [25, 42]. Concerning the b-adrenergic lipolytic drive, human fat cell lipolysis is mainly regulated by b2- and b1-adrenergic receptor stimulation [43]. No evidence could be provided for b3-adrenergic receptor-dependent lipolysis when isoproterenol, norepinephrine, or epinephrine were used to stimulate human fat cells in vitro [43, 44]. The drug previously used (i.e., CGP12177, which possesses both agonist and antagonist properties at b-adrenergic receptors) to assess the presence of b3-adrenergic effects in human fat cells [45, 46] acts, in fact, via an atypical state of the b1-adrenoceptor when this compound is used at higher concentrations for fat cell stimulation [47, 48]. Conﬁrmation of the lack of b3-adrenergic responsiveness in fat metabolism in humans has also been provided by in vivo studies [49–51]. A number of controversial results have been published concerning a polymorphism in codon 64 (Trp64Arg) of the b3-adrenergic receptor gene and the development of obesity and obesity-related disorders [52]. The subjects bearing the b3-adrenergic receptor variant, even the heterozygotes, had lower resting activity of the autonomic nervous system – a dysfunction that could be responsible for the lower resting metabolic rates described in some obese patients [53]. The studies performed until now have been unable to convincingly demonstrate a physiological contribution of a b3-adrenergic receptor to the regulation of lipolysis, lipid mobilization, and energy expenditure and lipid oxidation in humans. Concerning physiological regulations, sympathetic nerve ﬁbers traveling in cutaneous nerve fascicles exert a regulatory inﬂuence on subcutaneous fat tissue in humans [54]. Profound unresponsiveness to neurally stimulated lipolysis has been described in the subcutaneous adipose tissue of obese subjects [55]. Reduced b2adrenergic lipolytic responsiveness has been reported in fat cells from subjects with isoproterenol resistance [56] and in obese subjects [37]. Increased antilipolytic responsiveness linked to a2-adrenergic receptor stimulation has also been found in subcutaneous adipocytes of the obese of both sexes [36, 38]. The lipolytic defects have been conﬁrmed in in vivo studies [57–59]. Using in situ microdialysis, a speciﬁc impairment in the capacity of b2-adrenergic receptors agonists to promote lipolysis has been reported in the abdominal subcutaneous adipose tissue of obese adolescent girls [60]. It is suspected that a dysfunction of the b2-adrenergic pathway or of the b2adrenergic receptor density may play a role in the etiology or maintenance of a relatively increased fat mass and, consequently, obesity [61]. Further evidence for a putative role of b2-adrenergic receptors in the etiology of obesity is also provided by the discovery of three polymorphisms (the Gln27Glu, Arg16Gly variants and two polymorphic sites, T ! C substitution at 47 and T ! C substitution at 20, located in the 50 -leader cistron) of the b2-adrenergic receptor gene that are associated with obesity [62–65]. Interference for the b2-adrenergic receptor gene in the distribution of visceral adipose tissue, but not subcutaneous, was

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proposed [66]. Increased body weight, body fat, and obesity have also been described in men bearing the Gln27Gln genotype [67]. However, obese individuals bearing the Gln27Gln b2-adrenergic receptor phenotype may beneﬁt from physical activity to reduce their weight [68]. Some discrepancies, probably related to the poor delineation of obese phenotypes by the various investigators that have approached the problem, still persist [69, 70]. To conclude, these ﬁndings suggest that polymorphisms in the coding and noncoding sequences in the human b2-adrenergic receptor gene could be of major importance for obesity, energy expenditure, and b2-adrenergic receptordependent lipolytic function. The b1-adrenoceptor gene contains two nonsynonymous single nucleotide polymorphisms, Ser49Gly and Gly389Arg, which are both functional in human cells lines [71, 72]. Polymorphism of the b1-adrenoceptor gene inﬂuences long-term weight gain and the incidence of adult-onset overweight in women [73]; functional assays on lipolysis were not performed. To conclude, full badrenergic activation of human fat cell lipolysis usually requires synergistic activation of b1- and b2-adrenergic receptors [74]. A b2-adrenergic defect could be sufﬁcient to alter the normal b-adrenergic responsiveness, even if b1-adrenergic effects are found to be preserved in obese subjects [61]. In addition any reduction of b2-adrenergic receptor-mediated lipolytic response will disturb the normal functional balance existing between a2- and b-adrenergic receptor-mediated effects in human fat cells. Thus, the reduction of the b-adrenergic receptor-dependent lipolytic responsiveness initiated by the physiological amines will be ampliﬁed, thus leading to human fat cells with very weak lipolytic responses. Concerning a2-adrenergic receptors, activation of a2-adrenergic receptors by epinephrine and norepinephrine impairs the b-adrenergic component of catecholamine-induced lipolysis [75, 76]. In human subcutaneous fat cells where a2-adrenergic receptors outnumber b-adrenergic receptors, the preferential recruitment of the a2-adrenergic receptors at the lowest catecholamine concentrations inhibits lipolysis [32, 34]. The strongest a2-adrenergic effect has been observed in the adipocytes from subcutaneous adipose tissue from men and women (Figure 9.2a); a2-adrenergic receptors are particularly expressed in subcutaneous adipocytes from obese subjects [36, 39]. Moderate weight loss leads to a higher adipose cell lipolytic efﬁciency that is associated with changes at receptor levels (mainly increased b2- and decreased a2-sensitivities) [77]. One of the major determinants of a2-adrenergic receptor expression is fat cell hypertrophy; a positive correlation exists between human fat cell diameter and a2-adrenergic receptor number while the inverse relationship is observed for b1/b2-adrenergic receptors. The higher the fat cell size, the lower the lipolytic responsiveness is (Figure 9.2b). It is a phenomenon that was also observed in fat cells of various animal models (golden hamster, rabbit) [78]. Interestingly, in animal models, strong fat cell size reduction induced by calorie restriction is associated with a reduction of a2-adrenergic receptor expression [79]. The mechanisms driving this cell-size-related regulation of fat cell adrenergic receptors expression are unknown. Utilization of the in situ microdialysis technique has allowed a better demonstration of the relative contribution of the b1/b2- and a2-adrenergic receptors in the control of lipid mobilization in vivo [49, 80, 81]. Administration of an a2-agonist

9.3 Adrenergic Control of cAMP Production, Lipolysis and Lipid Mobilization Receptor number (fmol/mg protein)

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Figure 9.2 Heterogeneity of human fat cell adrenergic receptor distribution: incidence of the anatomical location and of fat cell size on fat cell adrenergic receptor distribution. (a) Comparative study of the distribution of b1/b2and a2-adrenergic receptors in crude white fat cell membranes from omental, abdominal, and femoral fat deposits from women. The a2- and b1-/b2-adrenergic receptor numbers were determined from saturation binding studies on crude fat cell membranes using [3H]RX821002 and [3H]CGP12177, respectively. Bmax, the total number of binding sites, was determined from Scatchard analysis of the saturation data. The values are means standard errors (bars) of 10–17 separate experiments. P < 0.05; P < 0.01; P < 0.001; significantly different from

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the corresponding values determined in omental fat cells (by Students paired t-test). (b) Relationships existing between b1/b2- and a2adrenergic receptor (AR) expression and mean fat cell diameter in human fat cells originating from omental and subcutaneous fat deposits. The a2- and b1-/b2-adrenergic receptor numbers were determined using saturation binding studies on intact fat cells with [3H]RX821002 and [3H]CGP12177, respectively. The a2-adrenergic receptor expression was positively correlated to the increment of fat cell diameter while an inverse correlation was found for the b1/b2-adrenergic receptors. A similar relationship has also been observed in hamster and rabbit white fat cells.

(clonidine) directly in the microdialysis probe has not been fully conclusive to attribute a physiological role to a2-adrenergic receptors due to potent vasoconstriction induced by clonidine [82]. Exercise-induced lipolysis is impaired in subcutaneous adipose tissue in obese men and women. The physiological stimulation of adipocyte a2-adrenergic receptors during exercise contributes to this impairment since the blunting of lipid mobilization was suppressed by local administration of an a2-adrenergic receptor antagonist [83] (Figure 9.3a and 9.3b). In heavily trained men, it was impossible to reveal any a2-adrenergic effect in the reduced subcutaneous fat deposits of such subjects [84] (Figure 9.3c). Sex-related differences in the lipid-mobilizing efﬁcacy of physical exercise have been revealed in obese women.

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Figure 9.3 Demonstration of the a2adrenergic receptor-dependent modulation of lipid mobilization in human subcutaneous adipose tissue during exercise. Comparative study of exercise-induced lipid mobilization in subcutaneous adipose tissue of men and women. Time course of extracellular glycerol concentration changes in subcutaneous adipose tissue, calculated from dialysate glycerol levels, during 60-min cycle-ergometer exercise in lean (a), obese (b), and trained (c) men. Comparison was performed with changes occurring in subcutaneous adipose tissue, during 45 min cycle-ergometer exercise in obese women (d). The nonselective a1/a2-adrenergic receptor antagonist, phentolamine, was or was not added to the perfusion medium of the microdialysis probe. Data are means

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standard errors from seven to 10 separate experiments. Physiological stimulation of adipocyte a2-adrenergic receptors strongly impairs exercise-induced lipid mobilization in subcutaneous adipose tissue in obese subjects (the effect is stronger in male than in female adipose tissue). The blunting of lipid mobilization induced by exercise-induced norepinephrine (Nepi)/epinephrine (Epi) release is completely suppressed by the local administration of a a1/a2-adrenergic receptor antagonist. Plasma levels of catecholamines are given at rest (R) and at the end of exercise (E). Epinephrine is suspected to be an important regulator in the control of lipid mobilization through activation of antilipolytic a2-adrenergic receptors in human subcutaneous adipose tissue during exercise.

The weak a2-adrenergic responsiveness observed in abdominal subcutaneous adipose tissue of obese women was also reduced after a moderate hypocaloriﬁc diet [85]. Gender differences in the adrenergic regulation of lipid mobilization have been reported in nonobese patients performing short submaximal exercise bouts (75% maximum O2 uptake, 30 min duration); however, the nature of the differences has not been interpreted by the authors [86]. Mechanisms remain still poorly understood.

9.4 Control of cAMP Production by Adenylyl Cyclase Inhibitors – Inhibition of Lipolysis

9.4 Control of cAMP Production by Adenylyl Cyclase Inhibitors – Inhibition of Lipolysis

Neuropeptides, paracrine factors, and autacoid agents (adenosine, prostaglandins, and their metabolites) originating from the adipocytes themselves, preadipocytes, endothelial cells, macrophages, and sympathetic nerves terminals act on human fat cells. Adenosine and prostaglandins are released by the adipocytes and various cells of the stroma vascular fraction of adipose tissue [87]. There are ﬁve different antilipolytic receptors negatively coupled to adenylyl cyclase in human fat cells: a2-adrenergic receptors (previously discussed), adenosine A1 receptors, EP3 prostaglandin E2 receptors, NPY/PYY receptors, and nicotinic acid receptors. The basal mechanisms underlying adenylyl cyclase inhibition and antilipolytic effects via inhibitory receptors seem to be similar for all the receptor classes. When the agonist interacts with the inhibitory receptor, activation of the Gi protein (i.e., the ai/bc subunits trimer) of a GTP-binding protein promotes dissociation of subunits, adenylyl cyclase inhibition and a decrease in intracellular cAMP. Adenosine and prostaglandins E2 and E1 exert potent antilipolytic effects in human fat cells. Concerning adenosine and prostaglandins, these compounds are probably metabolized very rapidly in vitro as well as in vivo [88, 89]. Nevertheless, substantial amounts of adenosine were found in the interstitial ﬂuid of adipose tissue [90]. Endogenous inhibitory agents may therefore have a stronger role on the control of triacylglycerol hydrolysis than previously suspected. Gia2 subunit appears to be the major transducer of antilipolytic responses in adipose rodent tissue, it provides a tonic suppression of basal adenylyl cyclase activity in vitro. This action is assessed by enhancement of lipolysis induced by the pertussis toxin treatment of the adipocytes in vitro [91, 92] and in vivo when using Gia2speciﬁc antisense RNA [93]. Unrestrained fat cell adenylyl cyclase proceeds at increased rates. It is suggesting that a certain degree of inhibition might be necessary for fat cell adenylyl cyclase to be susceptible to stimulation. Under in vivo conditions, in the presence of endogenous ligands, Gi-dependent inhibitory pathways may exert some permanent inhibition since the pathways driven by prostaglandins, catecholamines (i.e., a2-adrenergic receptors) and adenosine are always activated at low concentrations of the agents [78, 94]. A convincing demonstration that such mechanisms operate in vivo in human adipose tissue remains to be established. Two peptides of nervous (NPY) and gut origin (PYY) exert antilipolytic effects in human isolated fat cells. NPY/PYY receptor stimulation, via Gi-protein coupling, inhibits adenylyl cyclase activity and cAMP production [95]. Differences exist in human NPY/PYY receptor distribution according to the anatomical location of adipose tissue; the highest level of expression was found in subcutaneous adipocytes [96]. The human fat cell NPY/PYY receptor is a NPY-Y1 receptor subtype that when stimulated sustains a strong antilipolytic effect. In addition, its stimulation is also associated with a positive action on leptin secretion by human fat cells [97]. NPY is expressed by sympathetic neurons both in vivo and ex vivo, and its inhibitory effect on lipolysis has been conﬁrmed when neurons and fat cells are treated in coculture [98]. The speciﬁc circumstances under which sympathetic neurons release NPY to adipocytes in the periphery are unknown.

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Nicotinic acid has been used in the treatment of dyslipidemia for many years. These beneﬁts result from the ability of nicotinic acid to inhibit lipolysis in adipocytes and thereby reduce serum NEFA levels [99, 100]. Its effect on the inhibition of lipolysis via the activation of a Gi-protein-coupled receptor has been described in rodent adipocytes [91, 101]. Nicotinic acid and several of its derivatives exert potent antilipolytic effects in human fat cells (Lafontan et al., unpublished results). Nicotinic acid receptors were identiﬁed in rodent fat cell plasma membranes using the usual binding techniques with radiolabeled [3H]nicotinic acid [102]. A G-protein-coupled receptor that binds to nicotinic acid with the expected afﬁnity was identiﬁed by three groups. The receptor, termed GPR109A (HM74A in humans and PUMA-G in mice), is coupled to Gi-proteins and is mainly expressed in adipocytes and immune cells [103–105]. The antilipolytic effect of nicotinic acid disappeared in mice invalidated for the PUMA-G gene. D-b-Hydroxybutyrate was the ﬁrst endogenous ligand described for the PUMA-G receptor. It speciﬁcally activates the PUMA-G receptor at concentrations observed in serum during fasting in mice and its effect is abrogated in PUMA-G-deﬁcient mice [106]. Such a result supports the concept that the nicotinic acid receptor mediates the antilipolytic effects of D-b-hydroxybutyrate during starvation. However, an antilipolytic effect of D-b-hydroxybutyrate and acetoacetate was not observed in human isolated fat cells at relevant physiological concentrations (Lafontan et al., unpublished results).

9.5 Insulin: A Major Antilipolytic Agent Controlling cAMP Degradation

Insulin plays a major role in the control of NEFA disposal. It regulates the rate of lipolysis and NEFA efﬂux (i.e., inhibits lipolysis), as well as fat storage (i.e., increases the rate of resynthesis of triacylglycerols from NEFA; the re-esteriﬁcation effect) and glucose uptake by fat cell. Studies in the past 20 years have elucidated the main features of how insulin acts at the molecular level and this has been covered in a number of recent reviews [107]. Insulin is a major regulator of lipolysis; the supply of NEFAs to other tissues is rapidly and strongly inhibited by an elevation of the plasma insulin concentrations. The cellular mechanisms involved in the inhibition of lipolysis by insulin have been mainly delineated in rat fat cells; a similar mechanism is expected in human adipocytes. Insulin activates the cGMP-inhibited low-Km cAMP phosphodiesterase (PDE)-3B in rat fat cells. Insulin-induced activation of the PDE-3B is mediated via phosphorylation by the serine protein kinase Akt/protein kinase B (PKB), which is phosphorylated in response to insulin stimulation [108, 109]. PDE-3B appears to be associated with caveolae in adipocytes and this localization seems to be functionally important [110]. In fact, insulin induces formation of macromolecular complexes involved in activation of PDE-3B and its interaction with Akt/PKB. This recruitment of PDE-3B in macromolecular complexes may be critical for regulation of speciﬁc cAMP pools and signaling pathways by insulin (e.g., lipolysis) [111]. That PDE-3B is important in regulating cAMP signaling pathways, including lipolysis and insulin-induced antilipolysis, was conﬁrmed by the invalidation of PDE-3B gene in

9.6 Natriuretic Peptides Control cGMP Production, Lipolysis, and Lipid Mobilization in Humans

mice [112]. A number of circulating factors (such as TNF-a, interleukins (ILs), insulin itself, fatty acids, and glycation products) have been shown to inﬂuence the effects of insulin at the target cell level, and could lead to hyperglycemia and type 2 diabetes upon deregulation of their action [113]. Moreover, alterations in the level of expression of proteins involved in insulin signaling will also alter insulin responses in human fat cells. A low insulin receptor substrate (IRS)-1 expression in human fat cells is a marker of insulin resistance and risk for type 2 diabetes, and is associated with evidence of early vascular complications [114]. Striking adipose location-related differences, modulated by obesity, have been found in fat cell responsiveness to insulin. Insulin-induced antilipolysis and activation of NEFA re-esteriﬁcation are blunted in omental compared with subcutaneous fat cells. Various functional differences have been identiﬁed at the insulin receptor level and the postreceptor level of the insulin signaling cascade [115, 116]. Other partners of the insulin signaling cascade such as PDE-3B, responsible for the antilipolytic action of insulin in fat cells, and protein tyrosine phosphatases (PTPs), involved in the dephosphorylation of the insulin receptor, could also contribute to the modulation of insulin action. Endogenous PTP activity, including PTP-1B, is increased in omental adipose tissue and may contribute to the relative insulin resistance of this fat depot [117]. Increases in baseline systemic NEFA ﬂux have been reported in upper-body obese women. They have been partly attributed to a decreased sensitivity to the antilipolytic effect of insulin, independent of fat cell size, and to increased lipolytic rates associated with subcutaneous fat cell hypertrophy [118]. Subcutaneous abdominal adipocytes are more resistant to the antilipolytic effect of insulin than gluteal adipocytes, independently of cell size [119]. The regional heterogeneity of insulinregulated NEFA release has been conﬁrmed in vivo. Visceral adipose tissue is more resistant to insulin antilipolytic effects than leg and nonsplanchnic body fat [120]. Nevertheless, visceral fat may be a marker for, but not the source of, excess postprandial NEFA in obesity since the increased postprandial NEFA release observed in upper-body obese women and type 2 diabetics originates from the nonsplanchnic upper-body fat, not from visceral fat [121–123].

9.6 Natriuretic Peptides Control cGMP Production, Lipolysis, and Lipid Mobilization in Humans 9.6.1 Natriuretic Peptides

The natriuretic peptide family, identiﬁed for its natriuretic properties, is represented by ANP, BNP, and the C-type natriuretic peptide (CNP). ANP and BNP are essentially synthesized by atrial and ventricular cardiomyocytes, and regulate a variety of physiological events. They possess an important role in blood pressure regulation and volume homeostasis, and have natriuretic and vasodilating properties [124]. They

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also inﬂuence sympathetic nervous system (SNS) activity and the renin–angiotensin system. Most of the effects are mediated by the stimulation of cGMP production by target cells [124, 125]. The functions of ANP are not simply restricted to blood pressure homeostasis, they also seem to play an important role in the immune system [126]. Production of ANP and BNP is stimulated in pathological conditions; plasma levels of these peptides are increased in subjects with left ventricular hypertrophy, asymptomatic ventricular dysfunction, and overt heart failure [124]. NPRs either present a guanylyl cyclase activity (NPR-A and NPR-B) or not (NPR-C). NPR-A and NPR-B are the active forms of the receptors since their stimulation activates the guanylyl cyclase function of the receptor protein and the production of the second messenger, cGMP. NPR-C, which has a short cytoplasmic domain without guanylyl cyclase activity, inﬂuences plasma natriuretic peptide levels, ANP half-life, and its systemic effects [127]. ANP receptors have been identiﬁed in various tissues, including rat fat cells. NPRA have been identiﬁed in rat fat cells [128, 129] and a mild increment of intracellular cGMP was reported in rat fat cells [130]. Human adipose tissue expresses NPR-A and NPR-C mRNAs [131]. NPR-C and NPR-A protein levels and receptor function have never been explored, and no biological response had yet been reported in human fat cells when our ﬁrst studies were initiated. 9.6.2 Lipolytic Effect of Natriuretic Peptides

The original ﬁnding that was at the origin of numerous subsequent studies was the discovery of the lipolytic action of natriuretic peptides. Natriuretic peptides exert potent lipolytic effects similar to those induced by the b-adrenergic receptor agonist of reference, isoproterenol. The relative order of lipolytic potency of the peptides was ANP > BNP >> CNP (Figure 9.4a). The presence of a NPR-A receptor was revealed by the binding studies performed on human fat cell membranes using [125I]ANP as a radioligand [132]. Natriuretic peptides promoted a strong and sustained increment of intracellular cGMP in mature human adipocytes [133] as well as in adipocyte precursors differentiated into adipocytes [134]. The signal transduction pathway stimulated by ANP to promote lipolysis in human fat cells is strictly connected to an increase in intracellular cGMP concentrations. The nonhydrolyzable analog of cGMP, 8-bromo-cGMP, mimicked the lipolytic effects of ANP. Since PKA could be activated by cGMP, it was veriﬁed that ANP does not stimulate PKA activity and that inhibition of PKA by H-89 does not affect ANP-induced lipolysis. ANP-mediated lipolysis did not involve cross-talk between cGMP and PKA. It is a cGMP-dependent protein kinase (PKG), identiﬁed as cGMP-dependent kinasec (GK-I), which promotes perilipin and HSL phosphorylation, and which is at the origin of ANP-induced lipolysis. Inhibition of cGK-I by the cGMP analog 8-pCPT-cGMPS, inhibited HSL activation and lipolysis [134]. This result in isolated human fat cells conﬁrms early data in a rat cell-free system where phosphorylation and activation of HSL by a puriﬁed cGMP-dependent protein kinase was demonstrated [135, 136]. Neither the

9.6 Natriuretic Peptides Control cGMP Production, Lipolysis, and Lipid Mobilization in Humans

Figure 9.4 In vitro and in vivo effects on natriuretic peptides on isolated fat cell lipolysis and lipid mobilization in humans. (a) Comparison of the lipolytic effects of natriuretic peptides in isolated human fat cells. Values are means standard error. An asterisk indicates significantly different from basal value. (Adapted from [132].) (b) Changes in extracellular glycerol concentration and in ethanol ratio (ethanol dialysate concentration/ ethanol perfusate levels 100), which reveals changes in local blood flow, during the infusion of human ANP (10 mmol/l) in a microdialysis probe implanted in human subcutaneous

adipose tissue (Adapted from [132].) (c) Effect of intravenous infusion of human ANP (50 ng/ min/kg during 60 min) on plasma NEFA and glycerol levels (Adapted from [140].). (d) Comparison of the mean changes in extracellular glycerol concentration values in abdominal subcutaneous adipose tissue during two successive exercise bouts performed at 35% (exercise 1) and 60% (exercise 2) of VO2max and during recovery. Control microdialysis probe was perfused with Krebs–Ringer buffer. For the study of local b-adrenergic receptor blockade, a microdialysis probe was supplemented with propranolol (100 mmol/l) (Adapted from [145].).

mitogen-activated protein kinases (MAPK) inhibition (extracellular signal-regulated kinase (ERK)) nor p38 MAPK was involved in the ANP-mediated HSL phosphorylation. It was observed that the lipolytic effect of natriuretic peptides is independent of control by insulin. Insulin treatment of human fat cells has no incidence on ANPinduced lipolytic response [132, 133, 137]. The occurrence of natriuretic peptideinduced lipolysis has primate fat cell speciﬁcity. Natriuretic peptides do not stimulate lipolysis in various other species (mice, rat, dog and rabbit) [138].

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9.6.3 Induction of Lipid Mobilization by Administration of Pharmacological Doses of ANP

Infusion of human ANP in a microdialysis probe implanted in human subcutaneous adipose tissue promoted an increment in extracellular concentration of glycerol as well as vasodilatation of the vessels draining the fat deposit [132]. Both events contribute to a coordinated enhancement of lipid mobilization (Figure 9.4B). The lipid-mobilizing effect of an intravenous infusion of human ANP, as well as various metabolic and cardiovascular parameters were studied in young obese and normalweight subjects (Figure 9.4c). These in vivo studies showed that ANP is a potent lipidmobilizing hormone that acts independently of the activation of the SNS. Apparently, obesity did not modify the lipid-mobilizing effect of ANP in young obese subjects [139, 140]. Hemodynamic and metabolic response to physiologically relevant ANP concentrations was explored in healthy normal-weight young men. The increase in serum NEFA and glycerol concentrations were correlated with ANP plasma concentrations as well as glycerol increment in adipose tissue. Indirect calorimetry indicated an increase in lipid oxidation concomitantly with a decrease in carbohydrate oxidation, without changes in overall energy expenditure. ANP briskly stimulates lipid mobilization and oxidation at plasma concentrations that are usually encountered in conditions such as heart failure [141, 142]. 9.6.4 Contribution of ANP to the Physiological Control of Lipid Mobilization in Humans

Exercise-induced lipid mobilization was considered to mainly depend on SNS activation and catecholamine action in man. In addition, SNS-induced inhibition of insulin release also contributes to the lipid-mobilizing effect during exercise. A putative physiological contribution of ANP/BNP to exercise-induced lipid mobilization was hypothesized. During exercise, the SNS is activated and, concomitantly, ANP is released from the exercising heart. Strenuous endurance exercise is followed by increases in plasma BNP [143, 144]. Lipid mobilization assays were performed using in situ microdialysis in subcutaneous adipose tissue in healthy young men during two successive exercise bouts performed at 35 and 60% VO2max after placebo, local (i.e., in the microdialysis probe) or oral nonselective bantagonist treatment. In placebo-treated subjects, exercise promoted an increment in extracellular glycerol concentration. Infusion of propranolol (a nonselective bantagonist) in the microdialysis probe only partially reduced the extracellular glycerol concentration increase promoted by exercise (Figure 9.4d). It suggests a possible contribution of another lipid-mobilizing pathway. Indeed, oral b-adrenergic receptor blockade (i.e., tertatolol given per os 1 h before the beginning of exercise) did not prevent exercise-induced lipid mobilization in subcutaneous adipose tissue. Moreover, exercise-induced increase in plasma ANP was potently ampliﬁed by tertatolol administration. A positive correlation was found between extracellular glycerol concentration and plasma ANP levels but also between extracellular cGMP and extracellular glycerol concentration [145]. Since full

9.7 Other Lipolytic Pathways

blockade of fat-cell b-adrenergic receptors was veriﬁed, the existence of another partner was strengthened. The results demonstrate that exercise-induced lipid mobilization, resistant to local propranolol or oral tertatolol, is partly related to the action of ANP. Oral b-adrenergic receptor blockade promotes strong exercise-related ANP release by the heart, which explains the remaining lipid mobilization. ANP, concomitantly with the SNS, contributes to the control of lipid mobilization during exercise. Due to the lack of a suitable NPR-A receptor antagonist, usable in clinical studies, it is difﬁcult to determine the relative contribution of both pathways to the physiological control of lipid mobilization. Putative contribution of other biological compounds (parathyroid hormone (PTH), growth hormone, and IL-6) known to exert lipolytic effects in human fat cells was excluded [145].

9.7 Other Lipolytic Pathways 9.7.1 Growth Hormone

Exposure to growth hormone leads to increased plasma NEFA, ketone bodies, insulin-like growth factor-1, insulin, and glucose. Growth hormone defects in humans are associated with reduced lean body mass and increased fat mass; both can be normalized by growth hormone treatments [146]. Growth hormonedeﬁcient patients exhibit a reduction of lipolysis and plasma NEFA concentrations [147]. The physiological contribution of growth hormone to the control of human adipose tissue lipolysis and lipid mobilization has remained elusive, and is not yet entirely clear. Growth hormone receptors are not known in human fat cells; ﬁve human growth hormone receptor mRNA variants have been identiﬁed in fat cells [148]. In vitro studies have shown that growth hormone stimulates lipolysis in human adipocytes [149, 150]; the effect is delayed (2–3 h) when compared with that of catecholamines and the exact mechanism of action is not fully established. The transducing pathways are suspected to involve those used by catecholamines (i.e., cAMP- and PKA-dependent pathways). Growth hormone-dependent modiﬁcations of the interactions between adenylyl cyclase and Gia2 have been reported. Consequently, the growth hormone-related relief of Gi-dependent inhibition of cAMP production increases lipolysis [151, 152]. In rat adipocytes growth hormone increases lipolysis partly through the b-adrenergic system, including increases in both b1- and b3-adrenergic receptor function, and partly through enhanced adenylyl cyclase function, perhaps via a decrease in Gai protein expression, and increased expression of HSL [153]. Finally, in addition to its lipolytic effects, growth hormone produces a pronounced inhibition of adipose tissue lipoprotein lipase activity – the enzyme that plays the main role for hydrolyzing triglycerides in the vessels in the adipose tissue and thus for triglyceride accumulation in adipose cells.

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Pulsatile and continuous growth hormone administration, leading to plasma concentration rises seen during exercise in humans, promote a signiﬁcant increase in NEFA after 2–3 h, reﬂecting stimulation of lipolysis and ketogenesis [154, 155]. Small physiological growth hormone pulses increase interstitial glycerol concentrations in both femoral and abdominal adipose tissue [156]. Moreover, the normal nocturnal rise in plasma growth hormone concentrations also leads to site-speciﬁc regulation of lipolysis in adipose tissue [147, 157]. The lipolytic sensitivity to growth hormone shows increases during fasting [158]. The nocturnal mean peak of growth hormone preceded that of NEFA by 2 h; it is a lag-time that ﬁts with that found after a growth hormone bolus administration [159]. Growth hormone-dependent stimulation of lipolysis probably represents a physiological adaptation to stress (maintenance of basal lipolysis during fasting and exercise) [160]. Growth hormone and cortisol are cosecreted during stress conditions. It is probable that both hormones are involved in the regulation of adipose tissue metabolism during fasting and stress. Acute lipolytic effects of cortisol have been reported [161, 162]. Growth hormone and cortisol stimulate systemic and regional lipolysis independently and in an additive manner when coadministered [163, 164]. A small synthetic peptide sequence of human growth hormone (AOD-9041) has been shown to increase human and rodent fat cell lipolysis in vitro. It is efﬁcient on lipid mobilization after chronic oral administration in rodents; mechanisms of action remain to be clariﬁed in humans [165]. To conclude, growth hormone, although it possesses some delayed lipolytic and lipid-mobilizing properties, can be considered as a weaker regulator of lipolysis than catecholamines and insulin. 9.7.2 IL-6

The cytokine IL-6 is produced during muscle contraction and released into the blood [166]. The subcutaneous adipose tissue was also shown to secrete IL-6 and this secretion was correlated with the body mass index of the subjects [167]. Circulating IL-6 levels might reﬂect, at least in part, adipose tissue IL-6 production [168] and it was proposed that locally secreted IL-6 could act on adipocytes by a paracrine/autocrine mechanism. Human adipocytes express both IL-6 and its receptor system consisting of the IL-6 receptor and the signal transducing protein gp130 [169]. IL-6 stimulates lipolysis in human adipocytes [170] and exerts antiinsulin actions. It was found that IL-6-treated adipocytes have reduced insulinstimulated lipogenesis and glucose transport, and fail to maintain their adipocyte phenotype (e.g., downregulation of various adipogenic markers). Likewise, expression of insulin receptor-b and IRS-1 is reduced as is insulin-induced activation of insulin receptor-b, Akt/PKB and ERK1/2. Expression of suppressor of cytokine signaling-3, a potential inhibitor of insulin signaling, is also induced by IL-6 [171, 172]. Recombinant human IL-6 infusion, leading to plasma IL-6 concentrations of about 140 pg/ml in healthy volunteers, resulted in an increment of plasma NEFA and glycerol concentrations, and an increased rate of appearance of NEFA and glycerol measured by isotope dilution techniques. The increase in lipolysis is

9.7 Other Lipolytic Pathways

observed after more than 6 h [173]. Plasma cortisol concentrations were increased by 50%, transient changes in epinephrine were also observed during IL-6 infusion, while putative concomitant changes in growth hormone levels were not determined. It could be premature to include IL-6 inside a physiological loop of lipolytic process regulation [174]. 9.7.3 TNF-a

TNF-a is a macrophage-secreted product that has been suggested to signal the loss of body mass through the decrease in adipose tissue and muscle. TNF-a promotes hydrolysis of intracellular triacylglycerols in rodent fat cells [175]. Stimulation of lipolysis by TNF-a is not direct, since it becomes apparent only after long-lasting exposure of human and rodent adipocytes to the cytokine [176]. TNF-a stimulates triglyceride hydrolysis by multiple intracellular interferences with insulin signaling, G-protein expression, and perilipin [8]. TNF-a mechanisms of action have been explored in rodent fat cells with altered expression of TNF-a receptors (i.e., tumor necrosis factor-a receptor TNF-R1 and TNF-R2). Experiments were performed on preadipocyte cell lines established from wild-type mice (TNF-R1 þ / þ ; TNF-R2 þ / þ ) and from mice lacking TNF-R1 (TNF-R1/), TNF-R2 (TNF-R2/), or both (TNFR1/; TNFR2/) to demonstrate the role of the different TNF-a receptors in the induction of the lipolytic effects. TNF-a-induced lipolysis as well as inhibition of insulin-stimulated glucose transport are predominantly mediated by TNF-R1 [177, 178]. One important effect of TNF-a is suppression of the effect of insulin through inactivation of IRS-1 (i.e., reduction of the amount of IRS-1 protein and inhibition of its tyrosine phosphorylation). The inactivating serine-phosphorylation of IRS-1 is mediated by p42/44 MAPKs activation, at least in 3T3-L1 adipocytes [179, 180]. In human fat cells, TNF-a activates the three mammalian MAPKs in a distinct time- and concentration-dependent manner. TNF-a-induced lipolysis is mediated by only p44/42 ERKs and Jun kinase, but not by p38 kinase [181]. Blunting the endogenous tonic inhibition of lipolysis through downregulation of three Gai-protein subtypes is another possible mechanism since the amounts of Gs-protein or b-subunits of G-protein complex remain unaffected [182]. The mechanism involves proteolytic degradation mediated through the proteasome pathway [183]. Reduction of Gai coupling will raise basal lipolysis and reduce the effect of inhibiting ligands (i.e., adenosine, prostaglandins). Nevertheless, in contrast to ﬁndings in rodent cells, Gai does not appear to be essential for TNF-a-induced lipolysis in human adipocytes [184]. Experiments in rodent and human fat cells have demonstrated that TNF-a could also activate lipolysis by activating perilipin phosphorylation and decreasing its levels at the lipid droplet surface. A rather complex mechanism involves: (i) activation of ERK pathway (i.e., p42/44 MAPKs and Jun kinase) which mediates reduction of perilipin expression and facilitate lipases access to the lipid droplet [181, 184–186]; and (ii) reduction of PDE-3B expression followed by an elevated cAMP concentration leading to a concomitant increase in PKA activity and perilipin hyperphosphorylation that enables the lipases to reach lipid droplet [187].

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Although the inﬂuence of TNF-a on lipolysis is modiﬁed in obese rodents, the physiological and/or pathological importance of TNF-a signaling on lipolysis in humans remains rather debated [8]. 9.7.4 Other Lipolytic Peptides

PTH stimulates lipolysis in human fat cells [188–190]. The N-terminal 1–34 peptide portion of the hormone is responsible for adenylyl cyclase stimulation and cAMP production leading to stimulation of lipolysis. Furthermore, PTH stimulates lipid mobilization in vivo in man. Prolonged exposure to hypobaric hypoxia led to a potent reduction in lipid mobilization, through a decrease in the efﬁciency of b-adrenergic and PTH lipolytic pathways, as well as an increment in the a2-adrenergic receptormediated antilipolytic effects [150]. There appears to be no defect in the adenylate cyclase system in the fat cell in response to PTH in patients with pseudohypoparathyroidism. The effect of PTH appears at rather high PTH concentrations of the hormone which are clearly extraphysiological. There are no data concerning a physiological contribution of PTH in the control of lipid mobilization in humans. Lipolytic peptides (i.e., adrenocorticotropic hormone, a-melanocyte-stimulating hormone, lipotropin), which exert potent lipolytic effects in rodent fat cells via MC2 receptor stimulation, have no effect in human fat cells. Glucagon and glucagon-like peptide (GLP)-1, which also act in rodent fat cells, do not stimulate lipolysis in human isolated subcutaneous fat cells. Moreover, no signiﬁcant effect of either GLP-1 or glucagon on either lipolysis rate or blood ﬂow was detected in muscle or adipose tissue during local or experimental intravenous hyperglucagonemia [191, 192]. Leptin has been shown to increase glycerol release from mature adipocytes [193]. Nevertheless, leptin (within a concentration range of 25–250 ng/ml) has no direct lipolytic effect in human adipocytes either in children or adults [194]. Cachexia-inducing tumors release a lipid-mobilizing factor (LMF) that promotes release of glycerol when incubated with murine adipocytes. Induction of lipolysis by LMF was associated with an increase in intracellular cAMP levels. The serum and urine of cachectic cancer patients contain LMF, the activity of which is correlated with the extent of weight loss [195]. Zinc-a2-glycoprotein (ZAG), a protein of 43 kDa, acts as a LMF to stimulate lipolysis in adipocytes, leading to cachexia in mice implanted with ZAG-producing tumors. ZAG was detected in the major fat deposits of mice and in 3T3-L1 adipocytes. ZAG gene expression and protein were also found in human fat cells (visceral adipose tissue and subcutaneous adipose tissue). Murine and human ZAG share up to 100% identity in speciﬁc regions hypothesized to be important in lipid metabolism [196]. ZAG is a new adipose tissue protein factor that may be involved in the modulation of lipolysis in adipocytes [197]. Human adipocytes (i.e., originating from the SGBS human cell line) express and secrete ZAG, with ZAG expression being regulated particularly through TNF-a and the peroxisome proliferator-activated receptor-c nuclear receptor [198]. Nevertheless, its physiological role remains to be established in human fat cells and in human physiology.

9.8 Future Trends and Pharmacological Prospects

9.8 Future Trends and Pharmacological Prospects

To conclude, there are only two hormone systems, namely catecholamines and natriuretic peptides, that have acute stimulatory effects on lipolysis and fat mobilization in humans. Adipose tissue (e.g., fat cells and the vascular bed of adipose tissue) is a new and unsuspected target organ of natriuretic peptides. Two strictly independent pathways control cAMP and cGMP production, respectively. Induction of lipolysis and lipid mobilization must be now included in the numerous physiological actions of natriuretic peptides. Further fundamental and clinical studies will be required to answer the various questions raised by the discovery of the metabolic effects of natriuretic peptides. The effect of b-adrenergic antagonist treatment on natriuretic peptide release merits further study. Finally, all the drugs known to modulate natriuretic peptide-dependent pathways should be evaluated for their putative metabolic side-effects when given for the management of cardiovascular diseases. Deregulation of lipid metabolism has long been recognized as essential in the development of obesity and the metabolic syndrome. Lipolysis plays a pivotal role in controlling the quantity of triglycerides stored in fat depots and in determining plasma NEFA levels. Critical steps in this catabolic process constitute targets for strategies to ﬁght against obesity and to improve the poor metabolic proﬁle of patients with the metabolic syndrome. Activators of lipolysis could present a pharmacological interest to induce mobilization in fat deposits that are resistant to hormone-induced lipolysis. However, they must be associated with agents that stimulate the oxidation of fatty acids by skeletal muscle and energy expenditure. Concomitant stimulation of fat oxidation in skeletal muscle is an important alternative strategy. The blockade of a2adrenergic receptors has been considered for the enhancement of lipid mobilization. Administration of a a2-adrenergic receptor antagonist promotes SNS activation and blocks cell a2-adrenergic receptors in fat cells and other cells expressing this adrenergic receptor subtype. A strong lipid mobilization is induced by their blockade. Since the launch of nicotinic acid (niacin) as a lipid-lowering drug some time ago, the suppression of lipolysis to decrease free fatty acid levels has attracted much interest. The recent cloning of the receptor for nicotinic acid that is mainly expressed in adipose tissue has undoubtedly led to important screening efforts to identify agonists with fewer side-effects than niacin and its long-lasting form, acipimox. Inhibition of HSL is also attractive as the enzyme has little homology with other mammalian lipases and shows a rather limited tissue distribution. The effect of chronic treatment with these in rodent models of obesity and dyslipidemia is now awaited.

Acknowledgments

The author thanks all the past and present members of the laboratory (INSERM U317 and 586) for their contribution to research. Thanks go out more speciﬁcally to Drs. M.

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Berlan, C. Carpene, P. Mauriege, J. Galitzky, P. Valet, J.-S. Saulnier-Blache, G. Tavernier, A. Bouloumie, I. Castan, D. Langin, C. Sengenes, C. Moro, F. Crampes, I. De Glisezinski, I. Harant, and V. Stich who contributed to the described studies. Studies in the authors laboratory are supported by INSERM French National Institute for Medical Research and funds from Projects of the Fifth and Sixth Framework Programs of European Commission. The author apologizes to many colleagues for not being able to cite all relevant references because of space limitations.

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2 inhibitors in 3T3-L1 adipocytes. Mol Endocrinol., 14, 1557–1569. Fujishiro, M., Gotoh, Y., Katagiri, H., Sakoda, H., Ogihara, T., Anai, M., Onishi, Y. Ono, H. et al. (2003) Three mitogenactivated protein kinases inhibit insulin signaling by different mechanisms in 3T3-L1 adipocytes. Mol Endocrinol., 17, 487–497. Ryden, M., Dicker, A., Harmelen, V.V., Hauner, H., Brunnberg, M., Perbeck, L., L€ onnqvist, F., and Arner, P. (2002) Mapping of early signaling events in tumor necrosis-alpha-mediated lipolysis in human fat cells. J. Biol. Chem., 277, 1085–1091. Gasic, S. and Green, A. (1995) Gi Downregulation and heterologous desensitization in adipocytes after treatment with the a2-agonist UK14304. Biochem. Pharmacol., 49, 785–790. Botion, L.M., Brasier, A.R., Tian, B., Udupi, V., and Green, A. (2001) Inhibition of proteasome activity blocks the ability of TNF alpha to down-regulate Gi proteins and stimulate lipolysis. Endocrinology, 142, 5069–5075. Ryden, M., Arvidsson, E., Blomqvist, L., Perbeck, L., Dicker, A., and Arner, P. (2004) Targets for TNF-alpha-induced lipolysis in human adipocytes. Biochem. Biophys. Res. Commun., 318, 168–175. Souza, S.C., Vargas, L.M.d., Yamamoto, M.T., Lien, P., Franciosa, M.D., Moss, L.G., and Greenberg, A.S. (1998) Overexpression of perilipin A and B blocks the ability of tumor necrosis factora to increase lipolysis in 3T3-L1 adipocytes. J. Biol. Chem., 273, 24665–24669. Souza, S.C., Palmer, H.J., Kang, Y.-H., Yamamoto, M.T., Muliro, K.V., Paulson, K.E., and Greenberg, A.S. (2003) TNF-a induction of lipolysis is mediated through activation of the extracellular signal related kinase pathway in 3T3-L1 adipocytes. J. Cell. Biochem., 89, 1077–1086. Zhang, H.H., Halbleib, M., Ahmad, F., Manganiello, V.C., and Greenberg, A.S. (2002) Tumor necrosis factor-a stimulates lipolysis in differentiated

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Part Three Endocrine Functions of Adipose Tissue

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10 Leptin Secretion and Action Robert V. Considine 10.1 Introduction

As discussed in several chapters in this book, adipose tissue is an active endocrine organ releasing a varied array of metabolites, hormones, cytokines, and other factors. Although the potential for adipose tissue to act in an endocrine manner to regulate energy intake was suggested by Kennedy over 65 years ago [1], it was the discovery of leptin synthesis and release [2] that focused and accelerated investigation into the endocrine function of adipose tissue. Shortly following the discovery of adipose tissue leptin synthesis, a number of laboratories demonstrated that recombinant leptin inhibited food intake and caused weight loss in mice (reviewed in detail in [3]). A receptor for leptin was found in the hypothalamus and a point mutation within the gene implicated as the defect resulting in obesity in db/db mice [4, 5]. Taken together, these observations, as well as numerous other related ﬁndings, have established a major role for leptin in regulation of body weight. Further, studies to understand the central neural pathways responsive to leptin have provided extensive insight into the hypothalamic regulation of energy intake and energy expenditure. Defects in leptin synthesis and signaling such as found in ob/ob and db/db mice are rarely the cause of obesity in humans [6]. In contrast, human obesity is characterized by elevated serum leptin levels (hyperleptinemia) and resistance to the appetitesuppressing effect of leptin. Evidence suggests that hyperleptinemia contributes to obesity-related hypertension and other comorbidities. This chapter reviews the synthesis of leptin and its actions in the central nervous system. The concept of leptin resistance is discussed, as is evidence for the contribution of selective leptin resistance to development of comorbidities in obesity. The use of leptin to induce weight loss or to ameliorate the metabolic abnormalities associated with lipodystrophy are also reviewed.

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Figure 10.1 Serum leptin is determined by adiposity and is significantly reduced with weight loss. Serum leptin levels were measured in 10 women and 10 men before and 6 months after Roux-en-Y bariatric surgery. BMI was

significantly decreased 33.1 2.3% (56.3 to 36.1 kg/m2) in the men and 32.3 1.6% (46.4 1.0 to 8.9 2.8 kg/m2) in the women 6 months postsurgery.

10.2 Leptin Synthesis

Circulating leptin concentrations are highly positively correlated with adiposity, measured as body mass index (BMI), percent body fat, or fat mass (kg), in humans and also in rodent models [7]. As a consequence of the strong correlation between body fat and serum leptin, changes in adiposity result in changes in serum leptin. As shown in Figure 10.1, the reduction in fat mass following bariatric surgery is associated with a signiﬁcant decrease in serum leptin. An increase in adipose tissue during weight gain results in greater serum leptin [8]. These observations provide support for the concept that leptin is a signal of the size of energy stores in the body. Despite the strong correlation between serum leptin and fat mass in general population studies there is a considerable degree of variation in serum leptin at any given fat mass. Factors that contribute to the variation in serum leptin are gender, differential distribution of adipose tissue into the visceral or subcutaneous depots, and insulin/glucose metabolism. 10.2.1 Gender and Body Fat Distribution Determine Serum Leptin

Women have signiﬁcantly higher serum leptin concentrations than men with an equivalent amount of body fat [9]. One factor contributing to this difference is the deposition of body fat into different adipose tissue depots. Women have greater amounts of subcutaneous adipose tissue than men, who tend to have greater visceral

10.2 Leptin Synthesis

adipose tissue mass, particularly when obese. Leptin gene (LEP) expression and leptin secretion are greater from subcutaneous than visceral adipocytes (reviewed in [7]), thus more leptin is released into the blood from the greater subcutaneous adipose tissue mass in women. Gondal steroids also contribute to greater serum leptin in women. In vivo and in vitro studies have suggested that testosterone and estrogens can regulate leptin synthesis and release from adipose tissue independently of their effects to inﬂuence deposition of adipose tissue into the subcutaneous or visceral depots. In general, androgens reduce, and estrogens increase, leptin synthesis in adipose tissue [9]. A suppressive effect of testosterone on serum leptin is supported by studies of pubertal development in males, in whom serum leptin levels decrease as testosterone increases [10]. Further, administration of testosterone lowers serum leptin in hypogonadal men [11, 12] and in female-to-male transsexuals [13]. In vitro, exposure to dihydrotestosterone for 48 h decreases leptin secretion from visceral adipose tissue pieces [14, 15]. Evidence for a stimulatory effect of estrogen on serum leptin are observations that estrogen therapy (along with antiandrogens) increases leptin in male-to-female transsexuals [13] and that estradiol stimulates leptin secretion from visceral adipose tissue pieces in vitro [14, 15]. The mechanism(s) through which gonadal steroids directly regulate leptin synthesis and release from adipocytes remain to be fully elucidated. 10.2.2 Caloric Intake, Insulin, and Glucose Influence Serum Leptin

Serum leptin measured in the morning following an overnight fast tends to be fairly constant from day to day. However, over a 24-h period serum leptin exhibits a diurnal pattern with a peak at around 0200 h in both lean and obese individuals [16, 17]. If meal times are shifted by 6 h such that dinner is eaten at 2400 h instead of 1800 h, the peak in leptin is also shifted by 6 h. Further, complete day/night reversal results in peak leptin occurring at 1400 h [17]. These observations suggest that the leptin peak is entrained to food intake, and the requisite increase in insulin and glucose that accompanies eating. Additional evidence for insulin and glucose regulating serum leptin is provided by the ﬁnding that hyperinsulinemic/ euglycemic clamps of differing duration increase serum leptin [18, 19] and that feeding high fat/low carbohydrate meals, which result in smaller postprandial excursions in insulin and glucose than meals of standard carbohydrate content, reduce serum leptin [20]. Hexosamine (UDP-GlcNAc) biosynthesis in adipocytes has been suggested as a mechanism through which insulin and glucose regulate leptin synthesis. An increase in adipose tissue UDP-GlcNAc with infusion of glucosamine, uridine, or free fatty acids during a hyperinsulinemic/euglycemic clamp in rats was associated with an increase in serum leptin [21]. Transgenic mice with elevated adipose tissue hexosamine biosynthesis due to overexpression of the rate-limiting enzyme in UDP-GlcNAc biosynthesis have elevated Lep mRNA and serum leptin [22]. In obese humans subcutaneous adipose tissue UDP-GlcNAc is elevated 3.2-fold, and a

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signiﬁcant positive correlation between BMI and adipose tissue UDP-GlcNAc has been reported [23]. Further, increasing UDP-GlcNAc synthesis in human subcutaneous adipocytes in vitro stimulates leptin release and reducing UDP-GlcNAc decreases leptin secretion [23]. As described above, serum leptin measured after an overnight fast is fairly constant from day to day. However, a rapid decrease in serum leptin levels with short-term fasting (24–72 h) has been observed in both animals [24] and humans [25–27]. Leptin levels rise again with refeeding, and the fall in leptin can be blocked by maintenance of blood sugar at approximately 90 mg/dl during the fasting period [25, 26]. The rapid fall in leptin with fasting is disproportionately greater than the small reduction in adipose tissue mass that occurs over the same time period. Thus, it has been suggested that the fall in leptin with fasting is a signal to the central nervous system that coordinates the hypothalamic neuroendocrine response to caloric deprivation [28]. Evidence that leptin coordinates the neuroendocrine response to fasting was originally derived through replacement experiments in rodents [24]. In 48-h fasted mice, thyroid and reproductive hormone levels are reduced, and glucocortiocoids increased. These neuroendocrine adaptations occur to promote survival by reducing energy expenditure and managing the stress of caloric deprivation. Preventing the starvation-induced fall in leptin, by administration of recombinant leptin to achieve levels similar to the fed state, substantially blunted the reduction in gonadal and thyroid hormones, and attenuated the increase in glucocorticoids. Chan et al. [27] have demonstrated that replacement of leptin during complete caloric restriction in men can prevent the fasting-induced reduction in testosterone and partially prevent the suppression of the hypothalamic–pituitary–thyroid axis. These observations establish a role for leptin in regulating hypothalamic–pituitary function in response to caloric restriction, in addition to leptins role as a marker of energy stores in the fed state. 10.2.3 Transcriptional Regulation of Leptin Synthesis in Adipocytes

LEP mRNA is greater in large adipocytes than in small adipocytes isolated from the same piece of adipose tissue and leptin secretion strongly correlates with fat cell volume [29–31]. Thus, elevated leptin levels in obese individuals result from both greater secretion from larger adipocytes and an increased number of adipocytes. Hexosamine biosynthesis may be one link between adipocyte size and leptin synthesis [23]. Many hormones and cytokines also regulate leptin synthesis when tested in isolation. However, the effects of tumor necrosis factor (TNF)-a and cortisol have been the best studied and appear to be relevant in vivo. TNF-a, which is elevated in the adipose tissue of obese subjects, inhibits leptin release from cultured rodent and human adipocytes [32, 33]. Cortisol stimulates leptin synthesis and secretion from adipocytes in vivo and in vitro (e.g., [34, 35]). Synthesis of cortisol by 11b-hydroxysteroid dehydrogenase in the adipose tissue itself [36] likely regulates leptin secretion in a paracrine manner.

10.4 Leptin Action in the Central Nervous System

10.3 Leptin Receptors

The leptin receptor is present in almost all tissues examined. The best-studied isoform, the hypothalamic leptin receptor Ob-Rb, is a class I cytokine receptor. Leptin binding to Ob-Rb promotes Janus kinase (JAK)-dependent signaling through signal transducer and activator of transcription (STAT) proteins, primarily STAT-3. This leptin receptor has also been observed to activate phosphoinositol-3-kinase and phosphodiesterase 3B signaling pathways in the hypothalamus, although the role of JAK and STAT proteins in this process is not fully understood [37–39]. Ob-Ra (originally termed the short leptin receptor) is the other major isoform of leptin receptor, which is derived by alternative splicing of the LEPR gene. The extracellular domains of Ob-Ra and Ob-Rb are identical; however, the intracellular domain of Ob-Ra is truncated and lacks the structural motifs to initiate JAK–STAT signaling. Ob-Ra is highly expressed in cerebral microvessels comprising the blood–brain barrier where it functions to transport leptin from the blood to the brain [40]. Expression of Ob-Rb mRNA is greatest in the hypothalamus and strong evidence supports the concept that this receptor isoform signals for leptin action within the central nervous system [41]. In contrast, expression of Ob-Ra is more readily detectable than is Ob-Rb in non-neural tissue. This signiﬁcant difference in expression levels must be taken into consideration when interpreting effects of leptin elicited by direct interaction of the hormone with tissues and cells. However, based on observations in leptin receptor transfected cells, it appears that limited expression of Ob-Rb is sufﬁcient to provide competent leptin signaling [42].

10.4 Leptin Action in the Central Nervous System

Within the hypothalamus Ob-Rb is highly expressed in the arcuate, paraventricular, ventromedial, and dorsomedial nuclei [41, 43]. Leptin signaling in the hypothalamus is primarily mediated by two leptin-responsive populations of neurons present in the arcuate nucleus (Figure 10.2). One group of neurons express pro-opiomelanocortin (POMC), which is processed to a-melanocyte-stimulating hormone (MSH), and cocaine- and amphetamine-regulated transcript (CART). a-MSH and CART inhibit food intake and promote weight loss in a leptin-responsive manner. The second population of neurons contain both neuropeptide Y (NPY) and agouti-related peptide (AGRP). NPY and AGRP stimulate feeding and are inhibited by leptin [43]. The leptin-responsive neurons present in the arcuate nucleus innervate other hypothalamic areas, including the paraventricular nucleus (PVN), perifornical area (PFA), and lateral hypothalamic area (LHA), which contain secondary neurons that express different neuropeptides that regulate food intake [44]. The presence of leptin receptors in the PVN, PFA, and LHA suggests that leptin may regulate food intake or other processes through mechanisms initiated independent of the arcuate

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Cerebral Cortex

PVN DMH

Parasympathetic VMH ARC

NPY AGRP

LHA

and Sympathetic

αMSH

PBN

+

LEPTIN

ME NTS

LEPTIN

Pituitary Stalk

LEPTIN

Figure 10.2 Schematic drawing of the hypothalamus and brainstem in sagittal section illustrating leptin signal pathways. Leptin enters the hypothalamus through the median eminence (ME), but can also access other areas of the brain via transport across the blood–brain barrier. In the arcuate nucleus (ARC) leptin activates neurons containing a-MSH and CART, which inhibit feeding. Leptin also binds to arcuate neurons expressing NPY and AGRP, and inhibits release of these orexigenic peptide neurotransmitters. Leptin-responsive neurons in the arcuate nucleus project to the PVN and the LHA for integration with other neural signals. The effects of leptin signaling in the

hypothalamus are then relayed to the body via parasympathetic and sympathetic neurons. Hypothalamic leptin receptors (black semicircle) are also present in the dorsomedial (DMH) and ventromedial (VMH) nuclei, PVN, and LHA. Leptin receptors have also been detected in hindbrain structures such as the NTS, which receives chemosensory and mechanosensory input from the gut about meal size. Leptin may regulate NTS activity to influence meal size via descending connections from the hypothalamus or by directly binding to its receptor in this nucleus. (Figure modified with permission from [41].)

nucleus. In support of this concept, POMC neuron-speciﬁc leptin receptor knockout mice exhibit only modest increases in body weight [45], suggesting that leptin receptors located on neurons other than arcuate POMC neurons are important in body weight homeostasis. Gastrointestinal signals (mechanosensory, chemosesory, and hormonal) are the principal determinants of meal size. These neural signals are ﬁrst processed in the nucleus of the solitary tract (NTS) and parabrachial nucleus located in the dorsal medulla [46]. Recently, investigators have begun to focus in detail on the ability of leptin to regulate neuronal activity in the NTS. Reconstitution of viable leptin receptors in the arcuate nucleus of leptin receptor-deﬁcient Koletsky rats suggests that leptin acts in the hypothalamus to regulate NTS activity via descending projections [47]. However, leptin receptors have been detected in the NTS and injection of leptin directly into this nucleus inhibits feeding [48], suggesting leptin may directly signal in the hindbrain. Leptin could access the NTS from the blood via the

10.5 Leptin Resistance in Obesity

fenestrated capillaries of the adjacent area postrema [49]. Further work is necessary to fully understand leptin signaling in the hindbrain. In addition to reducing food intake, leptin has also been shown to increase energy expenditure via hypothalamic mechanisms. Leptin-induced weight loss in ob/ob mice results from an increase in oxygen consumption, body temperature, and locomotor activity, in addition to the reduction in food intake [50]. There is little evidence that leptin alters locomotor activity or increases body temperature when administered to lean or obese mice in which leptin synthesis is not genetically blocked. However, leptin administration to lean and obese rodents does increase sympathetic nerve activity to thermogenic brown adipose tissue, as well as nonthermogenic tissues including the kidney and adrenals [51]. Leptin activation of the sympathetic nervous system likely has a pathologic role in obesity-related hypertension, as discussed in detail in Section 10.6.1.

10.5 Leptin Resistance in Obesity

Mutations in both leptin [2] and the leptin receptor [4, 5] result in obesity in rodent models. However, inactivating mutations in the human LEP or LEPR gene causing obesity are extremely rare and have been identiﬁed in only a handful of families [6]. Rather, serum leptin levels are elevated in obese individuals due to the greater amount of adipose tissue. Thus, the concept of leptin resistance, in analogy to insulin resistance, was proposed to explain the observation that elevated leptin levels did not reduce food intake or prevent body weight gain in obese humans [3]. Several molecular possibilities have been put forth to explain the mechanism of leptin resistance. In rodents, consumption of a high-fat diet decreases the weight loss response to peripherally injected leptin by impairing Ob-Ra-mediated transport of leptin across the blood–brain barrier [52, 53]. Although defective leptin transport occurs in this model, the extent to which this defect contributes to leptin resistance is not entirely clear considering that the arcuate nucleus is located very near the median eminence where Ob-Ra-facilitated leptin transport may not be necessary for leptin access. A second mechanism for leptin resistance involves suppressors of cytokine signaling (SOCS) – a group of early genes activated by the JAK–STAT signal transduction pathway that act in a negative feedback loop to limit cytokine signaling. SOCS-3 is a potent inhibitor of leptin receptor-initiated JAK–STAT signaling in cultured cell lines and SOCS-3 mRNA in the hypothalamus is increased by leptin treatment [54]. Agouti mice are resistant to the appetite curbing effects of central leptin administration and SOCS-3 mRNA is elevated in leptin-responsive arcuate neurons in these mice, suggesting that increased SOCS-3 expression is a cause of leptin resistance in these animals [54]. Of importance, it appears that leptin resistance can exhibit a distinct anatomic distribution. Defective leptin-stimulated STAT3 phosphorylation can be demonstrated in the arcuate nucleus at the same time that other hypothalamic and extrahypothalamic sites (see Figure 10.1) remain leptinsensitive [55]. The selective anatomic distribution of resistance to leptin signaling in

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the hypothalamus is of particular importance in understanding the effects of leptin on blood pressure. A third molecular mechanism for leptin resistance may involve the activity of protein tyrosine phosphatase (PTP)-1B. In Cos-7 cells expressing leptin receptor, coexpression of PTP-1B inhibits tyrosyl phosphorylation of JAK-2 and STAT-3. In contrast, ﬁbroblasts engineered to express leptin receptor but lacking PTP-1B have enhanced leptin signaling [56]. PTP-1B knockout animals are resistant to obesity and exhibit enhanced leptin-induced hypothalamic STAT-3 phosphorylation [56, 57], suggesting that PTP1-B can regulate neuronal leptin signaling in vivo.

10.6 Metabolic Complications of Hyperleptinemia in Obesity

As discussed above, the concept of leptin resistance was originally proposed to explain the observation that elevated leptin did not reduce food intake or prevent body weight gain in obese humans. However, it is now recognized that resistance to leptin action may be primarily centered within the neuronal pathways regulating food intake and that other leptin-responsive processes remain intact in the obese state. Thus, hyperleptinemia resulting from leptin resistance within pathways regulating energy intake can have pathologic effects on other regulatory systems and contribute to the metabolic complications of obesity. 10.6.1 Leptin and Obesity-Related Hypertension

Leptin promotes weight loss in rodents via sympathetic activation of brown adipose tissue and oxidation of fatty acids. Leptin signaling in the central nervous system also increases the activity of renal, lumbar, hindlimb, and adrenal gland sympathetic nerves. Sympathoactivation with either intracerebroventricular or chronic intravenous leptin administration has been demonstrated to increase mean arterial blood pressure and heart rate in a number of rodent studies [51]. Furthermore, arterial blood pressure in leptin-deﬁcient ob/ob mice, which is lower than in wild-type littermates, is substantially increased with leptin treatment. In contrast, obese hyperleptinemic agouti mice and transgenic skinny hepatic-leptin-overexpressing mice both have elevated blood pressure [58]. Although inhibition of food intake by leptin is impaired in obese agouti mice, the leptin-induced increase in renal sympathetic nerve activity is not different in agouti mice and their lean littermates [59, 60]. Finally, in high-fat-fed mice the effect of intraperitoneal or intracerebroventricular leptin to decrease food intake and body weight was impaired, but leptin-induced renal sympathetic nerve activity was preserved [61]. Taken together, these observations demonstrate that leptin regulates blood pressure via sympathetic activation and that hyperleptinemia in obesity can result in hypertension. Further, leptin regulation of blood pressure via stimulation of renal sympathetic nerve activity appears to be

10.6 Metabolic Complications of Hyperleptinemia in Obesity

intact in the presence of resistance to the weight reducing effects of the hormone, supporting the concept of selective central leptin resistance in obesity. Leptin regulates sympathetic nerve activity via a-MSH and the melanocortin-4 receptor (MC-4R). The MC-4R receptor antagonist SHU-9119 blocks both acute and chronic effects of leptin to stimulate renal sympathetic nerve activity and heart rate [62–65]. However, MC-4R antagonism has also been shown to block leptin effects on food intake and body weight [62, 66], implying that the same neural pathway mediates both the metabolic and sympathetic effects of leptin. How can such an observation be reconciled with the ﬁnding of leptin resistance in metabolic pathways but leptin sensitivity in sympathetic pathways? One possibility is that resistance to the metabolic effects of leptin is not mediated by melanocortin signaling. As discussed in Section 10.4, POMC neuron-speciﬁc leptin receptor knockout only results in mild obesity, implicating other neural pathways in leptin regulation of food intake, possibly where leptin resistance could be focused. Alternatively, it is possible that leptin engages MC-4R signaling for both metabolic and sympathetic effects, but that the anatomic location of the neurons mediating each response is different. With intracerebroventricular administration of SHU9119 into the third ventricle it is not possible to determine if antagonism of MC-4R function blocks the sympathetic or metabolic responses to leptin via the same hypothalamic nuclei. In support of the idea that other hypothalamic nuclei may be engaged by leptin, a recent study has shown that injection of leptin into the ventromedial and dorsomedial hypothalamus can increase renal sympathetic activity, arterial pressure, and heart rate [67]. These hypothalamic nuclei have previously been shown to express normal leptin sensitivity in the presence of leptin resistance in the arcuate nucleus [55]. Evidence against the hypothesis of separate nuclei for the metabolic and sympathetic effects of leptin is provided by a recent report demonstrating that injection of leptin directly into the arcuate nucleus of normal rats increased renal sympathetic nerve activity [68]. While illustrating the central role of the arcuate nucleus in the metabolic and sympathetic effects of leptin, this study did not directly address the contribution of other nuclei in mediating the sympathetic effects of leptin in obese leptin-resistant animals. Thus, the exact mechanism(s) that provide for selective leptin resistance and maintenance of leptin-derived sympathetic effects remain to be fully elucidated. 10.6.2 Other Possible Pathologic Effects of Leptin

Elevated leptin levels in obese humans have been suggested to increase risk of thrombosis, and to contribute to key aspects of atherogenesis. However, as recently reviewed [69], these consequences of hyperleptinemia are highly debated with many studies ﬁnding no effect of leptin on thrombosis and atherogenesis. Additional work, with careful attention to the concentration of leptin used, will be needed to determine if leptin has a role in the pathophysiology of these vascular complications in obesity.

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10.7 Leptin Therapy in Humans

Early observations that leptin treatment resulted in substantial weight loss in rodents suggested that the hormone might be useful in treating human obesity. However, the ability of leptin to mediate weight loss in obese, leptin-resistant humans has been modest. In contrast, leptin has been found to be a useful therapeutic in states of human leptin deﬁciency. 10.7.1 Leptin, Weight Loss, and Human Obesity

Two clinical trials have been conducted to test the therapeutic potential of recombinant leptin in treating common obesity. In the ﬁrst trial daily leptin administration resulted in modest weight loss after 24 weeks of treatment in a group of lean and obese subjects [70]. Weight loss was highly variable by subject, but in general higher doses of recombinant leptin caused greater weight loss. In a second trial of obese men only, pegylated recombinant leptin caused a reduction in appetite, but no signiﬁcant decrease in body weight with 12 weeks of therapy [71]. An important consideration in this second study is that serum leptin levels were only elevated at two timepoints over the 12-week treatment period, suggesting that weight loss was not achieved because the leptin dose was not high enough. However, taken together, these two studies suggest that leptin can stimulate weight loss in humans if a sufﬁcient dose can be given that does not evoke signiﬁcant side-effects. Structural analogs or modiﬁcations to the hormone may lead to an improved pharmacology with regard to weight loss in generalized obesity. 10.7.2 Congenital Leptin Deficiency

Defects in the LEP gene resulting in human obesity have been identiﬁed in ﬁve different kindreds. The ﬁrst two cases to be identiﬁed, an 8-year-old girl weighing 86 kg and a 2-year-old boy weighing 29 kg, were cousins in a highly consanguineous Pakastani family [72]. The LEP gene in these children contained a homozygous frameshift mutation resulting from deletion of a single guanine nucleotide in codon 133, resulting in a truncated leptin protein that is not secreted [73]. This same mutation was subsequently found in four other individuals from three different unrelated families of Pakistani origin [74]. Leptin therapy in three of the children with congenital leptin deﬁciency has resulted in signiﬁcant and sustained loss of body fat. Beneﬁcial effects on appetite, hyperinsulinemia, hyperlipidemia, and immune cell number have also been observed. Leptin therapy also facilitated appropriately timed pubertal development [74]. Leptin deﬁciency has also been described in four members of a Turkish kindred [75, 76]. In this family a C ! Tsubstitution, changing arginine at position 105 to

10.7 Leptin Therapy in Humans

a tryptophan, impairs leptin secretion. All subjects were hyperphagic and severely obese. In addition, the three adults had hypothalamic hypogonadism. Leptin treatment in the three adults resulted in signiﬁcant weight loss and, in the one subject with diabetes, an improvement in hyperinsulinemia [77]. Overall, these ﬁndings strongly support a role for leptin to reduce obesity and improve metabolism in individuals with congenital leptin deﬁciency. 10.7.3 Lipodystrophic Leptin Deficiency

Lipodystrophy, a partial or complete loss of adipose tissue, results in hyperphagia, insulin resistance, diabetes, hypertriglyceridemia, nonalcoholic hepatic steatosis, and low leptin levels [78]. Leptin replacement therapy in lipodystrophic animal models improved a number of the metabolic abnormalities resulting from the lack of adipose tissue [79, 80], supporting its use in humans. Leptin administration to lipodystrophic patients improves insulin resistance, lowers hemoglobin A1c, and decreases fasting serum triglycerides [81, 82]. Liver and muscle lipid content is signiﬁcantly reduced with leptin treatment [82–84] as are semiquantitative scores of nonalcoholic steatohepatitis [85]. Animal studies suggest that leptin stimulates fatty acid oxidation in liver and muscle, thus improving insulin sensitivity of these tissues [86, 87]. Highly active antiretroviral therapy (HAART) for HIV-1 infection has been associated with insulin resistance, hyperlipidemia, and lipodystrophy [88]. Lee et al. hypothesized that the relative leptin deﬁciency in lipodystrophic HAARTtreated subjects might contribute to their insulin resistance and other metabolic abnormalities [89]. Leptin therapy in seven HIV þ men with HAART-induced lipoatrophy improved insulin resistance and high-density lipoprotein. Body weight and fat mass decreased with treatment due to a reduction in central trunk fat. Leptin may therefore be of use in treating the metabolic abnormalities associated with HAART treatment. 10.7.4 Hypothalamic Amenorrhea

Functional hypothalamic amenorrhea occurs when a relative energy deﬁcit due to weight loss, excessive exercise, or eating disorders disrupts the secretion of hypothalamic gonadotropin-releasing hormone, resulting in the absence of menstrual cycles, low estrogen levels, and low or normal levels of gonadotropins [90]. Due to the limited amount of adipose tissue, leptin levels are low in women with hypothalamic amenorrhea [91]. In eight women with hypothalamic amenorrhea leptin treatment increased mean luteinizing hormone levels, luteinizing hormone pulse frequency, and estradiol, and improved a number of measures of ovarian function [92]. Leptin therapy also increased insulin-like growth factor-1, insulin-like growth factor-binding protein-3, bone alkaline phosphatase, and osteocalcin, suggesting that leptin had beneﬁcial effects on bone metabolism in these subjects.

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10.8 Conclusions

Leptin is synthesized and released into the circulation in proportion to the amount of body fat, gender, and distribution of body fat into the subcutaneous or visceral depots. Elevated serum leptin in obesity results from the greater fat mass in the body and greater leptin synthesis in adipocytes of obese subjects. Adipocyte size, hexosamine biosynthesis mediated by insulin-stimulated glucose uptake, and elevated production of cortisol in adipose tissue all contribute to the increase in leptin synthesis by adipocytes in obesity. Leptin acts in the brain to regulate food intake and energy expenditure. Hypothalamic leptin signal pathways are the best understood at present, but intriguing work on leptin action in the hindbrain is now emerging. Resistance to leptin in the hypothalamus has been suggested to explain the observation that elevated leptin levels in obesity do not reduce food intake or body weight. However, leptin resistance in the central nervous system appears to be selective, with leptininduced activation of the sympathetic nervous system intact and contributing to hypertension in obese subjects. Recombinant leptin administration is an effective therapeutic for the metabolic complications of congenital or relative hypoleptinemia and leptin has some limited efﬁcacy as an antiobesity agent in subjects with general obesity. Future work is needed to better understand the effect of leptin to regulate the activity of the NTS to modify meal size. Administration of leptin in combination with gut hormones (glucagon-like peptide-1, peptide YY, and amylin) may prove to be more effective in mediating weight loss than using leptin alone.

Acknowledgments

Work of the authors cited in this chapter was supported in part by grants from the NIH (DK51 140 and M01 RR00 750 (Indiana University General Clinical Research Center)), the American Diabetes Association, and the Showalter Trust.

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11 Adiponectin Jonathan P. Whitehead and Ayanthi A. Richards 11.1 Introduction

Adipocytes secrete a plethora of bioactive molecules collectively termed adipokines. Adiponectin is the most abundant of these and circulates at high concentrations, around 5–30 mg/ml in humans. It has an admirable range of properties, including insulin-sensitizing, anti-inﬂammatory, antiatherogenic, and anticancer, and is also cardioprotective. Unlike most other adipokines, circulating levels of adiponectin are reduced in obesity and associated diseases, making it an attractive therapeutic target [1]. In support of its therapeutic potential, administration of recombinant adiponectin ameliorates metabolic complications in mice [2, 3], whilst the beneﬁcial effects of the insulin-sensitizing thiazolidinediones (TZDs) in humans are at least partly due to the improvement in adiponectin proﬁles [4]. Adiponectin, also termed Acrp30 (adipocyte complement-related protein of 30 kDa), ADIPOQ, apM1 (adipose most abundant gene transcript-1), or GBP28 (gelatin-binding protein of 28 kDa), was originally identiﬁed by four groups [5–8]. Its expression is largely restricted to adipocytes and is induced over 100-fold during adipocyte differentiation [5, 6]. The beneﬁcial effects of adiponectin are mediated through the ubiquitously expressed adiponectin receptors, AdipoR1 and AdipoR2, which represent a new class of membrane receptor [9]. APPL1 (adaptor protein containing phosphotyrosine-binding protein, pleckstrin homology domains, and leucine zipper 1) has recently been identiﬁed as a key signaling molecule that transduces the adiponectin signal from its receptors to downstream effectors [10]. Of these, the intracellular energy sensor AMP-activated protein kinase (AMPK) is central to many of adiponectins effects [11, 12], whilst activation of the peroxisome proliferator-activated receptor (PPAR)-a transcription factor is also pivotal [13]. Numerous clinical and epidemiological studies have demonstrated a decrease in circulating adiponectin levels in chronic inﬂammatory states such as obesity [14], insulin-resistance [15], type 2 diabetes [16], hypertension [17], cardiovascular disease [18], and liver disease [19, 20] (Figure 11.1). Polymorphisms at the adiponectin locus are predictors of circulating adiponectin levels, insulin sensitivity, and

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Figure 11.1 Circulating adiponectin levels in various states. See text for details.

atherosclerosis [21], whilst rare mutations in the adiponectin gene, called ADIPOQ, can impair adiponectin secretion and cause type 2 diabetes [22]. Males have lower circulating adiponectin levels than females, an observation that has been linked to the increased risk of insulin resistance and atherosclerosis in men [23–26], and testosterone accounts for this sexual dimorphism, with elevated adiponectin levels in hypogonadal men being reduced upon testosterone supplementation [25–27]. Many, if not all, of these correlations have been conﬁrmed in rodents, and the generation and characterization of adiponectin knockout mice has further established the importance of adiponectin in the etiology of the various disease states [28–33]. Adiponectin acts on multiple tissues and cell types to regulate numerous processes. It coordinates carbohydrate and lipid metabolism, primarily by its actions in liver and skeletal muscle, acting to decrease hepatic glucose output and increase peripheral glucose disposal, and increase fatty acid oxidation/decrease circulating fatty acids [3, 11, 12, 34]. Adiponectin acts on the endothelium to promote vasodilation [35] and has direct protective effects on the myocardium [36]. Recent evidence also supports an important role for adiponectin acting centrally in the brain [37]. At the molecular level adiponectin is no-less impressive, showing an array of modiﬁcations unparalleled by other adipokines. An overwhelming body of evidence has revealed that the resultant structural complexity is critical for both production and action of adiponectin, and gives rise to the variety of multimeric forms present in the circulation. In this chapter, we highlight current understanding of adiponectin structure and function, and its associations with health and disease.

11.2 Adiponectin Structure and Post-Translational Modifications

The adiponectin protein shares considerable homology with complement factor C1q and collagens, and consists of four domains (Figure 11.2): an N-terminal signal peptide that is cleaved upon entry into the endoplasmic reticulum; a variable region that shows greatest amino acid sequence divergence between species; a collagenous domain, so named because of its sequence and structural homology with collagen;

11.2 Adiponectin Structure and Post-Translational Modifications

Figure 11.2 Domain structure of human adiponectin. PTMs and highly conserved proline residues are detailed. The collagenous domain contains 22 G-X-Y or G-X-X collagen repeats involved in collagen triple-helix

formation. The globular domain exhibits a high degree of homology with subunits of C1q and TNF-a. aa, amino acid. See text for additional details.

and a C-terminal globular domain, involved in receptor binding. Synthesized as a single subunit, adiponectin undergoes extensive post-translation modiﬁcations (PTMs) during transit through the endoplasmic reticulum and Golgi. These PTMs are essential for multimerization, which occurs prior to secretion and results in formation of trimers, hexamers (collectively termed low-molecular-weight (LMW) multimers or sometimes referred to as LMW and medium-molecular-weight, respectively) and larger high-molecular-weight (HMW) multimers [22]. The structure of the various adiponectin multimers was elegantly illustrated by Lodishs group, using freeze-etch cryoelectron microscopy [38]. Adiponectin trimers are generated upon formation of a triple helix by noncovalent interactions within the collagenous domain and hydrophobic interactions between the globular heads [39]. Hydroxylation of multiple conserved proline residues in the collagenous domain appear to be involved in this process, since pharmacological inhibition of proline hydroxylation or expression in bacteria (which cannot perform these eukaryotic modiﬁcations) prevents trimer formation [40]. Numerous groups have demonstrated the importance of a conserved cysteine residue in the variable domain (Cys36 in human and Cys39 in mouse) that mediates the intertrimer disulﬁde bonds necessary for formation of hexamers and HMW multimers [22, 38, 41]. In addition, hydroxylation and subsequent glycosylation of four conserved lysine residues located within the collagenous domain are required for formation of the HMW multimers, since mutation of the glycosylated residues does not compromise trimer and hexamer formation, but abrogates formation of the HMW multimers [40, 42]. The presence of an a2–8-linked disialic acid moiety has also been reported although the site(s) and structural/functional signiﬁcance of this modiﬁcation are still to be determined [43, 44].

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11.3 Significance and Bioactivity of Adiponectin Multimers

The multimeric nature of circulating adiponectin has been demonstrated by a variety of biochemical separation techniques including velocity sedimentation, size-exclusion chromatography, and semidenaturing sodium dodecylsulfate– polyacrylamide gel electrophoresis. These studies have revealed that circulating adiponectin exists as a mixture of the trimer, hexamer, and HMW multimers [22, 38, 41], and that the multimers do not undergo interconversion once secreted [41]. A small amount of a truncated C-terminal globular form of adiponectin has been detected in human serum following immunoprecipitation [45] and Kadowakis group demonstrated that leukocyte elastase, a protease secreted by monocytes, is capable of producing such truncated fragments [46]. Whilst the physiological signiﬁcance of this truncated form remains unclear, as it has not been widely reported, a model that may account for the limited presence of such truncated forms of adiponectin was proposed by Scherer, based on a number of experimental observations [47]. Scherers group found that HMW multimers were cleared more rapidly than LMW oligomers following a metabolic challenge in mice. A recombinant form of adiponectin (C39S) that is incapable of forming oligomers larger than trimers and is sensitive to proteolytic cleavage following secretion was more biologically active than HMW adiponectin and subject to more rapid clearance. These and other observations that indicate an important role for HMW adiponectin in insulin sensitivity led them to propose that HMW adiponectin may be converted to a more biologically active form, through reduction and proteolytic cleavage [47]. Using the biochemical approaches mentioned above it has now been shown that decreases in total adiponectin observed in pathological states such as insulin resistance [48–50], the metabolic syndrome [51], hepatitis C genotype-3 infection [48] as well as testosterone-related decreases [25, 47] typically reﬂect a selective loss of HMW multimers and that effective therapeutic intervention is accompanied by an increase in the ratio of HMW to total adiponectin [47, 52–54]. Recently, several commercial enzyme-linked immunosorbent assays that facilitate measurement of the various multimers have been developed [55–57], making it possible to determine adiponectin proﬁles in a high-throughput manner. As a result, we have begun to see reports describing adiponectin proﬁles in larger cohorts, which further emphasizes the correlation between the decline in HMW adiponectin and cardiovascular disease [58, 59], insulin resistance, and the metabolic syndrome [60]. The above clinical and epidemiological observations are corroborated by direct investigations of the relative potencies/bioactivities of the LMW and HMW multimers in functional assays, in vitro and in vivo. Increasing evidence indicates that different adiponectin oligomers act on different target tissues and promote distinct biological actions. It is therefore important to take note of the source (bacterial or mammalian) and form (total, LMW or HMW) of recombinant adiponectin used in these functional investigations. In each of the following sections we present evidence supporting a role for the various adiponectin oligomers in key target tissues, followed

11.4 Adiponectin and Liver

by a brief description of relevant disease states with particular emphasis on metabolic dysregulation.

11.4 Adiponectin and Liver

A large body of evidence indicates that the HMW forms of adiponectin are the major effectors in liver. Scherer et al. ﬁrst demonstrated that recombinant adiponectin produced from mammalian cells, which produce the HMW multimers, reduced hepatic glucose output in diabetic mice [3]. In contrast, bacterially produced adiponectin, which is unable to form the HMW multimers [40, 42, 61], was virtually inactive. Furthermore, puriﬁed HMW adiponectin, but not LMW, reduced blood glucose in adiponectin null mice [47]. Consistent with this, Xu et al. demonstrated that hepatic AMPK activation and alleviation of hyperglycemia in diabetic mice was directly proportional to the concentration of HMW multimers administered [42]. Finally, unlike wild-type adiponectin, a glycosylation-defective mutant that could form only trimer and hexamer [40, 42] failed to enhance insulin-mediated suppression of glucose output in isolated primary rat hepatocytes [62]. Scherers group also showed that the beneﬁcial effects of TZDs are largely due to the increase in HMW adiponectin acting primarily on the liver in mice [47] and humans [54]. HMW adiponectin has also been proposed to be the most efﬁcacious adiponectin multimer for alleviation of fatty liver disease in obese mice [63]. Administration of adiponectin puriﬁed from mammalian cells ameliorated both alcoholic and nonalcoholic fatty liver disease (NAFLD) [64]. It also reduced hepatic production of proinﬂammatory cytokines such as tumor necrosis factor (TNF)-a [64], due at least in part to a direct effect on the Kuppfer cells [65]. In the human HepG2 hepatoma cell line, adiponectin promoted the expression of apolipoprotein A-I and ABCAI (ATP-binding cassette transporter) [66], which may facilitate increased assembly of high-density lipoprotein (HDL), and so provide a potential mechanism for the observed correlation between HDL and adiponectin [67, 68], particularly HMW adiponectin [51, 58]. The above functional studies provide a consensus regarding adiponectin and liver function; however, clinical and epidemiological investigations have generated mixed results with respect to adiponectins association with chronic liver disease (CLD). While several studies reported that circulating adiponectin levels were reduced in humans with nonalcoholic steatohepatitis [19, 20] and NAFLD [69, 70], adiponectin levels have also been shown to be elevated in subjects with chronic cirrhosis, independent of etiology [71, 72], and with increasing inﬂammation [73]. Interestingly, adiponectin levels were no longer found to correlate with insulin sensitivity or body massindex (BMI) in patients with CLD [74]. High adiponectin levels have beendetected in bile from patients with biliary obstruction and bile duct ligation in mice promoted a rapid increase in circulating levels, suggesting that biliary excretion contributes to adiponectin clearance and this may contribute to adiponectin accumulation, at least in states of cholestatic liver disease [74]. Further investigation is warranted to understand the complex interactions between adiponectin and the etiology of CLD.

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11.5 Adiponectin and Skeletal Muscle

Unlike the liver, the most physiologically relevant form(s) of adiponectin in skeletal muscle remain unclear. Lodishs group reported that bacterially produced globular adiponectin reduced circulating free fatty acids and glucose following a high fat meal or lipid injection [11, 45]. In vitro studies performed on isolated muscle strips and C2C12 myotubes showed that these effects were mediated by activation of AMPK, leading to increased fatty acid oxidation and glucose uptake. In contrast to globular adiponectin, full-length adiponectin produced in bacteria was relatively ineffective [11, 45]. Using similar forms of recombinant adiponectin, Kadowakis group found that chronic administration of globular adiponectin promoted partial restoration of insulin sensitivity in mouse models of obesity and lipoatrophy through increased expression of key molecules (CD36, acyl-CoA oxidase and uncoupling protein-2) and fatty acid oxidation in skeletal muscle. Once again, the relative efﬁcacy of full-length adiponectin was limited [2]. Lodish et al. also found that different adiponectin oligomers, produced in mammalian cells, activated discrete signaling pathways in C2C12 myotubes and muscle strips [38, 45, 61]. HMW and hexameric adiponectin promoted activation of nuclear factor-kB (NF-kB), but the trimeric isoform did not. In contrast, trimeric adiponectin, but not HMW or hexameric forms, stimulated phosphorylation and activation of AMPK [38, 45, 61]. Additional support for the potential importance of LMW oligomers in mediating the metabolic effects of adiponectin in skeletal muscle came from Kadowaki et al., when they identiﬁed the adiponectin receptors (see Section 11.13). However, given that these studies employed adiponectin puriﬁed from bacterial systems the physiological relevance is unclear. A more recent study found that HMWadiponectin puriﬁed from human serum had the greatest binding afﬁnity and most potent effect on AMPK activity in C2C12 myotubes, suggesting this may be the more physiologically relevant form [75]. There is reduced mitochondrial mass in skeletal muscle of mice lacking adiponectin and individuals with a family history of type 2 diabetes. Adiponectin has been shown to stimulate mitochondrial biogenesis in human myotubes, via AMPKdependent increases in PPAR-c coactivator-1a, providing a potential explanation [76]. Experimental evidence also suggests that adiponectin resistance may develop in skeletal muscle, at least in obesity, with compromised adiponectin signaling and action observed in isolated human skeletal muscle/cultured myotubes from obese subjects [77, 78] and obese mice [79].

11.6 Adiponectin and the Vasculature

All the adiponectin multimers are implicated in regulation of the endothelium and vasculature. Matsuzawa et al. demonstrated that adiponectin adheres to injured vascular endothelium [80] and also found that bacterially produced adiponectin blocked TNF-a-induced monocyte adhesion to endothelial cells, via inhibition of

11.7 Adiponectin and the Brain

induction of cell adhesion molecules and NF-kB activation [81, 82]. Several groups have subsequently shown that all forms of adiponectin are able to promote nitric oxide production, and suppress hyperglycemia-induced oxidative stress and inﬂammation in endothelial cells [35, 83, 84]. Thus, states of hypoadiponectinemia are associated with endothelial dysfunction [85], decreased endothelium-dependent vasodilation, and hypertension [17]. In normal tissues angiogenesis is a controlled process and this control is perturbed in many tumors. In this setting adiponectin has antiangiogenic activities, through inhibition of endothelial cell proliferation and migration and the promotion of endothelial cell apoptosis [86]. In contrast, injured vasculature is characterized by accelerated endothelial cell turnover and increased angiogenesis, especially in the context of ischemia. Adiponectin regulates the endothelial response to injury [87] by suppression of endothelial cell apoptosis [49] and stimulation of angiogenesis [88, 89]. The clinical relevance of the differing molecular weight multimers is again demonstrated by the more potent vascular protection afforded by the HMW multimers [49]. Adiponectin also attenuates smooth muscle proliferation and new intima formation in an oligomer-speciﬁc manner, and it binds to a number of growth factors [87, 90, 91] as well as a variety of circulating factors, including lipopolysaccharide (LPS) [92], albumin [56, 75], and cytokines such as monocyte chemoattractant protein (MCP)-1 [93]. Matsuzawas group demonstrated that adiponectin had direct effects on macrophages, with macrophage-to-foam cell progression, phagocytic activity, and LPS-induced TNF-a production all blocked by treatment with adiponectin [94, 95]. Walshs group have also recently shown that adiponectin protects against systemic inﬂammation by a mechanism involving macrophage-dependent clearance of early apoptotic cells through a receptor-dependent pathway involving calreticulin [96]. Collectively, these observations suggest that adiponectin is likely to provide important regulation by direct and indirect effects.

11.7 Adiponectin and the Brain

Several recent reports indicate an important role for LMW adiponectin in the brain. Although some studies concluded adiponectin did not cross the blood–brain barrier (BBB) [97, 98] this may be at least partly explained by the low levels of adiponectin found in the cerebrospinal ﬂuid (CSF), which are below the levels predicted by the baseline permeability of the BBB/blood–cerebrospinal barrier [99]. Nevertheless, it has become apparent that LMW, but not HMW, forms of adiponectin are found in the CSF in humans [100, 101] and mice [37], although only at concentrations around 1000-fold lower than those in serum. Unlike serum adiponectin, levels in the CSF showed no gender differences or correlation with insulin resistance [37, 100], probably because the reduction in circulating levels reﬂects a selective loss of HMW adiponectin. Interestingly, Kadowakis group reported that both circulating and CSF levels of adiponectin were increased in mice after fasting and reduced upon

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Figure 11.3 Major targets of adiponectin. FA, fatty acid; See text for details.

refeeding [37]. Importantly, they also showed that intracerebroventricular administration of adiponectin hexamer stimulated AMPK in the arcuate hypothalamus, via AdipoR1, increased food intake, and blocked leptins effects on these parameters. These observations prompted the intriguing proposal that adiponectin may have evolved as a starvation factor, serving to reduce energy expenditure and increase food intake during periods of fasting [37]. Such a suggestion had previously been proposed by Behre [102] and was also put forward by Scherer, who reported that a transgenic mouse line with elevated adiponectin levels grew morbidly obese, yet remained insulin sensitive [103]. It is unclear how observations made in fasted mice will translate to free-living humans, given the latters general reluctance to endure sustained periods of negative energy balance. Feeding does not typically alter adiponectin in humans [104, 105], although one study found that fasting increased circulating adiponectin levels [106], suggesting the regulatory mechanisms are likely to be conserved between species. Collectively (see Figure 11.3), these studies suggest that the HMW multimers are responsible for mediating the majority of adiponectins metabolic effects in peripheral tissues. In contrast, adiponectins central effects appear to be mediated by the LMW forms. Thus, understanding the processes governing the regulation of adiponectin multimer formation, secretion, and metabolism/clearance in both healthy and disease states is central to the identiﬁcation of novel therapeutic targets and the development of adiponectin-based treatments.

11.8 Adiponectin Expression and Secretion

Expression of adiponectin occurs from an intermediate stage of adipogenesis onwards [5, 6] and it may act in an autocrine fashion to promote adipogenesis [107]. Adiponectin production is common to adipocytes in both white and brown adipose

11.9 Adiponectin Secretion

tissues [47, 108]. Studies of isolated human explants or adipocytes suggest adiponectin secretion from visceral or subcutaneous depots is comparable [109] although it may be subject to depot-speciﬁc regulation [110], and differences have been reported for cells derived from visceral and subcutaneous depots, following differentiation in vitro [111]. Given the overwhelming evidence demonstrating that expression and secretion of adiponectin, particularly HMW adiponectin, is compromised in a variety of disease states, the next section describes our current understanding of adiponectins secretory pathway, followed by a discussion of some of the factors that have been shown to regulate adiponectin expression/secretion.

11.9 Adiponectin Secretion

While expression of adiponectin is largely restricted to adipocytes, other mammalian cell types can efﬁciently produce the same complement of multimers following ectopic expression [40]. Thus, the basic cellular machinery for production and secretion of adiponectin multimers would seem to be conserved. However, the possibility that adipocyte-speciﬁc regulation of various steps in the synthesis, multimerization and secretion of adiponectin remains. The molecular details of adiponectins secretory journey through the adipocyte are still relatively poorly deﬁned (Figure 11.4). Initial, pulse–chase experiments revealed that acute insulin treatment stimulated adiponectin secretion in a phosphoinositide-3-kinase-dependent manner [5, 112, 113]. Early comparisons of the intracellular distribution and secretion of adiponectin with other proteins suggested that adiponectin may enjoy a unique trafﬁcking itinerary and/or be regulated by distinct mechanisms [5, 112]. Support for the latter comes from a recent report demonstrating signiﬁcant biochemical overlap between adiponectin, adipsin, and transferrin receptors [114]. Important new insights into the molecular mechanisms facilitating transit of adiponectin through the secretory pathway are provided by two recent studies that demonstrate adiponectin is subject to thiol-mediated retention in the endoplasmic reticulum [115, 116]. Adiponectin binds directly to the resident endoplasmic reticulum chaperone, ERp44, via an intermolecular disulﬁde bond involving Cys36 (Cys39 in mouse) and this interaction is proposed to delay its secretion, facilitating PTMs and multimerization. The oxidoreductase Ero1-La is a preferred partner for ERp44, capable of using oxygen to generate disulﬁde bonds that it transfers to proteins such as ERp44 and protein disulﬁde isomerase, and displaces adiponectin from ERp44. Although ERp44 and Ero1-La are widely expressed, levels of both are increased during differentiation of adipocytes [115, 116]. Experimental approaches to manipulate ERp44 and/or Ero1-La provide evidence that both proteins play a key role in determining the secretion of adiponectin multimers. Scherers group found that increasing ERp44 levels increased the intracellular retention of adiponectin. Conversely, reducing ERp44 had the opposite effect [115]. Farmer et al. found that increasing Ero1-La increased adiponectin secretion and this was largely through an increase in the secretion of HMWadiponectin, whilst knockdown of Ero1-La reduced

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Figure 11.4 Summary of adiponectin multimerization and the secretory pathway. PTMs, and trimer, hexamer, and HMW multimer formation occur in the endoplasmic reticulum (ER) and Golgi prior to secretion. The ERp44/Ero1-La balance regulates production and secretion of adiponectin in a multimer-

specific fashion, and expression of ERp44/Ero1La is affected by factors including nutrient/ metabolic status and TZDs. Other proteins implicated in adiponectin trafficking are also shown. SIRT1, sirtuin-1. See text for additional details.

adiponectin secretion [116]. Additional studies demonstrated that expression of both ERp44 and Ero1-La are regulated in response to various stimuli. Levels of both proteins are reduced in obesity and in males – two situations where levels of HMW adiponectin are decreased. Conversely, both proteins are induced in response to PPAR-c agonists, which preferentially increase secretion of HMW adiponectin [47]. Moreover, the NAD-dependent deacetylase sirtuin-1, which regulates PPAR-c, was shown to represent a direct link between expression of ERp44 and Ero1-La and nutrient status [116]. Finally, insulin reduces the interaction of ERp44 with adiponectin [115], providing a potential mechanism for the observed increase in adiponectin secretion in response to acute insulin. These ﬁndings establish a mechanism by which effectors may speciﬁcally alter parameters of adiponectin secretion in a multimer-speciﬁc fashion, without perturb-

11.10 Ectopic Adiponectin Expression

ing secretion of other proteins, through an effect on the retention mechanisms rather than an effect on vesicle trafﬁcking per se. One such effector is testosterone. Matsuzawas group ﬁrst demonstrated that testosterone inhibited adiponectin secretion [26] and Xu et al. went on to show that this was due to a selective inhibition of HMW multimer secretion [25]. Although the levels of ERp44 and Ero1-La were not determined in these studies, given the above, it is tempting to speculate that changes in the levels and/or functional status of these proteins affect retention and secretion of HMW adiponectin as described. Changes in this process of intracellular retention are also likely to be of particular relevance in scenarios that affect cellular redox potential. Moving through the secretory pathway, several groups have demonstrated that secretion of adiponectin is blocked by treatment with the ADP ribosylation factor (ARF)-guanine nucleotide exchange factor) inhibitor, Brefeldin A, demonstrating the absolute dependence of adiponectin secretion on ARF activity and transit through the Golgi [40, 114, 117]. Consistent with this, ARF6 and the small G-protein Rab11 have recently been reported to play a role in basal and insulin-stimulated adiponectin secretion from 3T3-L1 adipocytes [114], while adiponectin also shows some colocalization with ARF1 [117]. The vesicle SNARE (soluble N-ethyl maleimide-sensitive fusion protein attachment protein receptor), vps10 tail-interacting protein-1a (Vti1a), which is localized to the trans-Golgi network (TGN) in some cell types and associates with glucose transporter 4 (GLUT4) vesicles in adipocytes, has also been implicated in adiponectin secretion [118]. Knockdown of Vti-1a impaired both basal and insulin-stimulated adiponectin secretion, as well as insulin-stimulated glucose uptake, prompting the suggestion that Vti-1a may be involved in an early sorting step common to both adiponectin and GLUT4 [118]. The recently discovered Golgilocalizing c-adaptin ear homology ARF-binding protein (Golgi-localizing c-adaptin ear homology ADP ribosylation factor-binding protein GGA) family of adapter proteins are known to play a key role in the differential regulation of secretion from the TGN. They function by facilitating sorting and recruitment of speciﬁc cargo molecules into clathrin-coated exocytic vesicles in an ARF-GTPase-dependent manner. GGA1, but not GGA2 or GGA3, has recently been implicated in adiponectin secretion [117]. Adiponectin-containing vesicles were speciﬁcally precipitated by the cargo binding domain of GGA1 and a dominant negative GGA1 mutant inhibited adiponectin secretion whilst coexpression of wild-type GGA1 enhanced adiponectin secretion [117].

11.10 Ectopic Adiponectin Expression

Several reports indicate adiponectin can be expressed in a number of nonadipose cells, often in response to stress. Adiponectin mRNA was detected in mouse liver parenchyma cells following administration of carbon tetrachloride [119] and in human liver biopsies from patients with steatosis [120]. Adiponectin expression was also induced in skeletal muscle following treatment with inﬂammatory

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cytokines, in both in vivo and in vitro settings, prompting the suggestion that the adiponectin response may represent a local anti-inﬂammatory mechanism and a means of increasing energy supply [121]. Expression of adiponectin has been detected in osteoblasts [122], which are derived from pluripotent precursor cells that can also give rise to adipocytes. Finally, regulated expression of adiponectin has been demonstrated in cytotrophoblast and syncytiotrophoblast cells in human placenta [123, 124], leading to the suggestion that adiponectin may play an important role in the regulation of energy metabolism at the materno-fetal interface.

11.11 Regulation of Expression and Secretion

Numerous investigators have sought to identify the mechanisms that contribute to changes in adiponectin expression and secretion. Early longitudinal studies, performed in rhesus monkeys, demonstrated that the reduction in circulating adiponectin levels paralleled the development of obesity and insulin resistance [125]. Importantly, the reduction in adiponectin levels occurred in the absence of any changes in adiponectin mRNA expression, strongly supporting the notion that the major effects are mediated at a post-transcriptional level, at least in this setting. Although in vitro studies demonstrate that acute insulin promotes adiponectin secretion, in vivo evidence from fat-speciﬁc insulin receptor knockout mice [126] and humans with severe loss of insulin receptor function [127, 128] or type 1 diabetes [129, 130] suggest that chronic insulin reduces circulating adiponectin levels. One possibility is that lack of insulin action leads to a reduction in reactive oxygen species (ROS) and that this accounts for the increase in adiponectin [128]. 11.11.1 Oxidative stress

Increased oxidative stress has been widely documented to contribute to the reduced levels of adiponectin via effects on transcription and secretion [131–134], and is implicated in multiple forms of insulin resistance [135]. In obese states, adipose tissue-speciﬁc changes in the balance of pro- and antioxidative enzymes results in increased levels of ROS within adipose tissue and increased systemic markers of oxidative stress [131]. This correlates with altered adipokine levels, including reduced adiponectin. Complementary studies in 3T3-L1 adipocytes showed that ROS levels were increased during differentiation and following treatment of cells with fatty acids. Treatment with NADPH oxidase inhibitors prevented the increase in ROS, indicating that NADPH oxidase is a major protagonist. Increased ROS were shown to directly reduce both PPAR-c and adiponectin mRNA expression and secretion, while increasing expression of other genes such as those for plasminogen activator inhibitor-1, interleukin (IL)-6, and MCP-1, and treatment with the antioxidant Nacetyl cysteine (NAC) reversed these effects [131]. Importantly, treatment of obese KKAy mice with the NADPH oxidase inhibitor apocynin for 3 weeks reduced both

11.11 Regulation of Expression and Secretion

systemic and adipose tissue ROS levels, without altering food intake or body weight, and increased mRNA expression and circulating levels of adiponectin suggesting such approaches may have therapeutic potential [131]. Similar effects have also been reported using angiotensin II type I receptor blockers. Treatment of obese mice with olmasartan reduced adipose ROS and NADPH oxidase levels, and increased circulating adiponectin levels, as well as improving other adipokine proﬁles [136]. In humans undergoing peritoneal dialysis, a 3-month treatment with candersartan reduced markers of oxidative stress and increased adiponectin levels [137]. Recent evidence indicates that induction of CCAAT/enhancer-binding protein (C/EBP) homologous protein (CCAAT/enhancer-binding protein homologous protein CHOP), which is an endoplasmic reticulum stress-induced protein, plays a key role in attenuation of adiponectin mRNA expression in the face of increasing ROS and endoplasmic reticulum stress [138, 139]. CHOP attenuates adiponectin promoter activity by direct antagonism of C/EBP-a binding. Increased suppressor of cytokine signaling-3 expression has also been implicated in the reduced expression of adiponectin observed in vitro [140] and in obese mouse models [141]. 11.11.2 Activators of PPARc – TZDs and Fish Oils

TZDs increase adiponectin levels via a combination of events occurring at the transcriptional [54] and post-transcriptional level [142]. TZDs such as pioglitazone reduce ROS [135], which may account for many of their beneﬁcial effects, including increased adiponectin levels. The efﬁcacy of TZDs in humans was shown to correlate more strongly with the increase in circulating levels of HMW adiponectin rather than changes in total adiponectin levels per se [47], and the TZD-induced increases in ERp44 and Ero-1La expression mentioned earlier [115] may contribute to the increase in HMW adiponectin. Fish oils and n-3 polyunsaturated fatty acids have also been shown to increase adiponectin levels in mice through PPAR-c activation, suggesting they may act via a similar mechanism [143, 144]. Studies from Scherers group, using adiponectin knockout mice, highlight the importance of adiponectin in the context of TZD-mediated improvements in metabolism, with obese, adiponectin knockout mice failing to respond to TZDs [32]. 11.11.3 Weight Loss

Improvements in insulin sensitivity following modest weight loss (8–9 kg) or exercise are not associated with changes in adiponectin levels [145] or multimer distribution [146]. However, more signiﬁcant weight loss (10% or greater reduction in BMI) has been reported to increase adiponectin levels [52, 147–149], and promote a shift in the multimer distribution, with increased levels of HMW adiponectin and a concomitant reduction in LMW levels [49, 52]. A recent study suggests that the weight loss-induced increase in circulating adiponectin can occur in the absence of changes in adiponectin mRNA levels, consistent with an important role for

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post-transcriptional mechanisms [150]. Changes in adiponectin do not appear to be involved in improved insulin sensitivity associated with exercise, in the absence of changes in fat mass [149]; however, emerging evidence suggests that changes in expression of the adiponectin receptors may play a role [151]. 11.11.4 Other Agents

The cannabinoid receptor subtype-1 antagonist rimonabant increases adiponectin expression and secretion in vivo and in 3T3-L1 adipocytes [152, 153], leading to the suggestion that rimonabant may mediate its effects at least partly by increasing adiponectin, independent of effects on appetite, although these data are not supported by work from all groups [154]. b-Adrenergic agonists (and cAMP) promote reduced expression and secretion of adiponectin, both in vivo and in vitro [155], whilst adiponectin itself has been reported to be involved in a negative feedback loop, with exogenous adiponectin reducing expression of adiponectin and AdipoR2, but not AdipoR1, both in vivo and in vitro [156]. In vitro systems have been widely employed to examine the effects of a variety of agents on adiponectin expression and secretion. Experiments in 3T3-L1 cells suggest that insulin, TNF-a, or dexamethasone, but not growth hormone, angiotensin II or tri-iodothyronine, reduce adiponectin mRNA expression [157], whilst growth hormone and prolactin suppress adiponectin expression from human adipose tissue [158]. As described, oxidative and endoplasmic reticulum stress reduce adiponectin expression [131, 159], whilst concomitantly increasing expression of proinﬂammatory cytokines [160]. Proinﬂammatory cytokines in turn increase oxidative stress and thereby decrease adiponectin expression producing a vicious cycle. Interestingly, recent evidence demonstrates this can occur without changes in multimer distribution [161]. Antioxidants such as NAC are able to restore adiponectin expression and secretion and also prevent other cytokine-induced effects such as activation of NF-kB [162]. In contrast to most cytokines, IL-15 increases adiponectin secretion [163] whilst the widely used antidiabetic drug metformin reduced adiponectin expression in 3T3-L1 adipocytes [164].

11.12 Adiponectin Clearance

Relatively little is known about how adiponectin is metabolized and cleared from the circulation. Once in the circulation, adiponectin oligomers are stable and do not undergo exchange from one form to another [41, 105]. Studies in mice and rabbits suggest the HMW multimers are cleared more rapidly than LMWoligomers [41, 105]. Recent investigations in humans are consistent with this and also highlight how changes in total adiponectin reﬂect changes in HMW multimers [165]. Adiponectin has been detected in urine from type 2 diabetic subjects and healthy males [166]. Urinary adiponectin levels, which are around three orders of magnitude lower than

11.13 Adiponectin Receptors and Downstream Effectors

serum levels, were signiﬁcantly elevated in patients with macro-albuminuria and correlated with urinary albumin. These observations support a model where leakage of circulating adiponectin is largely responsible for the high urinary adiponectin [166]. Intriguingly serum adiponectin levels were also elevated in patients with macro-albuminuria, suggesting there may be a compensatory mechanism to assuage microvascular damage in the advanced stages of diabetic nephropathy by increased production of adiponectin. As in the case of CLD, where altered biliary clearance of adiponectin has been implicated [74], the relationship between adiponectin and chronic or end-stage kidney disease appears complex. Increased adiponectin levels have recently been associated with a range of positive and negative outcomes, presenting something of an epidemiological conundrum in this setting [167]. It is clear from the above that expression of the adiponectin gene is regulated by multiple factors. Once translated, the protein undergoes a series of PTMs as it passes through the secretory pathway and these are intimately involved in the adiponectin multimerization process. As with adiponectin expression, multimerization and secretion are subject to multiple levels of regulation. Although relatively little is known about mechanisms governing adiponectin clearance, studies suggest this may also be altered in pathophysiological states. Interventions at any or all of these steps represent attractive therapeutic opportunities. In the next section we describe the adiponectin receptors. Increasing evidence indicates that these are responsible for mediating the majority of adiponectins effects.

11.13 Adiponectin Receptors and Downstream Effectors

The adiponectin receptors, termed AdipoR1 and AdipoR2, were identiﬁed by expression cloning by Kadowaki et al. [9]. They share a high degree of homology and are conserved from yeast to humans [168]. Structural predictions for AdipoR1 and AdipoR2 identiﬁed both as members of a novel class of seven transmembrane domain proteins, termed progestin and AdipoQ receptors [169]. While they lack signiﬁcant homology with other proteins they are distantly related to G-proteincoupled receptors (GPCRs); however, they exhibit reversed topology with an intracellular N-terminus and do not signal via classic GPCR pathways (G-proteins or calcium) [9]. A large body of evidence supports the hypothesis that these adiponectin receptors serve to transduce adiponectin signals. Typically, increasing AdipoR1 and/ or AdipoR2 expression increases adiponectin binding, signaling, and action [9]. Conversely, reducing receptor expression has reciprocal effects [9]. Until recently a lack of suitable antibodies has limited investigation of AdipoR1 and AdipoR2 proteins in relevant tissues, and most studies have combined measurement of mRNA levels with readouts of adiponectin action to infer relative signiﬁcance and regulation of the two receptors [79, 122, 170–174]. Notable ﬁndings from these studies include the early observations that expression of AdipoR1/R2 is reduced by insulin [79]; AdipoR1 expression is reduced in adipose tissue from obese subjects [174]; TZDs do not change expression of AdipoR1/R2 [175]; ﬁbrates increase

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expression of AdipoR1/R2 in adipose tissue [176] but not skeletal muscle [177]; and feeding reduces expression of AdipoR1/R2 [79, 178]. Collectively these studies demonstrate that, like adiponectin, expression of the adiponectin receptors is subject to regulation in response to multiple factors. The characteristics of adiponectin binding to the two receptors have still to be rigorously examined. In the original study, Kadowakis group employed recombinant adiponectin (full-length or globular) puriﬁed from bacteria. Notwithstanding, differences were observed with AdipoR1-binding globular adiponectin with high afﬁnity whilst AdipoR2 showed intermediate afﬁnity for both globular and full-length adiponectin [9]. A more recent report using puriﬁed trimer, hexamer, and HMW adiponectin from human serum demonstrated that HMW adiponectin was the most potent activator of AMPK in C2C12 myotubes, which express high levels of AdipoR1 [75]. Kadowakis group also recently reported that manipulation of AdipoR1 and/or AdipoR2 in mice altered metabolism in a manner consistent with a central role for both receptors in adiponectin action [179]. The two receptors appeared to have speciﬁc downstream targets in liver, with AdipoR1 linked more closely to AMPK whereas AdipoR2 was linked more closely with PPAR-a, prompting speculation that functional differences may be accounted for by AdipoR1- or AdipoR2-speciﬁc intracellular binding proteins [179]. In an independent, study Linden et al. found that AdipoR1/ mice were obese, glucose-intolerant, and showed decreased energy expenditure consistent with a positive role for AdipoR1 in energy metabolism; however, AMPK activity was not altered [180]. Perhaps more unexpectedly, they found that AdipoR2/ mice displayed improved insulin sensitivity, exhibiting resistance to diet-induced obesity and increased energy expenditure [180]. Whilst the latter ﬁndings are difﬁcult to rationalize in the context of the body of evidence in favor of a positive role for both AdipoR1 and AdipoR2 they are supported by another recent study from Reifel-Miller et al., who found that ablation of AdipoR2 reduced diet-induced weight gain, dyslipidemia, and insulin resistance [181]. Collectively, these studies emphasize the important metabolic role of AdipoR1 and AdipoR2, and also highlight the limitations of our current understanding and the need for additional investigations to further deﬁne the molecular details and physiological roles of these receptors.

11.14 Adiponectin Signaling

Adiponectin stimulates the activation of a number of intracellular kinases including AMPK, p38 mitogen-activated protein kinase (MAPK) and c-Jun N-terminal kinase (JNK) via AdipoR1 and AdipoR2 [182]. The signaling pathways are poorly deﬁned, but Dong et al. recently identiﬁed APPL1 as a key transducer of the adiponectin signal [10]. APPL1 binds constitutively to both AdipoR1 and AdipoR2, and adiponectin increases this interaction [10]. The phosphotyrosine-binding domain of APPL1 is required for binding, although the receptors are not phosphorylated on

11.15 Conclusions

tyrosine residues [10], and binding presumably occurs via the conserved region of the cytoplasmic tail of the receptors. Adiponectin-stimulated phosphorylation of AMPK and p38 MAPK was increased in C2C12 cells overexpressing APPL1 and decreased in cells with reduced APPL1 [10]. APPL1 has recently been shown to interact directly with AKT2 in 3T3-L1 adipocytes, and it appears to act as a point of convergence between the insulin and adiponectin signaling cascades [10, 182, 183]. APPL1 is also required for the adiponectin-induced, AMPK-dependent phosphorylation and activation of endothelial nitric oxide synthetase in human endothelial cells [84]. Thus, APPL1 has already been implicated in adiponectin-stimulated glucose uptake, fatty acid oxidation [10], and nitric oxide-induced vasodilatation [84], and is likely to be involved in many more of adiponectins pleiotropic actions. Like APPL1, the energy sensor and metabolic regulator AMPK is implicated in most of adiponectins actions and is itself an attractive therapeutic target for obesityrelated diseases. It is a heterotrimer made up of a catalytic a-subunit, and regulatory b- and c-subunits, with distinct isoforms of each subunit (a1/a2, b1/b2 and c1/c2/ c3) present at varying levels in different tissues. Activation of AMPK is complex and can be promoted by a combination of events that include a decrease in the cellular AMP/ATP ratio [184], phosphorylation by upstream kinases, of which four have now been identiﬁed (LKB1 [185], calmodulin-dependent protein kinase kinases [186, 187], TAK1 [188], and ataxia telangiectasia mutated kinase [189]), and reduced accessibility to phosphatases [190]. As such, the molecular details by which adiponectin stimulates AMPK remain to be deﬁned and it seems likely that different mechanisms will exist in different cell types. Adiponectin induces fatty acid oxidation in cultured muscle cells (C2C12) by sequential activation of an AMPK–p38 MAPK–PPAR-a pathway [191], whilst the AMPK–p38 MAPK axis also results in NF-kB-dependent, IL-6 production in human synovial ﬁbroblasts [192], suggesting this may be a common pathway. Adiponectin activates JNK in a range of cancer cell lines [193], suppressing constitutive activation of signal transducer and activator of transcription-3, which may contribute to adiponectins anticancer effects [194], and in osteoblasts [195], where JNK activation was linked to proliferation. Interestingly, Goldstein et al. found that full-length and globular adiponectin reduced glucose-induced ROS in human endothelial cells via AMPK-independent, cAMP/protein kinase A (PKA)-dependent pathways [83] and have more recently demonstrated that both AMPK-dependent and cAMP/PKAdependent pathways are involved in adiponectins suppression of IkB activation induced by TNF-a or glucose in these cells [196]. These data highlight how multiple parallel signaling pathways may mediate adiponectins effects and future investigations should help to elaborate the molecular details in different cell types.

11.15 Conclusions

The association between hypoadiponectinemia and metabolic disease provides an ideal scenario for adiponectin therapy. Therapeutic approaches must give due

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consideration to our evolving appreciation of the complexity of adiponectin biology and its pleiotropic actions. Hypoadiponectinemia is characterized by a preferential loss of HMWadiponectin, the major effector in peripheral tissues such as liver, whilst the LMW oligomers, which mediate adiponectins central actions, are relatively unchanged. Hence, strategies to selectively increase the circulating levels of HMW adiponectin appear desirable. This may be achieved by simple administration of recombinant HMW adiponectin or by alternative therapies such as the TZDs, which selectively increase the production and secretion of HMW adiponectin from adipocytes. Whilst emerging evidence suggests that adiponectin resistance may be a feature of obesity, ﬁndings to date, albeit mainly from animal studies, suggest that adiponectin treatment will be sufﬁcient to ameliorate many of the complications. In conclusion, the prospect of adiponectin therapy or regulation of adiponectin multimer distribution by affecting production, secretion, clearance or action is an attractive one for the treatment of many facets of obesity-associated diseases. Continued basic and clinical research efforts are warranted to realize the maximum potential of adiponectin-based strategies.

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B.D., Snowden, M.A., and Carling, D. (2007) Investigating the mechanism for AMP activation of the AMP-activated protein kinase cascade. Biochem. J., 403, 139–148. Yoon, M.J., Lee, G.Y., Chung, J.-J., Ahn, Y.H., Hong, S.H., and Kim, J.B. (2006) Adiponectin increases fatty acid oxidation in skeletal muscle cells by sequential activation of AMP-activated protein kinase, p38 mitogen-activated protein kinase, and peroxisome proliferatoractivated receptor a. Diabetes, 55, 2562–2570. Tang, C.-H., Chiu, Y.-C., Tan, T.-W., Yang, R.-S., and Fu, W.-M. (2007) Adiponectin enhances IL-6 production in human synovial ﬁbroblast via an AdipoR1 receptor, AMPK, p38, and NF-kB pathway. J. Immunol., 179, 5483–5492. Miyazaki, T., Bub, J.D., Uzuki, M., and Iwamoto, Y. (2005) Adiponectin activates c-Jun NH2-terminal kinase and inhibits signal transducer and activator of transcription 3. Biochem. Biophys. Res. Commun., 333, 79–87. Kelesidis, I., Kelesidis, T., and Mantzoros, C.S. (2006) Adiponectin and cancer: a systematic review. Br. J. Cancer, 94, 1221–1225. Luo, X.H., Guo, L.J., Yuan, L.Q., Xie, H., Zhou, H.D., Wu, X.P., and Liao, E.Y. (2005) Adiponectin stimulates human osteoblasts proliferation and differentiation via the MAPK signaling pathway. Exp. Cell. Res., 309, 99–109. Wu, X., Mahadev, K., Fuchsel, L., Ouedraogo, R., Xu, S.-q., and Goldstein, B.J. (2007) Adiponectin suppresses IkB kinase activation induced by tumor necrosis factor-a or high glucose in endothelial cells: role of cAMP and AMP kinase signaling. Am. J. Physiol. Endocrinol. Metab., 293, E1836–E1844.

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12 Preadipocyte factor-1 and Adipose Tissue-Specific Secretory Factor/Resistin – Two Secreted Factors from Adipose Tissue: Role in Adipogenesis and Insulin Resistance Hei Sook Sul, Yuhui Wang, and Carolyn Hudak 12.1 Introduction

Although the principal mechanisms linking increased adiposity with insulin resistance are only partially understood, obesity is associated with major diseases such as diabetes and cardiovascular diseases. Adipose tissue development can be affected by genetic background, hormonal balance, diet, and physical activity. Adipose tissue mass can increase by increased fat cell size due to lipid accumulation as well as by increased number of fat cells arising from differentiation of preadipocytes into adipocytes. Peroxisome proliferator-activated receptor (PPAR)-c and CCAAT/ enhancer-binding protein (C/EBP) play critical roles in adipocyte differentiation [1–3]. Adipocyte differentiation and expression of these transcription factors can be regulated by various factors. In this regard, in addition to being the main site for energy storage in the body, adipose tissue has recently been appreciated as a major endocrine organ. Adipose tissue secretes a wide array of molecules that are involved in various physiological processes, including immune response, vascular function, and energy homeostasis, as well as growth and development of the adipose tissue itself. Among these factors are leptin, adiponectin, tumor necrosis factor (TNF)-a and other cytokines, including interleukin (IL)-6 and IL-8. In this chapter, we address the role of two secreted factors from adipose tissue – preadipocyte factor (Pref)-1 and adipose tissue-speciﬁc secretory factor (ADSF)/resistin. We discuss current knowledge on their role in the regulation of adipose tissue development, their signaling mechanisms, as well as their involvement in different pathological disorders such as obesity-related diabetes.

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12.2 Pref-1 Structure

We originally cloned Pref-1 by differential screening of a 3T3-L1 cDNA library, by the criteria of preadipocyte-speciﬁc expression and absence in other tissues [4]. Pref-1 is expressed in multiple mouse embryonic tissues, such as liver, lung, tongue, pituitary, and developing vertebrae [4], and placenta, and detectable amounts of circulating Pref-1 are found in maternal serum in concentrations that correlate with the number of fetuses [5]. Dlk-1, the human homolog of Pref-1, also identiﬁed as fetal antigen (FA)-1 [6] by puriﬁcation from fetal circulation, has been shown to be expressed in a wide array of human embryonic tissues [7]. However, after birth, expression of Pref-1 is rapidly abolished in most tissues and becomes restricted to certain cell types that include preadipocytes [4], thymic stromal cells [8] as well as some of the neuroendocrine type of cells, such as pancreatic islet b-cells [9], adrenal glands [10], and pituitary. Pref-1 is synthesized as a protein of 385 amino acids that contains a signal sequence at the N-terminus and a single membrane-spanning domain of amino acids 300–322 (Figure 12.1). The most striking structural feature of Pref-1 is the presence of six tandem epidermal growth factor (EGF)-like repeats in the extracellular domain that maintain both the conserved spacing of six cysteines for the formation of three disulﬁde bonds and other amino acids characteristic of the EGF-like repeat motif-containing proteins (Figure 12.1) [4]. The EGF-like motif was originally described for EGF and subsequently for other growth factors, which, by binding to EGF receptor, act as signals for cell proliferation and differentiation. However, Pref-1 does not contain the conserved amino acid residues that are required for EGF receptor binding. Rather, Pref-1 shares higher structural homology with another class of EGF-like repeat containing signaling proteins, the Notch/Delta/Serrate family that are involved in cell signaling and cell fate determination. Pref-1 however, lacks the DSL (Delta Serrate Lin12) domain that is conserved in all Notch ligands to mediate receptor–ligand interaction for Notch [11], indicating that Pref-1 would not act as a Notch ligand. In preadipocytes, multiple transmembrane forms of Pref-1, ranging from 50 to 60 kDa,arefound in thecell membrane due, inpart,to post-translational modiﬁcations

Figure 12.1 Domain structure of Pref-1 isoforms. S, signal sequence; Jm, juxtamembrane domain; Tm, transmembrane domain; Cy, cytoplasmic region; D and P, distal and proximal cleavage sites.

12.3 Pref-1 Inhibition of Adipocyte Differentiation

containing N-linked oligosaccharides as well as sialic acids. Moreover, there are four major alternative splicing products of Pref-1 (Pref-1A–D) (Figure 12.1) [12]. In addition to the largest full-length Pref-1 form, alternate splicing generates three major shorter forms of Pref-1, each containing in-frame deletions in the extracellular juxtamembrane region or EGF-like repeat domain (Figure 12.1). The relative abundance of the different spliced forms varies depending on the tissue or developmental stage investigated [13]. Pref-1 can be proteolytically cleaved at the extracellular domain at two sites to generate soluble forms of Pref-1 [13, 14]. Thus, the two larger alternate splicing forms of Pref-1, Pref-1A and -1B, are cleaved at a juxtamembrane as well as at a more N-terminal processing sites to generate a larger 50-kDa and a smaller 25-kDa soluble form. The smaller Pref-1C and -1D, due to the larger deletions that include the juxtamembrane processing site, are cleaved only at the N-terminal site to generate the smaller 25-kDa soluble form, but not the larger 50-kDa form. In this regard, FA-1, the protein in fetal circulation, likely corresponds to the larger soluble form of human Pref-1. Using various inhibitors and in vitro approaches, we found that one of the ADAM (a disintegrin and metalloproteinase) family members, TNF-a-converting enzyme (tumor necrosis factor-a-converting enzyme TACE, ADAM17), can cleave the Pref-1 at the juxtamembrane region to generate the large soluble form of Pref-1. We also found that both basal and stimulated cleavage was inhibited by the broad metalloproteinase inhibitor GM6001, suggesting that cleavage of membrane Pref-1 is dependent on a metalloproteinase. In addition, lentivirus-mediated overexpression of TACE increased Pref-1 cleavage to produce the large soluble form. Conversely, knockdown of TACE by transfecting small interfering RNA (siRNA) decreased the release of the large soluble form from the membrane form [15]. In addition, this cleavage was not detectable or was markedly decreased in cells bearing mutated TACE or in cells transfected with TACE siRNA. These data clearly demonstrate TACE mediated cleavage at the juxtamembrane and generation of the large 50-kDa soluble form of Pref-1. We also found that the release of the 50-kDa full extracellular domain was markedly enhanced by phorbol 12-myristate 13-acetate treatment in a dose- and time-dependent manner. This indicated that cleavage of Pref-1 is regulated by protein kinase C.

12.3 Pref-1 Inhibition of Adipocyte Differentiation

Pref-1 is highly expressed in 3T3-L1 preadipocytes, but decreases during differentiation and is absent in mature adipocytes [4, 16]. Pref-1 expression is downregulated speciﬁcally by the synthetic glucocorticoid dexamethasone – a component of the differentiation-inducing agents dexamethasone, methylisobutylxanthine, and insulin that are routinely used for adipocyte differentiation in vitro. Glucocorticoid receptors are present in preadipocytes, and glucocorticoids have been attributed to enhance adipocyte differentiation by increasing C/EBP-d and PPAR-c expression. We found a close correlation between dexamethasone-mediated downregulation of

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Pref-1 and efﬁcacy of dexamethasone on differentiation of 3T3-L1 preadipocytes. Thus, dexamethasone may induce adipocyte differentiation partly by suppressing Pref-1 expression [17]. Using various in vitro approaches, we have shown the inhibitory role of Pref-1 in adipocyte differentiation. In 3T3-L1 cells, constitutive expression of Pref-1 by stable transfection markedly lowers the degree of adipocyte differentiation. Pref-1 prevents lipid accumulation and expression of various adipocyte markers, such as fatty acid synthase (FAS), stearoyl-CoA desaturase (SCD), aP2 (adipocyte-selective fatty acid-binding protein), PPAR-c, and C/EBP-a. Conversely, decreasing Pref-1 levels by transfection of antisense sequence greatly enhances adipogenesis [4, 17]. This suggests that Pref-1 acts at an early stage during the differentiation process to inhibit adipogenesis [14]. Overall, these studies demonstrate that Pref-1 expression inhibits adipocyte conversion of 3T3-L1 cells and that downregulation of Pref-1 is a necessary step in adipocyte differentiation. The human homolog of Pref-1, Dlk-1, inhibits 3T3-L1 differentiation, demonstrating that Pref-1 and Dlk-1 are functionally equivalent [18]. Since Pref-1 can generate a soluble form by cleavage of the extracellular domain, the above-described transfection of full-length Pref-1 cannot distinguish the effect of the membrane form of Pref-1 from soluble Pref-1 in regulating adipocyte differentiation. However, addition of the large soluble form expressed in COS cells as well as Pref-1 extracellular domain fused to glutathione-S-transferase expressed in Escherichia coli are effective at inhibiting adipocyte differentiation. In addition, unlike Pref1A and -1B, which generate a 50-kDa soluble Pref-1, Pref-1C and -1D, that still produce 25-kDa cleavage product but not the large 50-kDa soluble form, are not effective at inhibiting adipocyte differentiation. These observations clearly show the inhibitory effect of the large soluble 50-kDa protein but not the full-length membrane form on adipocyte differentiation. Furthermore, transfection of a noncleavable transmembrane form of Pref-1A, constructed by the deletion of the 22 amino acids containing the putative cleavage site at the juxtamembrane region, conﬁrmed that the membrane form of Pref-1 cannot inhibit adipocyte differentiation. Since the smaller 25-kDa cleavage product is not effective at inhibiting adipocyte differentiation, the additional N-terminal cleavage may be a means to inactivate the biologically active 50kDa soluble Pref-1.

12.4 Mechanism for Pref-1 Function

We have used Pref-1 null mouse embryonic ﬁbroblasts (MEFs) to examine the signaling pathways for Pref-1 inhibition of adipocyte differentiation. As predicted, the degree of adipose conversion is higher in Pref-1 null MEFs as compared to wildtype MEFs. Compared to the 50% differentiation of wild-type MEFs into adipocytes, 90% of Pref-1 null MEFs differentiated into adipocytes. Furthermore, infection of lentivirus containing the soluble form of Pref-1 into Pref-1 null MEFs decreased the degree of differentiation, conﬁrming the inhibitory effect of Pref-1 on adipogenesis [15]. We recently demonstrated that the inhibitory effect of Pref-1 is mediated

12.5 In Vivo Effect of Pref-1 on Adipogenesis and Glucose/Insulin Homeostasis

through activation of the mitogen-activated protein kinase kinase (MEK)/extracellular signal-regulated kinase (ERK) pathway [19]. Addition of the large soluble Pref-1 increases the phosphorylation of ERK in a time- and dose-dependent manner. Both wild-type and Pref-1 null MEFs show a transient burst of ERK phosphorylation upon addition of adipogenic agents. Wild-type MEFs also show a low but signiﬁcant second increase in ERK phosphorylation peaking at day 2 that parallels the expression level of Pref-1. This second increase in ERK phosphorylation is absent in Pref-1 null MEFs. Furthermore, a speciﬁc MEK inhibitor or siRNA-mediated ERK depletion prevented the second ERK phosphorylation and enhanced adipocyte conversion of wild-type MEFs. Furthermore, treatment of Pref-1 null MEFs with Pref-1 restores the second ERK phosphorylation, resulting in inhibition of adipocyte differentiation [19]. These studies show that Pref-1 inhibition of adipogenesis is by activation of ERK/MEK pathway during differentiation. Since the cleaved Pref-1 ectodomain containing six EGF-like repeats, which are presumably involved in protein–protein interaction, is biologically active, we predict that Pref-1 acts as a ligand for a yet to be identiﬁed EGFrepeat-containing receptor to activate ERK/MEK pathway to inhibit adipocyte differentiation. Identiﬁcation of the Pref-1 receptor is critical for understanding the mode of Pref-1 action.

12.5 In Vivo Effect of Pref-1 on Adipogenesis and Glucose/Insulin Homeostasis

To determine the role of Pref-1 in vivo, we generated Pref-1 null mice [20] as well as transgenic mice overexpressing the large soluble form of Pref-1 in adipose tissue [21]. In Pref-1 null mice, both male and female mice weighed signiﬁcantly less than wild-type mice at weaning. However, Pref-1 null mice gained body weight more rapidly. The weight of major fat depots (inguinal, retroperitoneal, and gonadal) was signiﬁcantly higher in Pref-1 null mice, indicating that the accelerated body weight gain in Pref-1 null mice was due to an increase in adipose tissue mass. Histological analysis of fat depots revealed that adipocytes from Pref-1 null mice were bigger than those from wild-type littermates. Moreover, mRNA levels of various markers of adipocyte differentiation, including, C/EBP-a, SCD, FAS as well as ADSF/ resistin and other adipocyte-secreted factors were signiﬁcantly higher in adipose tissue from Pref-1 null mice, indicating enhanced adipogenesis. Pref-1 null mice also showed enlarged fatty liver as well as increased circulating levels of triglycerides, cholesterol, and free fatty acids – characteristics usually associated with obesity. Thus, studies of Pref-1 null mice demonstrate that ablation of Pref-1 enhances adipogenesis in vivo and support the proposed role of Pref-1 as a negative regulator of adipogenesis [20]. Transgenic mice overexpressing the soluble form of Pref-1 as a human immunoglobulin-c constant region (hFc) fusion protein in adipose tissue were generated using the aP2 promoter. These transgenic mice showed a marked decrease in adipose tissue mass, and reduced expression of adipocyte markers and adipocyte secreted factors. With decreased adipose tissue development, as observed in lipodystrophy

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mouse models, these mice suffered from hypertriglyceridemia, decreased glucose tolerance, and lower insulin sensitivity [21]. Mice expressing the Pref-1–hFc exclusively in liver under the control of the albumin promoter also showed a decrease in adipose mass and adipocyte marker expression, suggesting an endocrine mode of action of Pref-1 [21]. Data from these in vivo studies with Pref-1 knockout and transgenic mice models are consistent with in vitro studies, strongly demonstrating the inhibitory effect of Pref-1 on adipogenesis. The results from Pref-1 null and Pref1-overexpressing transgenic mice also suggest that proper development of adipose tissue is critical for maintenance of glucose/insulin homeostasis. Pref-1 is encoded by the gene dlk1 and, along with dat, gtl2, peg11, antipeg11, and meg8, is located in an imprinted syntenic region of mouse chromosome 12, human chromosome 14, and sheep chromosome 18. Paternal monoallelic expression of the Pref-1 gene, due to differential methylation, is well documented [22–24]. Given the role of imprinted genes in fetal growth and development in general, and the expression of Pref-1 during embryonic development in a variety of tissues, Pref-1 undoubtedly has a function beyond the regulation of adipogenesis [22–26]. Pref-1 null mice, in addition to increased adiposity, display greater than 50% perinatal lethality, and surviving animals show growth retardation, skeletal malformation, and eyelid defects [20]. In addition, Pref-1 transgenic mice not only have reduced adiposity but also show skeletal defects [21]. Pref-1 null and transgenic mice show distinct defects similar to maternal uniparenteral disomy (UPD) syndrome UPD12 and paternal UPD12 in mice, respectively, and syntenic maternal and paternal UPD14 syndromes in humans. Calipyge sheep with a mutation in chromosome 18 show decreased adiposity with elevated Pref-1 levels. Taken together, these results strongly suggest that Pref-1 is involved not only in adipogenesis, but also in embryonic development, and that the observed UPD syndromes are at least partly due to altered Pref-1 expression levels. In this regard, as during adipocyte differentiation, Pref-1 may function as a soluble factor, maintaining proliferating cells in an undifferentiated state during mouse embryonic development.

12.6 ADSF/Resistin: Identification and Structure

ADSF/resistin was identiﬁed by three independent groups via different approaches. ADSF/resistin was identiﬁed in our laboratory as ADSF that inhibits adipocyte differentiation of 3T3-L1 cells [27]. This protein was also identiﬁed as an adipocytesecreted hormone whose expression is suppressed by the insulin-sensitizing PPAR-c agonists thiazolidinediones and was named resistin [28]. Steppan et al. reported that the circulating levels of resistin were high in genetically obese mice and mice treated with recombinant resistin had increased insulin resistance, while administration of antibody against resistin improved insulin sensitivity. Thus, these authors suggested resistin as a potential link between obesity and insulin resistance. Yet, Holcomb et al. originally identiﬁed the same protein, FIZZ3, as a member of the FIZZ (found in inﬂammatory zone) family of cysteine-rich secreted proteins. These authors

12.7 ADSF/Resistin Expression and Function

originally found FIZZ1 in bronchoalveolar lavage ﬂuid from mice with ovalbumininduced pulmonary inﬂammation [29]. FIZZ2 was reported to have similar action to resistin in insulin resistance [30]. In this chapter, the protein is referred to as ADSF and/or resistin. ADSF/resistin is a 114-amino-acid polypeptide that has a unique cysteine repeat motif at the C-terminus [27]. It exists as both a monomer and homodimer, and it can form a hetero-oligomer with the two other members of this family [30–32]. This oligomerzation is mediated by disulﬁde bonds at cysteine residues [33]. A recent crystal structure study indicates that resistin circulates as two distinct assembly states, probably corresponding to hexamers as well as more bioactive trimers [34].

12.7 ADSF/Resistin Expression and Function

In rodents, ADSF/resistin mRNA is exclusively expressed in adipose tissue, including brown adipose tissue, and inguinal, gonadal, and retroperitoneal fat pads of white adipose tissue. ADSF/resistin mRNA is markedly increased at later stages of 3T3-L1 as well as primary rat preadipocyte differentiation into adipocytes [27]. In humans, however, resistin is abundantly expressed in peripheral monocytes and macrophages and, if expressed, at a very low level in adipocytes [35]. Treatment of 3T3-L1 cells with conditioned media from COS cells transfected with ADSF/resistin suppressed differentiation of these cells into adipocytes, demonstrating in vitro inhibition of adipogenesis by ADSF/resistin [27]. Similarly, another member of this protein family of FIZZ1 (also called resistin-related molecule-b) has also been shown to inhibit adipocyte differentiation of 3T3-L1 preadipocytes [31]. To examine ADSF/resistin function in adipogenesis in vivo, we generated transgenic mice overexpressing ADSF/resistin–hFc fusion protein, which functions in a dominant-negative manner [36]. The ADSF/resistin fusion protein not only forms homooligomers, but can also hetero-oligomerize with endogenous ADSF/resistin. The hFc domain probably blocks the interaction of ADSF/resistin with its putative interacting protein or receptor. These transgenic mice showed increased adiposity in a transgene dose-dependent manner, owing to enhanced adipocyte differentiation as well as adipocyte hypertrophy. Interestingly, although resistin null mice were reported to have normal adiposity, loss of resistin in ob/ob mice exacerbated obesity without changes in food intake – a phenotype consistent with the role of ADSF/resistin in inhibition of adipogenesis. In this regard, ADSF/resistin mRNA levels were reported to be at a lower level in adipose tissue of various genetically obese and diet-induced obesity mouse models [37]. However, the circulating ADSF/resistin level was reported to be higher in genetically obese and diet-induced obesity in mice. It is possible the half-life of circulating resistin may be affected in these mice. Regardless, the local concentration of ADSF/resistin in adipose tissue may be critical for its effect during the expansion of adipose tissue. ADSF/resistin mRNA levels are very low in fasting conditions or in streptozotocin-induced diabetes, but are increased drastically by refeeding or by insulin treatment [27], reﬂecting tightly controlled expression of

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ADSF/resistin by nutrition and hormones. Similar ADSF/resistin protein levels in the adipose tissue as well as in circulation by radioimmunoassay conﬁrmed the changes in ADSF/resistin mRNA levels during fasting/feeding as well as by glucose and/or insulin [38, 39]. ADSF/resistin, therefore, could be an adipocyte sensor for the nutritional state of animals and function as a feedback regulator in adipogenesis. The physiological role of resistin and its involvement in obesity-associated insulin resistance has been an ongoing debate. As mentioned above, correlating with insulin resistance, circulating resistin levels are higher in genetic and diet-induced obesity mouse models [38, 39]. Administration of resistin impaired glucose tolerance in normal mice, while a neutralizing resistin antibody improved insulin sensitivity in diet-induced obese mouse models [28]. Furthermore, despite the development of obesity, transgenic mice overexpressing dominant-negative ADSF/resistin fed either a chow or high-fat diet showed enhanced glucose tolerance and insulin sensitivity compared to wild-type mice [36]. These mice also had increased leptin and adiponectin levels in circulation, probably because of increased adiposity and adipogenesis. The complex phenotype of enhanced insulin sensitivity with increased adiposity observed in these transgenic mice could be caused by the chronic effect of the impairment of inhibitory function of ADSF/resistin in adipocyte differentiation. A direct effect of this dominant-negative ADSF/resistin on glucose/insulin homeostasis, however, cannot be ruled out. On a chow diet, mice lacking resistin showed a normal glucose tolerance. However, on a high-fat diet, resistin-deﬁcient mice had improved glucose tolerance, but fasting insulin levels or insulin tolerance were not altered in these mice. Loss of resistin also improved glucose homeostasis in leptindeﬁcient ob/ob mice [40]. The primary target of resistin action appears to be the liver, causing hepatic insulin resistance, with secondary effects on skeletal muscles and adipose tissue [30]. Feeding mice a high-fat diet for 3 weeks brought about an 80% increase in plasma resistin levels accompanying an increase in hepatic glucose production. This hepatic insulin resistance was reversed by resistin antisense oligonucleotide that normalized plasma resistin levels [41]. Acute infusion of resistin reconstituted insulin resistance also. Whole-body insulin resistance involving impaired insulin signaling in skeletal muscle, liver, and adipose tissue, resulting in glucose intolerance and hyperinsulinemia was also reported upon adenovirus-mediated hyper-resistinemia in rats [42]. Contrary to the observed relationship between resistin and insulin resistance, no correlation between either resistin expression or serum levels with serum insulin or glucose levels was reported in obese and insulin-resistant mouse models [43]. These discrepancies may be due to the methodology of measurements or the potentially differing function of various multimeric forms of resistin. Similar to leptin, which has been well documented for its central role in peripheral glucose homeostasis, central infusion of resistin to cerebral ventricle or mediobasal hypothalamus showed an increase in glucose production, that was abrogated in neuropeptide Y null mice suggesting the central function of resistin [44, 45]. Impaired phosphorylation of Akt and AMP-activated protein kinase as well as induction of suppressor of cytokine signaling-3 in hepatic insulin resistance

12.8 Conclusions

by resistin have been reported. However, these changes may represent a consequence of insulin resistance. Identiﬁcation of a potential membrane receptor that mediates resistin action would be necessary to better understand the underlying mechanism. The function of human resistin is even more complex not only with regard to adipogenesis, but also to the metabolic physiology [42, 46]. The biological role of human resistin in glucose metabolism is controversial at best. Several human studies have shown correlation of resistin levels with obesity and insulin resistance, while other studies found no association of resistin with insulin sensitivity or the metabolic syndrome [47–55]. However, it has been reported that resistin levels may be related to the degree of adiposity [56].

12.8 Conclusions

Both obesity and lipodystrophy are commonly associated with diverse pathologies, including diabetes and cardiovascular diseases. It has become evident that adipose tissue is an endocrine organ that secretes a wide variety of factors and dysfunction in secretion affects whole-body glucose homeostasis. Here, we have presented the current knowledge on the function of two adipocyte secreted proteins, Pref-1 and ADSF/resistin. Pref-1 is synthesized and released from preadipocytes in both rodents and humans. ADSF/resistin is secreted from adipocytes in rodents, while, in humans, it is produced from monocytes and macrophages. Although their signaling may differ, these two proteins inhibit adipocyte differentiation and directly or indirectly affect glucose/insulin homeostasis. The inhibitory role of Pref-1 on adipogenesis has been clearly demonstrated in vivo by the generation of Pref-1 null and Pref-1-overexpressing transgenic mice. Pref-1 null mice exhibit increased deposition of adipose tissue, with enlarged adipocytes with increased expression of adipocyte markers. Transgenic mice overexpressing Pref-1 show impaired adipose tissue development and lower insulin sensitivity. By a variety of in vitro approaches using 3T3-L1 cells and Pref-1 null MEFs, Pref-1 has been shown to be processed by TACE to generate a biologically active soluble form and the soluble form prevents differentiation into adipocytes through activation of MEK/ERK pathway. The role of ADSF/resistin in obesity-linked diabetes and cardiovascular diseases is, to date, an ongoing debate. Clearly, identiﬁcation of the receptors through which Pref-1 and ADSF/resistin signaling occur is critical, and would represent a major advance in understanding the molecular mechanisms underlying the function of these adipose tissue secreted factors. Acknowledgments

The work from the authors laboratory was supported by NIH grants to H.S.S. We thank Nabila Aboulaich for her contribution in the initial draft of the review.

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associated dlk. Differentiation, 64, 103–114. Kim, K.A., Kim, J.H., Wang, Y., and Sul, H.S. (2007) Pref-1 (preadipocyte factor 1) activates the MEK/extracellular signalregulated kinase pathway to inhibit adipocyte differentiation. Mol. Cell. Biol., 27, 2294–2308. Moon, Y.S., Smas, C.M., Lee, K., Villena, J.A., Kim, K.H., Yun, E.J., and Sul, H.S. (2002) Mice lacking paternally expressed Pref-1/Dlk1 display growth retardation and accelerated adiposity. Mol. Cell. Biol., 22, 5585–5592. Lee, K., Villena, J.A., Moon, Y.S., Kim, K.H., Lee, S., Kang, C., and Sul, H.S. (2003) Inhibition of adipogenesis and development of glucose intolerance by soluble preadipocyte factor-1 (Pref-1). J. Clin. Invest., 111, 453–461. Schmidt, J.V., Matteson, P.G., Jones, B.K., Guan, X.J., and Tilghman, S.M. (2000) The Dlk1 and Gtl2 genes are linked and reciprocally imprinted. Genes Dev., 14, 1997–2002. Takada, S., Tevendale, M., Baker, J., Georgiades, P., Campbell, E., Freeman, T., Johnson, M.H., Paulsen, M., and Ferguson-Smith, A.C. (2000) Delta-like and gtl2 are reciprocally expressed, differentially methylated linked imprinted genes on mouse chromosome 12. Curr. Biol., 10, 1135–1138. Wylie, A.A., Murphy, S.K., Orton, T.C., and Jirtle, R.L. (2000) Novel imprinted DLK1/GTL2 domain on human chromosome 14 contains motifs that mimic those implicated in IGF2/H19 regulation. Genome Res., 10, 1711–1718. Jensen, C.H., Teisner, B., Hojrup, P., Rasmussen, H.B., Madsen, O.D., Nielsen, B., and Skjodt, K. (1993) Studies on the isolation, structural analysis and tissue localization of fetal antigen 1 and its relation to a human adrenal-speciﬁc cDNA, pG2. Hum. Reprod., 8, 635–641. Laborda, J., Sausville, E.A., Hoffman, T., and Notario, V. (1993) dlk, a putative mammalian homeotic gene differentially expressed in small cell lung carcinoma and neuroendocrine tumor cell line. J. Biol. Chem., 268, 3817–3820.

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H.S. (2001) A cysteine-rich adipose tissuespeciﬁc secretory factor inhibits adipocyte differentiation. J. Biol. Chem., 276, 11252–11256. Steppan, C.M., Bailey, S.T., Bhat, S., Brown, E.J., Banerjee, R.R., Wright, C.M., Patel, H.R., Ahima, R.S., and Lazar, M.A. (2001) The hormone resistin links obesity to diabetes. Nature, 409, 307–312. Holcomb, I.N., Kabakoff, R.C., Chan, B., Baker, T.W., Gurney, A., Henzel, W., Nelson, C., Lowman, H.B., Wright, B.D., Skelton, N.J., Frantz, G.D., Tumas, D.B., Peale, F.V. Jr., Shelton, D.L., and Hebert, C.C. (2000) FIZZ1, a novel cysteine-rich secreted protein associated with pulmonary inﬂammation, deﬁnes a new gene family. EMBO J., 19, 4046–4055. Rajala, M.W., Obici, S., Scherer, P.E., and Rossetti, L. (2003) Adipose-derived resistin and gut-derived resistin-like molecule-beta selectively impair insulin action on glucose production. J. Clin. Invest., 111, 225–230. Blagoev, B., Kratchmarova, I., Nielsen, M.M., Fernandez, M.M., Voldby, J., Andersen, J.S., Kristiansen, K., Pandey, A., and Mann, M. (2002) Inhibition of adipocyte differentiation by resistin-like molecule alpha. Biochemical characterization of its oligomeric nature. J. Biol. Chem., 277, 42011–42016. Chen, J., Wang, L., Boeg, Y.S., Xia, B., and Wang, J. (2002) Differential dimerization and association among resistin family proteins with implications for functional speciﬁcity. J. Endocrinol., 175, 499–504. Banerjee, R.R. and Lazar, M.A. (2001) Dimerization of resistin and resistin-like molecules is determined by a single cysteine. J. Biol. Chem., 276, 25970–25973. Patel, S.D., Rajala, M.W., Rossetti, L., Scherer, P.E., and Shapiro, L. (2004) Disulﬁde-dependent multimeric assembly of resistin family hormones. Science, 304, 1154–1158. Patel, L., Buckels, A.C., Kinghorn, I.J., Murdock, P.R., Holbrook, J.D., Plumpton, C., Macphee, C.H., and Smith, S.A. (2003) Resistin is expressed in human macrophages and directly regulated by

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glycemia. Am. J. Physiol. Endocrinol. Metab. 288, E625–E632. Singhal, N.S., Lazar, M.A., and Ahima, R.S. (2007) Central resistin induces hepatic insulin resistance via neuropeptide Y. J. Neurosci., 27, 12924–12932. Muse, E.D., Lam, T.K., Scherer, P.E., and Rossetti, L. (2007) Hypothalamic resistin induces hepatic insulin resistance. J. Clin. Invest., 117, 1670–1678. Pravenec, M., Kazdova, L., Landa, V., Zidek, V., Mlejnek, P., Jansa, P., Wang, J., Qi, N., and Kurtz, T.W. (2003) Transgenic and recombinant resistin impair skeletal muscle glucose metabolism in the spontaneously hypertensive rat. J. Biol. Chem., 278, 45209–45215. Silha, J.V., Krsek, M., Skrha, J., Sucharda, P., Nyomba, B.L., and Murphy, L.J. (2004) Plasma resistin, leptin and adiponectin levels in non-diabetic and diabetic obese subjects. Diabet. Med., 21, 497–499. Silha, J.V. and Murphy, L.J. (2004) Serum resistin (FIZZ3) protein is increased in obese humans. J. Clin. Endocrinol. Metab., 89, 1977; author reply 1978–1977. Youn, B.S., Yu, K.Y., Park, H.J., Lee, N.S., Min, S.S., Youn, M.Y., Cho, Y.M., Park, Y.J., Kim, S.Y., Lee, H.K., and Park, K.S. (2004) Plasma resistin concentrations measured by enzyme-linked immunosorbent assay using a newly developed monoclonal antibody are elevated in individuals with type 2 diabetes mellitus. J. Clin. Endocrinol. Metab., 89, 150–156. Norata, G.D., Ongari, M., Garlaschelli, K., Raselli, S., Grigore, L., and Catapano, A.L. (2007) Plasma resistin levels correlate with determinants of the metabolic syndrome. Eur. J. Endocrinol., 156, 279–284. Lee, J.H., Chan, J.L., Yiannakouris, N., Kontogianni, M., Estrada, E., Seip, R., Orlova, C., and Mantzoros, C.S. (2003) Circulating resistin levels are not associated with obesity or insulin resistance in humans and are not regulated by fasting or leptin administration: cross-sectional and interventional studies in normal, insulinresistant, and diabetic subjects. J. Clin. Endocrinol. Metab., 88, 4848–4856.

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Lobig, M., and Forst, T. (2003) Evaluation of human resistin assays with serum from patients with type 2 diabetes and different degrees of insulin resistance. Clin. Lab., 49, 571–576. 53 Heilbronn, L.K., Rood, J., Janderova, L., Albu, J.B., Kelley, D.E., Ravussin, E., and Smith, S.R. (2004) Relationship between serum resistin concentrations and insulin resistance in nonobese, obese, and obese diabetic subjects. J. Clin. Endocrinol. Metab., 89, 1844–1848. 54 Utzschneider, K.M., Carr, D.B., Tong, J., Wallace, T.M., Hull, R.L., Zraika, S., Xiao, Q., Mistry, J.S., Retzlaff, B.M., Knopp, R.H., and Kahn, S.E. (2005) Resistin is not

associated with insulin sensitivity or the metabolic syndrome in humans. Diabetologia, 48, 2330–2333. 55 Iqbal, N., Seshadri, P., Stern, L., Loh, J., Kundu, S., Jafar, T., and Samaha, F.F. (2005) Serum resistin is not associated with obesity or insulin resistance in humans. Eur. Rev. Med. Pharmacol. Sci., 9, 161–165. 56 Pagano, C., Marin, O., Calcagno, A., Schiappelli, P., Pilon, C., Milan, G., Bertelli, M., Fanin, E., Andrighetto, G., Federspil, G., and Vettor, R. (2005) Increased serum resistin in adults with Prader–Willi syndrome is related to obesity and not to insulin resistance. J. Clin. Endocrinol. Metab., 90, 4335–4340.

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13 Adipose Tissue and Blood Pressure Regulation Lisa A. Cassis and Sara B. Police 13.1 Introduction

The modern industrialized environment has contributed to an epidemic rise in the prevalence of obesity and obesity-associated disorders, including hypertension. Hypertension affects more than 65 million adult Americans [1]. The majority of patients with hypertension are overweight, with a recently reported odds ratio (95% conﬁdence interval) of 1.73 and 3.39 for overweight and obesity, respectively, for independent association with hypertension [2]. A 10 kg greater body weight is associated with a 3.0 and 2.3 mmHg increase in systolic and diastolic pressure, respectively, increasing the risk of cardiovascular disease by 12% [3, 4]. Early reports in the Framingham Heart Study demonstrated that 78% of essential hypertension in men can be directly attributed to obesity [5]. Given the unabating epidemic of obesity in the United States, it is clear that obesity-related hypertension will increase in the future. Despite a strong association between obesity and hypertension, mechanisms linking the two diseases are not fully understood. Hemodynamic characteristics of obese hypertensive subjects include an increase in intravascular volume, increased cardiac output, endothelial dysfunction, and abnormal kidney function [6–15]. Candidate mediators contributing to these hemodynamic changes in obese hypertensives include activation of the sympathetic nervous system [16–18], the renin–angiotensin system (RAS) [19–21], elevated free fatty acids (for review, see [22]), oxidative stress [23–26], insulin resistance and hyperinsulinemia [27–30], and dysregulated production and function of various adipocyte-derived factors (Figure 13.1). For the purposes of this chapter, we focus on adipose tissue and its role in obesity-related hypertension. 13.2 Adipose Tissue Changes with Obesity: Relation to Blood Pressure Control

During positive energy balance, excess energy is stored in adipose tissue. This mechanism is important in normal physiology, where adipose tissue serves as an

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Obesity

Sympathetic Nervous System

ReninAngiotensin System

Free Fatty Acids

Insulin Resistance Hyperinsulinemia

Adipokines

Hypertension Figure 13.1 Mechanisms contributing to obesity-related hypertension. Obesity could increase blood pressure through multiple mechanisms, including stimulation of the sympathetic nervous system, activation of the

RAS, elevations in circulating concentrations of free fatty acids, insulin resistance and hyperinsulinemia, and elevated expression and release of various adipokines.

energy storage reservoir for the release of lipids during fasting that are oxidized as fuel for skeletal muscle. When energy intake exceeds energy expenditure, however, triglycerides accumulate in adipocytes, leading to adipocyte hypertrophy and hyperplasia. Enlargement of existing adipocytes with lipid is usually the ﬁrst mechanism utilized to handle excess energy stores [31], followed by recruitment of precursor cells to healthy adipocytes that can contribute to additional lipid storage. Dysfunction of adipose tissue with obesity thus represents the combined mechanisms of markedly hypertrophied adipocytes and impaired adipogenesis. Depending on the anatomic location of adipose tissue, these mechanisms are differentially altered with obesity. For example, subcutaneous adipose tissue typically retains the ability to increase proliferation and differentiation of adipocytes to accommodate a positive energy balance, while hypertrophy of the more metabolically active visceral adipose tissue is closely associated with hypertension. The concept of the adipocyte as a simple energy storage site for lipid has evolved considerably, primarily as a result of the discovery of leptin production and secretion by adipocytes [32]. It is now widely recognized that adipocytes produce a variety of factors that are present in the circulation and that can act in an autocrine/paracrine manner to regulate energy homeostasis. The list of factors produced by adipocytes grows longer each year (Figure 13.2) and has spurred the terminology adipokine to describe a variety of factors that exhibit a proinﬂammatory phenotype. With adipocyte hypertrophy, the production of many of these adipocyte-derived factors is dysregulated. Mechanisms for changes in adipokine expression and secretion with adipocyte hypertrophy are not well deﬁned, but have been suggested to include interactions between adipocytes and recruited macrophages, interactions with the extracellular matrix, and changes in cholesterol distribution from the cell

13.2 Adipose Tissue Changes with Obesity: Relation to Blood Pressure Control

Adipocytes

Adipocyte-derived factors

Leptin

Resistin

Adiponectin

TNFα

Angiotensinogen

Glucocorticoids

PAI1

IL6

MCP1

Visfatin

Figure 13.2 Adipocyte-derived factors. Adipocytes produced a variety of factors, termed adipokines due to their ability to promote inflammation. Adipocyte-derived factors can act in an autocrine or paracrine manner. IL6, interleukin-6.

membrane to the triacylglycerol droplet [33, 34]. The consequences of adipocyte hypertrophy on the expression of adipocyte-derived factors are pronounced, and many of these factors, as described below, can contribute to the control of blood pressure (Figure 13.3). 13.2.1 Adipocyte RAS in Obesity-Related Hypertension

We originally reported expression of angiotensinogen, the only known precursor to angiotensin II, in a variety of white and brown adipose tissues from rats [35, 36]. Follow-up studies in our laboratory examined mechanisms for processing of angiotensinogen to angiotensin II in adipocytes and adipose tissue, and demonstrated renin-like and angiotensin-converting enzyme (ACE) activity, and the release of angiotensin peptides from adipocytes [37–39]. Further studies in our laboratory demonstrated regulation of adipose angiotensinogen expression by nephrectomy [40, 41], angiotensin II [42, 43], diabetes [44], obesity [45–48], and cold acclimation [49, 50]. Additional research by a variety of investigators has demonstrated a complete RAS in rodent and human adipocytes (for reviews, see [51–54]). Evidence suggests that the adipocyte RAS may contribute to obesity-related hypertension. In mice with adipocyte-speciﬁc overexpression of angiotensinogen, adipocyte

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Obesity

Adipocyte Leptin RAS

Adiponectin Insulin PAI-1

SNS Endothelial dysfunction

Free fatty Resistin acids

11-βHSD1

RAS Vascular reactivity

Hypertension Figure 13.3 The role of adipokines in obesity-related hypertension. Several different adipokines have the ability to modulate blood pressure through regulation of the sympathetic nervous system (SNS), the RAS, by causing endothelial dysfunction, or by enhancing vascular reactivity.

growth was increased [55]. Importantly, in mice overexpressing angiotensinogen in adipocytes, plasma angiotensinogen concentrations increased and were associated with elevated blood pressure. Unexpectedly, mice with deﬁciency of either AT1a or AT2 receptors exhibit resistance to diet-induced obesity [56, 57]. In mice overexpressing 11bhydroxysteroid dehydrogenase (HSD)-1 selectively in adipose tissue, visceral adiposity was associated with an increase in adipose angiotensinogen expression and the development of hypertension [58]. In mice with diet-induced obesity, angiotensinogen expression was increased in visceral adipose tissue, but not in subcutaneous adipose or nonadipose tissues, but blood pressure was not examined [59]. Results from our laboratory demonstrated a site-speciﬁc increase in angiotensinogen mRNA abundance in a visceral adipose tissue of rats with diet-induced obesity and hypertension [45]. Importantly, obese hypertensive rats exhibited pronounced elevations in systemic angiotensin II concentrations, which correlated positively to adipose angiotensinogen mRNA abundance and to blood pressure [45]. Administration of an AT1 receptor antagonist resulted in a greater reduction in blood pressure in diet-induced obese rats withhypertension comparedtoleancontrols [60]. Additionalstudies examinedmechanisms for regulation of adipose angiotensinogen, focusing on endocrine feedback regulation by the end-product peptide, angiotensin II. Results demonstrated that angiotensin II regulates adipose angiotensinogen mRNA abundance through the AT1a receptor and that in states of high systemic angiotensin II (i.e., obesity), angiotensin II regulation of adipose angiotensinogen expression increases [42]. In humans, angiotensinogen mRNA abundance in omental, but not in subcutaneous adipose tissue correlated positively to waist-to-hip ratios [61]. In obese and lean

13.2 Adipose Tissue Changes with Obesity: Relation to Blood Pressure Control

patients, angiotensinogen gene expression was greater in visceral compared to subcutaneous adipose tissue and was increased in adipose tissue from obese compared to lean subjects [62]. In contrast, other studies have revealed either a slight decrease or no change in angiotensinogen gene expression in adipose tissue from obese compared to lean patients [63–65]. Recent studies in obese menopausal women demonstrated lower angiotensinogen gene expression in adipose tissue; however, expression of angiotensinogen decreased in adipose tissue of obese patients with weight loss [66]. Importantly, reductions in angiotensinogen gene expression with weight loss in obese patients were associated with decreased circulating angiotensinogen concentrations and reductions in blood pressure. Clinical studies comparing obese to lean subjects demonstrated a positive correlation between body mass index (BMI) and plasma angiotensinogen concentrations [19–21, 67, 68]. Plasma renin activity has been reported to increase in obese hypertensives [69] and recent studies demonstrate that direct renin inhibition with Aliskiren results in a reduction in arterial hypertension in obese patients [70]. In addition, a positive correlation between plasma ACE activity and BMI was demonstrated [20], while weight loss in obese subjects resulted in a reduction in plasma ACE activity [71]. In the placebo-controlled TROPHY trial for the treatment of obese patients with hypertension, 66% of patients administered lisinopril exhibited a reduction in diastolic blood pressure below 90 mmHg, compared to 43% of patients administered hydrochlorothiazide.ain [7, 10, 72] Similar results were obtained in the Candesartan Role on Obesity and Sympathetic System study, where the AT1 receptor antagonist candesartan decreased blood pressure, muscle sympathetic nerve activity, and improved insulin sensitivity, while hydrochlorothiazide exhibited favorable effects on blood pressure, but not on the other parameters [73]. Collectively, these results support a role for the adipocyte RAS in human obesity-related hypertension. 13.2.2 Leptin in Obesity-Related Hypertension

Leptin, a peptide hormone secreted by adipocytes, regulates food intake and energy expenditure through effects on the central nervous system. In the brain, leptin binds to the long form of the leptin receptor and activates pro-opiomelanocortin (POMC)/cocaine- and amphetamine-related transcript neurons in the hypothalamus, while inhibiting the activity of neuropeptide Y/agouti-related protein neurons (for review, see [74]). These neurons project to the paraventricular nucleus and lateral hypothalamic area, and to sympathetic preganglionic neurons in the medulla and spinal cord [75]. In addition to stimulating sympathetic outﬂow to thermogenic tissues for the control of energy expenditure, leptin also increases sympathetic activity to cardiovascular-relevant organs, including the kidneys and adrenal gland [76–78]. These effects of leptin have been demonstrated to result in an increase in arterial blood pressure when leptin is administered chronically into the central nervous system [79]. However, while both central and peripheral administration of leptin both stimulate the sympathetic nervous system, only central leptin administration increases blood pressure [76, 78]. These results have suggested that peripheral

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effects of leptin, including stimulation of nitric oxide, may counterbalance the central effects of leptin to stimulate the sympathetic nervous system, resulting in no change in arterial pressure when leptin is elevated peripherally. Many of the effects of leptin are mediated by a product of POMC processing, amelanocyte-stimulating hormone (MSH), which binds to melanocortin (MC)-4/5 receptors in the hypothalamus to elicit the metabolic effects of leptin [80, 81]. Administration of an MC-4/5 receptor antagonist to rats abolished the effects of chronic leptin administration to raise blood pressure [82]. Moreover, in mice with obesity from MC-4 receptor deﬁciency, intravenous leptin infusion did not increase blood pressure, suggesting that a functional MC-4 receptor is essential for the cardiovascular effects of leptin [83]. Circulating leptin levels are considered to be a marker of adiposity and plasma leptin concentrations are increased in obese compared to lean individuals [84]. The presence of obesity despite increased circulating concentrations of leptin has led to the hypothesis that obese humans are resistant to the metabolic actions of leptin [85]. However, in order for leptin to be considered a pivotal adipokine linking obesity to hypertension, the cardiovascular effects of leptin would need to be preserved in obese subjects with high circulating leptin concentrations. This hypothesis is supported by data from yellow obese agouti mice, which are resistant to the anorexigenic effects of leptin, but have an intact sympathetic excitatory response to leptin administration [86, 87]. Metabolic and cardiovascular effects of leptin have also been demonstrated to diverge based on the response of the baroreﬂex, where activation of the baroﬂex selectively inhibits leptin-induced renal sympathetic activation, but has no effect on sympathetic activation to brown adipose tissue [88, 89]. The mechanism for these diverging effects of leptin to increase sympathetic outﬂow for metabolic versus cardiovascular regulation involves differences in the neuronal pathways. For example, antagonists of the MC-4 receptor inhibit leptin-induced activation of renal sympathetic nerves, but have no effect on sympathetic activation to brown adipose tissue, while a corticotrophin-releasing factor receptor antagonist selectively inhibits leptin-induced sympathetic activation to brown adipose tissue [90]. If these data are extrapolated to obesity, it is conceivable that obese individuals have selective leptin resistance to the metabolic effects of leptin, while cardiovascular effects of leptin are maintained. However, this hypothesis has not been directly tested in human obesity. 13.2.3 Adiponectin in Obesity-Related Hypertension

Adiponectin was cloned independently by four groups in 1995 and 1996, and is expressed at the mRNA level predominantly in adipose tissue from experimental animals and humans [91–93]. Adipose tissue serves as the major source of circulating adiponectin, which is present in plasma at concentrations ranging from 3 to 30 mg/ml in adults [94]. Circulating adiponectin exists in two major oligomeric forms of a hexamer and a 400-kDa high-molecular-weight complex [95]. Unlike other adipokines, adiponectin levels in plasma are inversely correlated with BMI [96]. Moreover, the negative correlation of adiponectin with body weight becomes stronger when

13.2 Adipose Tissue Changes with Obesity: Relation to Blood Pressure Control

correlated to visceral, as compared to subcutaneous, adipose tissue [97]. Mechanisms for reductions in adipose expression of adiponectin with obesity are unknown; however, tumor necrosis factor (TNF)-a has been suggested as a contributor due to its ability to inhibit adiponectin promoter activity [98]. The signiﬁcance of reductions in circulating adiponectin with obesity arise from its ability to act as an insulin sensitizer [99–102], and anti-inﬂammatory and antiatherogenic protein [103, 104]. Circulating levels of adiponectin have been reported to decrease in patients with hypertension [105] and hypoadiponectinemia has been demonstrated as an independent risk factor for hypertension in cross-sectional studies [106, 107]. Moreover, in recent studies prospectively examining the 5-year development of hypertension in a nondiabetic Chinese cohort, an inverse relationship between plasma adiponectin concentration and the future development of hypertension was deﬁned [108]. Surprisingly, the predictive nature of plasma adiponectin concentrations with hypertension remained signiﬁcant even after adjustment for BMI. These results suggest that adiponectin may play a role in the pathogenesis of essential hypertension, including obesity-related hypertension. Mechanisms potentially contributing to hypertension in patients with hypoadiponectinemia include increased plasma fatty acid concentrations and insulin resistance, endothelial dysfunction from loss of nitric oxide [109], dysregulated vascular smooth muscle growth with resulting vascular hypertrophy [110, 111], and loss of suppression of renal sympathetic nerve activity [112]. Interestingly, recent studies demonstrate that adenoviral delivery of adiponectin to obese KKAy mice decreased blood pressure, while adiponectin deﬁcient mice exhibited hypertension, supporting an in vivo role for adiponectin in obesity-related hypertension [113]. 13.2.4 Insulin and Obesity-Related Hypertension

In addition to regulating glucose uptake into adipocytes, insulin increases lipid storage by promoting adipocyte differentiation, inhibiting lipolysis, and stimulating lipogenesis. With insulin resistance at hypertrophied adipocytes, lipolysis is increased and plasma levels of nonesteriﬁed fatty acids (NEFAs) rise. Elevations in NEFA have been demonstrated to increase sympathetic nerve activity, promote insulin resistance at skeletal muscle, cause endothelial dysfunction, increase oxidative stress in variety of cell types, and enhance vascular smooth muscle sensitivity to contractile agonists. In addition, insulin has direct effects through insulin receptors to regulate a variety of cardiovascular-relevant functions. At the vascular endothelium, insulin stimulates the production of nitric oxide [114] through insulin receptor tyrosine kinase-dependent phosphorylation of insulin receptor substrate-1 leading to Akt activation and phosphorylation of endothelial nitric oxide synthetase [115, 116]. At vascular smooth muscle, insulin decreases contractility associated with reductions in membrane RhoA [117], by inhibiting calcium inﬂux and promoting calcium efﬂux [118]. In the heart, insulin regulates a variety of metabolic processes (glucose uptake, glycolysis, lipid metabolism, protein synthesis), enhances cardiac contractility [119–122], and through several insulin signaling pathways can contribute to

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cardiac hypertrophy [118]. Moreover, a variety of studies demonstrate that physiological concentrations of insulin increase sympathetic nerve activity [123–125]. Given that insulin has opposing effects to increase vasodilation, but yet promote vasoconstriction, the net hemodynamic effect of insulin on blood pressure is minimal in healthy subjects. In states of insulin resistance, speciﬁc signaling pathways utilized by the insulin receptor are impaired, with decrements in phosphoinositide-3-kinase signaling and retained Ras/mitogen-activated protein kinase (MAPK)-dependent signaling effects of insulin [126, 127]. Interestingly, many of the prohypertensive effects of insulin occur through MAPK signaling, such that in states of hyperinsulinemia insulin may contribute to hypertension. 13.2.5 Plasminogen Activator Inhibitor-1 and Obesity-Related Hypertension

Plasminogen activator inhibitor (PAI)-1 is the major physiological inhibitor of tissuetype plasminogen activator and urokinase-type plasminogen activator, and as such inhibits ﬁbrinolysis. PAI-1 concentrations in plasma are elevated in type 2 diabetes, even after correction for insulin levels [128, 129]. The source of elevated PAI-1 with obesity is controversial, since both adipocytes and stromal vascular cells (i.e., macrophages) express PAI-1 [130–132]. In addition, stromal vascular cells in visceral adipose tissue express considerably more (5-fold) PAI-1 compared to subcutaneous adipose tissue [133]. However, plasma concentrations of PAI-1 are more closely related to fat accumulation and PAI-1 expression in the liver than in adipose tissue, suggesting the liver serves as a major source of PAI-1 production [134]. Several different mechanisms have been suggested to contribute to elevated PAI-1 expression with obesity, including TNF [135, 136], transforming growth factor-b [137, 138], oxidative stress [139], free fatty acids [140], and angiotensin II [141–143]. In mice with PAI-1 deﬁciency, both diet-induced [144, 145] and genetic obesity [146] are decreased, suggesting that PAI-1 can contribute to the development of obesity. Moreover, pharmacologic inhibition of PAI-1 results in a reduction in weight gain and improved insulin sensitivity in mice [147, 148]. While obesity-induced increases in PAI-1 expression have been mechanistically linked to insulin resistance and atherosclerosis, there have been no studies directly examining the role of PAI-1 in obesity-related hypertension. 13.2.6 Free Fatty Acids and Obesity-Related Hypertension

The release of free fatty acids from abdominal adipocytes increases with central obesity [149, 150]. Free fatty acids are released from adipocytes following hormonesensitive lipase (HSL)-mediated hydrolysis of triacylglycerol, and represent the major secretory product of adipocytes [151]. The activity of HSL is stimulated by badrenergic agonists, and decreased by insulin. With insulin resistance, activity of HSL increases, with a resulting release of free fatty acids from adipocytes into the

13.2 Adipose Tissue Changes with Obesity: Relation to Blood Pressure Control

systemic circulation. A variety of studies, performed in animals and humans, demonstrate that infusion of Intralipid results in an increase in blood pressure (for review, see [22]). Elevations in circulating levels of free fatty acids can inﬂuence blood pressure through multiple mechanisms. Elevations in blood pressure following infusion of oleic acid to rats were blunted by an a1-adrenergic antagonist [152]. Conversely, infusion of Intralipid in hand veins of normotensive subjects resulted in a lower dose of phenylephrine required to produce 50% of the maximal vasoconstrictor response [153], suggesting that free fatty acids promote a-adrenergic responsiveness. These data suggest that enhancing the responsiveness to a1-adrenergic agonists may serve as a mechanism linking elevated fatty acids to hypertension. Free fatty acids reduce endothelium-dependent production of nitric oxide and studies have demonstrated that infusion of Intralipid to lean, healthy individuals results in a reduction in methacholine-induced relaxation [154]. Moreover, elevated free fatty acids have been suggested to serve as a mechanism for impaired vasodilator responses to insulin [155]. Another potential mechanism linking elevations in free fatty acids to hypertension with obesity includes an increase in oxidative stress; however, this mechanism has not been directly tested in humans. Free fatty acids exhibit mitogen actions on vascular smooth muscle, potentially related to activation of protein kinase C and potentiation of the responsiveness to other growth factors (insulin-like growth factor-1) [156–160]. Finally, free fatty acids have been reported to activate the RAS, as evidenced by increased expression of angiotensinogen in adipocytes [161], enhanced mitogenic responses to angiotensin II [157, 162], and stimulation of the production of aldosterone from rat adrenal cells [163]. Collectively, these results suggest that elevations in circulating concentrations of free fatty acids could contribute to obesityrelated hypertension through multiple mechanisms. 13.2.7 Resistin and Obesity-Related Hypertension

Resistin was identiﬁed by three independent groups [164–166] and is a predominantly adipocyte-derived protein that is detectable in the systemic circulation [164]. In rodent models of obesity resistin levels were elevated in serum [164], and some studies have shown elevations in resistin with human obesity and diabetes [167]. However, conﬂicting data has also been reported, showing either a reduction in serum resistin levels and decreased resistin mRNA expression in adipose tissue from genetically obese db/db mice [168], or no change [169]. Resistin has been suggested to play a role in the regulation of glucose homeostasis. Administration of resistin protein resulted in a reduction in insulin action in C57BL/6 mice [164] and SpragueDawley rats [170]. Interestingly, resistin has been demonstrated to increase the production of the proinﬂammatory cytokines endothelin-1 and monocyte chemotactic protein (MCP)-1 in endothelial cells [171, 172], and stimulate foam cell formation in macrophages [173, 174]. However, it is unclear whether these effects of resistin contribute to a role of this protein in the pathogenesis of obesity, atherosclerosis, or diabetes. In prehypertensive subjects with normal BMI, plasma levels of resistin were increased [175]. Moreover, in patients with essential

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hypertension and type 2 diabetes mellitus, serum resistin levels were increased [176]. However, a role for resistin in the development of obesity-related hypertension in animal models or humans has not been deﬁned. 13.2.8 11b-HSD-1 and Obesity-Related Hypertension

The effect of cortisol at target tissues is modulated by the tissue-speciﬁc enzyme 11bHSD-1. The type 1 form of 11b-HSD is expressed by adipocytes and facilitates the conversion of inactive cortisone to active cortisol. The type 2 form of 11b-HSD catalyzes the metabolism of cortisol locally to cortisone, minimizing the effects of glucocorticoids and allowing actions at the mineralocorticoid receptor. 11b-HSD-1 is expressed to a greater extent in omental than subcutaneous adipose tissue [177] and thus contributes to the differential effects of glucocorticoids on regional adipose depots. In adipocytes, by promoting the formation of active cortisol, 11b-HSD-1 promotes the differentiation of preadipocytes to adipocytes. Transgenic mice overexpressing 11b-HSD-1 in adipocytes exhibit elevated plasma corticosterone, visceral obesity, and several aspects of the metabolic syndrome, including hypertension [58]. Interestingly, hypertension in these mice was associated with activation of the RAS and blood pressure was decreased by an AT1 receptor antagonist. Conversely, inhibition of 11b-HSD-1 in obese rodents improves glucose tolerance, insulin sensitivity, and lipid proﬁles [178]. While a role of 11b-HSD-1 in hypertension has been demonstrated in several experimental models, the enzyme does not appear to be as clearly linked to hypertension in humans [179].

13.3 Regional Adipose Deposition and Blood Pressure Regulation

In 1956, Vague ﬁrst described different patterns of fat distribution, coined android for upper-body obesity and gynoid for lower-body obesity [180]. As far back as 1956, android obesity was found to be more closely related to diabetes mellitus and coronary artery disease. It is generally well accepted that central adiposity (android obesity) is associated with greater cardiovascular risk than gynoid obesity. Subcutaneous fat comprises approximately 80% of total body fat [181], but is generally less metabolically active as compared to visceral adipose tissue [181, 182]. Both genetic and hormonal (sex hormones, corticosteroids, growth hormone, insulin, etc.) mechanisms may regulate the regional distribution of adipose tissue. With obesity, both subcutaneous and visceral adipose tissue increase in mass; however, the greater metabolic activity of visceral adipose tissue is more closely associated with metabolic disease. Using computed tomography to measure abdominal, thoracic and thigh fat, Hayashi et al. [183] demonstrated a signiﬁcant odds ratio of hypertension with increasing quartiles of intra-abdominal fat, which remained after adjusting for total subcutaneous fat area or waist circumference in Japanese Americans with a high degree of abdominal fat given their BMI. In Dallas Heart Study participants,

13.3 Regional Adipose Deposition and Blood Pressure Regulation

dual-energy X-ray absorptiometry scanning assessment of fat in the trunk area correlated positively with systolic blood pressure [184]. Recent studies in participants from the Framingham Heart Study using multidetector computed tomography assessment of both subcutaneous and visceral adipose tissue demonstrated that visceral adipose tissue was more strongly associated with an adverse metabolic risk proﬁle even after accounting for standard anthropometric indexes [185]. Mechanisms for these close associations of visceral adipose tissue to metabolic risk, including hypertension, are unknown, but are thought to relate to the greater metabolic activity of visceral adipose tissue which then drains into the portal circulation [186]. 13.3.1 Changes in Visceral Adipose Tissue in Obesity-Related Hypertension

With the exception of leptin and HSL, each of the adipokines described above is expressed in greater abundance in visceral compared to subcutaneous adipose tissue. For example, in young rats, mRNA abundance of approximately 20 genes, including HSL and angiotensinogen, was greater in perirenal visceral as compared to subcutaneous adipose tissue [187]. Similar results have been obtained using human adipose tissue [181, 188, 189]. Moreover, using the clamp technique to control delivery of nutrients to adipose tissue, glucose uptake was associated with an increased expression of resistin, adiponectin, leptin, PAI-1, and angiotensinogen in visceral, but not subcutaneous, fat [190]. However, only a few adipokines with regional variations in expression have been examined in the context of obesityrelated hypertension. We demonstrated a site-speciﬁc increase in mRNA abundance of angiotensinogen in rats with diet-induced obesity and hypertension [45]. Administration of an AT1 receptor antagonist to rats with diet-induced obesity hypertension resulted in normalization of blood pressure, supporting a role for the RAS in obesity-related hypertension [60]. In transgenic mice expressing 11bHSD-1 in adipose tissue, visceral obesity developed on normal mouse diet and was exacerbated by high-fat feeding [191]. Importantly, these mice developed hypertension which was associated with activation of the RAS [58]. To date, results support a role for increased expression and activity of speciﬁc adipokines in visceral adipose tissue as a potential link between obesity and hypertension. However, without methodologies to control the depot-speciﬁc expression of adipokines, it is difﬁcult to directly deﬁne the role of regional expression of adipokines in obesity-related hypertension. 13.3.2 Potential Role for Perivascular Adipose Tissue in Obesity-Related Hypertension

Following our original observation that adipose tissue, including periaortic adipose tissue, expressed angiotensinogen as the precursor to angiotensin II, we examined the role of periaortic adipose tissue as a potential modulator of the contractile response to a variety of agonists [192]. The general consensus prior to this time was

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that periarterial adipose tissue served primarily a mechanical, rather than a metabolic function [193, 194]. Our results demonstrated that in rat aortic rings with adherent periaortic adipose tissue, the contractile response to norepinephrine was reduced, supporting a role for periaortic adipose tissue as modulator of vascular responsiveness. Since this time, a variety of investigators have demonstrated that adipose tissue surrounding blood vessels serves a physiological and/or pathophysiologic role. In addition, alterations in the function of perivascular adipose tissue have been suggested to contribute to various diseases, including hypertension. In 2002, Lohn et al., [195] demonstrated that perivascular adipocytes secreted a factor, termed adventitial-derived relaxing factor (ADRF), which acts by a tyrosine kinasedependent mechanism to activate potassium channels in vascular smooth muscle. The release of ADRF resulted in attenuated vascular responses to angiotensin II, serotonin, and phenylephrine. Further studies focused on ADRF release from resistance arteries demonstrated hyperpolarization of vascular smooth muscle in rat mesenteric arteries by periadventitial fat [196], and that these effects were diminished in mesenteric arteries from spontaneously hypertensive rats [197]. Additional studies demonstrated that conditioned media from rat perivascular adipose tissue stimulates the proliferation of vascular smooth muscle cells and that with obesity, these effects are augmented [198]. Finally, recent studies in lipoatrophic A-ZIP/F1 transgenic mice demonstrated that enhanced contractile responses of blood vessels to agonists was associated with an absence of perivascular adipose tissue, potentially contributing to hypertension [199]. Interestingly, the mass of perivascular adipose tissue increased markedly along the length of the aorta when rats were fed a high-fat diet [200] and was suggested to contribute to obesity-associated atherosclerosis through stimulation of macrophage recruitment to adipose tissue [201]. Collectively, these results suggest that expanded mass of adipose tissue surrounding cardiovascular relevant organs, including blood vessels, may contribute to obesityrelated diseases, including hypertension. Future studies should address the pathophysiological relevance of periorgan adipose tissue in obesity-associated disease.

13.4 Conclusions

Adipose tissue is no longer a simple energy-storing site, but is capable of the production of a variety of factors that can inﬂuence blood pressure. Alterations in adipose production and release of proteins with obesity may contribute to obesityassociated diseases, including hypertension. Future areas for investigation include deﬁnition of the mechanisms for altered adipokine expression with obesity and the relative contribution of adipokines to obesity-related hypertension. The use of adipocyte-speciﬁc promoters to deplete adipokines in mice should facilitate our understanding of the role of these adipose-derived factors in obesity and obesityrelated hypertension.

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14 Adipokines, Inflammation, and Obesity Karine Clement 14.1 Introduction

Links between the inﬂammation of adipose tissue and its potential role in systemic complications of obesity were proposed in 1993 by Hotamisligil et al. [1]. In a study ﬁrst carried out in the mouse, this group showed that white adipose tissue (WAT) synthesizes a proinﬂammatory cytokine called tumor necrosis factor (TNF)-a, the expression of which was elevated in adipocytes of insulin-resistant and obese mice. The insulin sensitivity in the animals could be improved using TNF-a antibodies. These ﬁrst observations underlined the existence of a tie between a proinﬂammatory cytokine, produced and secreted by adipose tissue, and the development of insulin resistance in rodents. However, the neutralization of TNF-a did not appear to be very effective in controlling insulin resistance in humans. Nevertheless, a novel ﬁeld of research was opened linking inﬂammation and obesity domains that took on an even greater dimension after the discovery of the adipocyte hormone, leptin, in 1994. Leptin is a cytokine produced and secreted by WAT, which has not only an essential role in the control of energy metabolism via its role on hypothalamic pathways, but is also a key player in many metabolic activities in peripheral tissues (including the liver, pancreas, muscles and immune system) [2]. The discovery of leptin paved the way to the concept of adipocytokines or adipokines that was proposed to qualify the small proteins synthesized and secreted by adipose tissue. They are prone to circulate in the plasma and eventually to exert a systemic inﬂuence. In this context, they can interfere with the cross-talk between organs [3]. Among these proteins, a distinction has to be made between the molecules speciﬁcally or exclusively produced by adipose tissue depots and other molecules produced in abundance by tissues other than adipose tissue (like the liver). Within adipose tissue itself, one can still distinguish between adipokines produced speciﬁcally by adipocytes (like leptin, adiponectin, and serum amyloid A (SAA)), and those produced by other cell types of the nonadipocyte fraction. The identiﬁcation in 2003 that macrophages accumulate in adipose tissue of obese subjects has made inﬂammatory cells major contributors to adipokine production in adipose tissue.

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Excellent reviews have been published on this subject in the past few years, detailing particularly the roles and functions of adipokines [4, 5]. In this chapter, we cover certain aspects showing the contribution of adipose tissue in producing adipokines that may inﬂuence local and systemic biology.

14.2 Contribution of Adipose Tissue in Systemic Inflammation during Obesity

Obesity is characterized by the augmentation of adipose tissue mass; however, according to individual characteristics, adipose tissue depots are more or less abundant in certain regions of the organism; in the upper part of the body in android obesity, in the lower part of the body in gynoid obesity. The distribution of adipose mass in the upper part of the body confers an increased risk for obesity complications. The increase in fat mass, especially in the abdominal region, is usually associated with a moderate but chronic increase in circulating levels of inﬂammatory mediators. They include nonspeciﬁc markers like C-reactive protein (CRP) [6, 7], acute-phase inﬂammatory proteins, and proinﬂammatory cytokines. Adhesion and remodeling molecules of the extracellular matrix are part of these systemic changes since they can also exert, among many functions, pro- and anti-inﬂammatory, and prothrombotic effects [7–10] (Table 14.1). The liver and lymphoid organs are usual sites of production of these mediators. Recent developments have shown that adipose tissue secretes some of these active molecules and in this way might contribute to their systemic increase during the development of obesity (Table 14.1). These molecules represent a very heterogeneous panel of factors with diverse, complex, and sometimes redundant roles. They can participate in innate, adaptive immunity, growth, hematopoiesis, and metabolism, among others. The number of molecules known to be produced by adipose tissue is increasing regularly, especially because of the developments in the utilization of analytical systems exploring large-scale gene expression (DNA chips) or proteins secreted by the tissue depots [11, 12]. The most studied proteins, historically involved in interorgan cross-talk that connects weight homeostasis, insulin sensitivity, and many other key biological systems, remain leptin [13] and adiponectin [14]. In contrast to leptin, known as a proinﬂammatory mediator, adiponectin has compelling anti-inﬂammatory functions and is an insulin-sensitizer molecule. Other molecules like visfatin [15], resistin [16], or omentin [17] have also been described in adipose tissue of mice and humans. Mainly secreted by the visceral adipose tissue, they are suggested to contribute to the development of insulin resistance (resistin, visfatin) or, in contrast, to facilitate insulin action (omentin). Studies were mostly performed in cells or in rodents. Nonetheless, there is still a lot to be discovered regarding these adipokines in human adipose tissue biology, particularly their effective role in insulin sensitivity [18] since non-negligible differences might exist between humans and rodents. The potential role of the recently discovered molecules has recently been discussed in detail [19].

14.2 Contribution of Adipose Tissue in Systemic Inflammation during Obesity Modification of circulating inflammatory or adhesion markers during the course of obesity and weight loss brought about by diet.

Table 14.1

Name

Nonspeciﬁc markers CRP ﬁbrinogen orosomucoid Acute phase of inﬂammation haptoglobin SAA Cytokines/ILs IL-6 IL-8 IL-18 IL-10 IL-1Ra TNF-a MCP-1 MCP-4 MIF Other adipokines leptin visfatin resistin adiponectin omentin Adhesion proteins/extracellular matrix remodeling/ prothrombotics MMP-9 ICAM VCAM hepatocyte growth factor PAI-1 cathepsin S

Systemic modification

Effect of weight loss

Visceral (vis) versus subcutaneous (sc)

increase increase increase

decrease decrease decrease

increase increase

decrease decrease

vis > sc sc > vis

increase increase increase increase increase increase increase increase increase

decrease decrease decrease decrease decrease decrease or 0 decrease ? decrease

discussed vis > sc vis > sc vis ¼ sc not clear vis ¼ sc vis > sc ? vis ¼ sc

increase increase diminution diminution diminution

decrease decrease augmentation or 0 increase ?

sc > vis vis > sc (discussed) vis > sc (discussed) vis > sc vis sc

increase increase increase increase increase increase

decrease decrease decrease ? decrease decrease

— — — vis > sc vis ¼ sc

The expression and secretion of a myriad of other adipokines belonging to different family classes have been identiﬁed in adipose tissue (Table 14.2, A and B). They include members of the cytokine family, such as interleukins IL-1, IL-1Ra [20–22], IL-8 [23–25], IL-18 [26], and IL-10 [27, 28], growth factors like transforming growth factor (TGF)-b [29], proteins secreted in the acute phase of inﬂammation (IL-6, plasminogen activator inhibitor (PAI)-1 [8], haptoglobin, SAA [30], or the chemokines (monocyte chemoattractant proteins (MCP)-1, -3, and -4), angiopoietin, metallothionein), macrophage inﬂammatory protein (MIP)-1a) [31], and complement factors [32]. Other active biomolecules like retinol-binding protein-4 tend to be synthesized by the

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Table 14.2 Cytokine families expressed or secreted by the human adipose tissue.

Cytokine

Identified in the human adipose tissue (example)

References

Chemokines

MCP-1/CCL2 MCP-3 MCP-4 RANTES/CCL5 MIP-1a IL-6 IL-8 IL-1Ra IL-10 IL-18 Interferon family (IP-10/CXCL10) TNF-a vascular endothelial growth factor isoforms TGF-b hepatocyte growth factor leptin

[99] [12] [62] [106]

ILs

Interferons TNF family Growth factors

Others

[107] [108] [109] [11] [48] [110] [1] [111] [43] [112] [2]

WATand to be linked with insulin resistance [33, 34], although its precise role has been debated [35]. Some molecules were originally found in rodents or in mouse adipocyte models [5], and others were identiﬁed while studying the gene expression proﬁle of human adipose tissue. Using this technique, our group demonstrated, for example, that different isoforms of SAA, particularly the inducible forms SAA-1 and -2, can be produced by human adipose tissue. SAA was usually thought to be produced by the liver. SAA circulating levels increase in obese subjects, and correlate with adipose tissue and adipocyte size [30]. Proteins like the cysteine proteases of the cathepsins family also belong to the adipokine family since they are synthesized and secreted in excess by adipose tissue throughout obesity [36]. Obesity is therefore associated with a moderate but chronic increase of a cocktail of inﬂammatory factors and a decrease of production of some factors like adiponectin. The relative contribution of the different tissues (hepatic, the lymphoid system, subcutaneous and visceral adipose tissues, and eventually the muscle) in the delivery of adipokines to the systemic circulation is technically difﬁcult to examine and even more so during the different phases of obesity development. Even though the expression of a large number of cytokines is found in adipose tissue and some may inﬂuence its local biology, this does not necessarily mean that these molecules are signiﬁcantly secreted in the circulation to exert a major systemic role [37]. 14.3 Adipose Tissue Depots and Adipokine Production

Adipose tissue in the upper part of the body is a usual risk factor of type 2 diabetes, hypertension, dyslipidemia, and cardiovascular diseases [38]. Glucose intolerance is

14.4 Adipokines and Adipose Tissue Cell Types

signiﬁcantly more common in subjects with abdominal obesity than in those with fat mass accumulation in the lower part of their body. Plasma triglyceride levels are also signiﬁcantly more elevated in individuals with central obesity. By contrast, excess adipose tissue in the lower part of the body (i.e., gynoid obesity) is not associated with major metabolic consequences[39, 40]. Adipose tissue possesses anatomical speciﬁcities, particularly regarding adipocyte functions and roles (lipogenesis, lipolytic activity, and expression of developmental genes, hormonal response to insulin or to catecholamines, to sexual hormones, or to cortisol) as reviewed in [41]. The proﬁle of adipokine secretion is part of these dissimilar characteristics between the subcutaneous and visceral tissues. Leptin is preferentially secreted by the subcutaneous adipose tissue [42], while the expression of adiponectin, PAI-1, IL-8, IL-1b, MCP-1, and TGF-b is higher in visceral fat. By contrast, there are controversial reports concerning IL-6 and TNF-a appears to be equally synthesized by the different sites [43–48]. Visfatin, resistin, and omentin were shown to be preferentially expressed in the visceral tissue in humans [31, 49], but this was recently debated for visfatin [50]. It is important to mention that in severe forms of obesity in which all adipose tissue stores increase in volume, the part played by the visceral or the very abundant subcutaneous adipose tissue in the systemic delivery of inﬂammatory mediators is still not well known. Nevertheless, the distinct proﬁle of adipokine secretion between visceral and subcutaneous adipose tissues probably contributes to the increased risk of metabolic and cardiovascular complications, and to the development of other complications like hepatic steatosis and nonalcoholic steatosis in obese individuals. Finally, other adipose tissue depots in the so-called ectopic sites may contribute to the production of inﬂammatory mediators in the absence of obesity. In this regard, the local production of inﬂammatory molecules by epicardial adipose tissue and its proximity with coronary vessels could contribute to the development of coronary pathologies [51].

14.4 Adipokines and Adipose Tissue Cell Types

The adipose tissue is a heterogeneous tissue composed of several cell types – the mature adipocytes and many other cells (preadipocytes, ﬁbroblasts, endothelial cells, histiocytes, and macrophages) grouped together in the stromal vascular fraction (SVF). The various cell types remain to be precisely characterized as well as the relative change in their phenotypes and proportional number in line with the accumulation in human adipose tissue depots. Owing to this heterogeneity, the cellular source of inﬂammatory factors secreted by adipose tissue has been discussed. In vitro studies showed primarily that isolated adipocytes can express inﬂammatory factors like TNF-a [52]. The acute-phase protein SAA is overexpressed and secreted in abundance by isolated adipocytes of obese subjects, as is leptin, whereas adipocyte secretion of adiponectin diminishes in insulin-resistant and obese individuals. SAA and leptin production by adipose tissue depends on adipocyte size [30, 53, 54]. Adipocyte size inﬂuences the expression of inﬂammatory mediators

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as demonstrated by fractionation studies of adipocytes coupled with studies of gene expression proﬁles [53]. Adipocyte size, for example, determines the secretion of adipokines like IL-6 and IL-8 secreted by the most hypertrophic adipocytes, but also of MCP-1 and granulocyte colony-stimulating factor [55]. The size of adipocytes may inﬂuence the metabolism and obesity complications since it was shown, for example, that adipocyte hypertrophy precedes the development of type 2 diabetes [56]. Nevertheless, an increasing number of studies agree that the principal production site of inﬂammatory mediators is cells of the SVF [57]. By separating mature adipocytes from the stromal fraction, we characterized functional expression proﬁles of the SVF showing an enrichment of functions related to immunity and inﬂammation while the adipocyte proﬁle is mostly metabolic [58]. Studies published by North American and French groups prove that adipose tissue of obese subjects or animals is a site of major macrophage accumulation, which is proportional to the body mass index and adipocyte hypertrophy [31, 59, 60]. Macrophages of adipose tissue are undoubtedly a source of adipokine production, as described for visfatin and resistin [31]. Other cell types, such as endothelial cells, inﬂammatory cells (lymphocytes, natural killer (NK) cells), and preadipocytes, can also contribute to the production of adipokines, notably when exposed to a local inﬂammatory microenvironment. A study carried out in rodents shows the modiﬁcation of NK cell populations and lymphocyte subpopulations in mice fed with a high-fat diet [61]. Markers of activated T cells have also been identiﬁed in adipose tissue of obese subjects [62]. Studies are underway to characterize cell phenotypes, their origin, and their secretory capacities in human, especially in the different fat depots. It was proposed that macrophages and mature adipocytes show close gene expression proﬁles. In addition, adipocytes could exert macrophage-like properties, especially when exposed to a proinﬂammatory environment [63, 64]. However, the transplantation of bone marrow into irradiated mice has established that the vast majority of macrophage inﬁltration in adipose tissue originates from the circulation. In obese subjects, these macrophages typically aggregate in crowns around the adipocytes [27]. Macrophages express activation markers, but are they in a pro- or anti-inﬂammatory state? The macrophage phenotypes could be pro- or anti-inﬂammatory depending on the degree of obesity and of its evolution, as suggested by studies in mice showing that weight gain is accompanied by transformation from a macrophagic M2 (anti-inﬂammatory) phenotype towards an M1 (proinﬂammatory) proﬁle [65]. Consequently, secretion proﬁles of adipose tissue can change depending on the phenotype of the cell population inﬁltrating it during the different stages of obesity (installation, aggravation, maintenance, weight loss).

14.5 Adipokines, Macrophages, and the Biology of Adipocytes

Adipose tissue macrophages may contribute to maintaining the low-grade inﬂammatory state linked to obesity (Figure 14.1). The factors that induce the

14.5 Adipokines, Macrophages, and the Biology of Adipocytes

Figure 14.1 Typical aspect of macrophage infiltration in obese subjects. Note the distribution in rosettes of the macrophages, which are not seen on the nonobese, but which are more enhanced in the deep (omental)

tissue. A, adipocyte. The macrophages are stained with HAM56 antibody. These macrophages synthesize many cytokines locally. SC, subcutaneous.

inﬁltration and activation of macrophages in adipose tissue are probably multifactorial. The paracrine, autocrine, and endocrine signals as well as mechanical modiﬁcations (hypertrophy and adipocyte hyperplasia) could play a role in this phenomenon. 14.5.1 Chemoattraction

Many adipokines synthesized by adipose tissue are candidates to attract inﬂammatory cells. In vitro studies suggest that leptin itself at supraphysiologic levels induces adhesion proteins, hence facilitating the migration of monocytes [66]. Conversely, adiponectin inhibits this process, but only in models of aortic endothelial cells [67]. This potential action still needs to be demonstrated with adipose tissue cells. Very little is known about the role of selectins, integrins, and elements of adhesion to the extracellular matrix in the process of macrophage attraction in adipose tissue. Many adipokines could exert a stimulatory or inhibitory effect on the recruitment of macrophages into adipose tissue. The large-scale analysis of gene expression that can be done with DNA chips allows the identiﬁcation of candidate genes. For example, we observed in human studies that the levels of expression of MCP-1, colony-stimulating factor-3 and of the urokinase plasminogen activator (CD87) increase signiﬁcantly in adipose tissue of subjects with morbid obesity [27].

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MCP-1 (also known as CCL2) is a strong chemoattractant cytokine and it acts via its receptor CCR2. CCR2 can also link other molecules like MCP-3/CCL7. In a ﬁrst study, CCR2 (CCR2/) gene knockout mice show a reduction of macrophage inﬁltration in adipose tissue and improvement of insulin sensitivity. This led to the suggestion that the CCR2 receptor and the MCP-1 cytokine are major players in macrophage accumulation within adipose tissue [68]. Meanwhile, recent contradictory data suggest that MCP-1 might not be such a crucial candidate since knockout mice for CCR2 that have the same level of macrophage inﬁltration as the wild-type gain more weight and are insulin-resistant [69]. Other candidate molecules and other mechanisms are on track to be explored. Local hypoxia could also play an important role in the attraction and retention of macrophages within adipose tissue. We have shown that hypoxia-inducible factor (HIF)-1a, a transcription factor normally induced by hypoxia, is overexpressed in subcutaneous adipose tissue of obese subjects and that this expression is decreased during weight reduction. Tissue hypoxia induces macrophage attraction in solid tumors as well as in atheromatosed plaques. These observations suggest that adipose tissue of obese subjects could be hypoxic in some areas and a local expression of chemokines could be induced. It should be noted that leptin, which possesses indirect chemoattractant properties, is itself a known factor of induction of the HIF-1 gene [70]. 14.5.2 Paracrine Cross-Talk in the Adipose Tissue via Adipokines

Does the macrophage accumulation in adipose tissue really have a detrimental role? In other words, could macrophage accumulation be related to an adaptation process associated with the augmentation of fat mass? In such circumstances, macrophage accumulation could be necessary for the upkeep of the tissue and perhaps aim to limit its expansion? A recent study has established that macrophagic aggregates within adipose tissue are localized around the dead adipocytes, suggesting that one of their functions is to clean up the debris of necrotic cells [71]. We have previously shown, using electron microscopy, that adipocytes surrounded by macrophages have indeed granules of lipofuscin in their cytoplasm, suggesting accelerated aging of the cells [27]. In addition to their role in cleaning up old and useless cells, the accumulation of macrophages may be also useful for the formation of new vessels, particularly at the site of inﬂammation and in ischemic zones when adipose tissue grows. Macrophages could also control fat mass growth and modify the biology of adipocytes and preadipocytes via the local production of adipokines. This is demonstrated by several studies. The speciﬁc effects of adipokines on different adipocyte functions (increased lipolysis, modiﬁcations of adipocyte secretion, and induction of adipocyte insulin resistance eventually overcome by thiazolinediones) have been shown for TNF-a and IL-6 [72]. However, the SVF cells produce cocktails of inﬂammatory molecules. The consequence of local inﬂammation on preadipocyte biology has thus been studied. In the presence of a medium derived from human

14.5 Adipokines, Macrophages, and the Biology of Adipocytes

Figure 14.2 Cross-talk between macrophages, preadipocytes, and adipocytes via the chemokines and adipokines.

macrophages, human preadipocytes show a drastic change in their phenotype, acquiring a proinﬂammatory phenotype by secreting signiﬁcant amounts of IL-6 or IL-8; they grow [73], but differentiate badly [73, 74]. These cellular alterations are induced in cocultures without direct cellular contact, suggesting the key role of soluble factors (Figure 14.2). The mature adipocyte also endures profound modiﬁcations of its biology in coculture systems with activated macrophage medium. Other than the proinﬂammatory state, an increased lipolysis and a resistance to insulin are observed [75, 76]. TNF-a has been proposed to mediate these effects. From the molecular point of view, the nuclear factor-kappa B (NF-kB) pathway, implicated in the primary regulation of inﬂammatory responses, is induced in the preadipocyte [73] as in the adipocyte in the presence of medium from macrophages [77]. The NF-kB pathway is also brought into play in macrophages in contact with a medium from adipocytes. The Toll-like receptors (TLR-4) appear to be important actors that lead to the induction or suppression of genes orchestrating the inﬂammatory response. TLRs are a large family of receptors, including TLR-4, identiﬁed in the mid-1990s as a bacterial lipopolysaccharide receptor (endotoxin). Recent data have shown that free fatty acids produced by adipocytes after adrenergic stimulation are also strong inducers of TLR-4/NF-kB [77]. TLR-4 is hence expressed by the adipocytes and overexpressed during obesity [78]. Therefore, fatty acids produced in excess by the adipocytes, ﬁlled with triglycerides, and stimulated by the inﬂammatory medium, activate TLR-4, thus inducing these inﬂammatory changes. TLR-4 knockout mice are protected from these deleterious effects, especially from insulin resistance induced by lipid infusions [79].

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Based on these different studies, a dual effect of macrophages of adipose tissue could be expected: (i) a local beneﬁcial effect in clearing out the old adipocytes, and in the control and in limiting the development of fat mass, and (ii) in a simultaneous manner, a deleterious systemic effect via the increase in the production and secretion of adipokines, easing the production and progression of complications of obesity and the induction of insulin resistance.

14.6 Adipokines and Complications of Obesity

Inﬂammatory molecules could be candidates to exert a molecular link between adipose tissue and the metabolic, cardiovascular, hepatic, thrombotic complications and certain cancer types occurring during the natural evolution of human obesity. A myriad of candidate adipokines are proposed to play this role. The potential inﬂuence of adipokines on the complications of obesity is the subject of regular updating reports [80, 81]. In the cardiovascular ﬁeld, they can be considered as cardiovascular risk factors and even play a direct physiopathological role, eventually favoring the installation and aggravation of atherosclerosis. In terms of the relationship between abnormalities of cardiac function in obese subjects, the accumulation of visceral fat and low-grade inﬂammation has been suggested. Among the candidates secreted by adipose tissue, the increase in IL-6, IL-8, and MCP-1, and the decrease of adiponectin, are cited frequently. The studies of the physiopathological links between adipokines and cardiac function can be illustrated by the analysis of adiponectin effects in rodents. Adiponectin can indeed diminish the size of lesions observed in acute ischemic myocardial infarctus, exert angiogenic properties, and lower the atherogenesis plaque in genetically predisposed mice (apolipoprotein E/ invalidation) with an overexpression of adiponectin [82]. In the metabolic ﬁeld, several chemokines known to be produced by adipose tissue, like RANTES (Regulated upon Activation of Normal T cells Expressed and Secreted), IL-1b, and IL-8, as well as factors of oxidative stress, are increased in diabetic or glucose-intolerant patients and the amelioration of glycemia by insulin therapy reduces circulating levels of these molecules [83, 84]. The increase in the levels of TNF-a, IL-6, IL-1b, IL-8, resistin, and many other factors produced by macrophage activation could contribute to the deterioration of insulin sensitivity. Nonetheless, the precise relationship between the importance of macrophage accumulation in adipose tissue depots, adipokine secretion, and modiﬁcations of insulin sensitivity also need to be addressed in humans. Macrophage accumulation is clearly more abundant in visceral tissue [58] than in subcutaneous tissue and this could explain some of the risks associated with the accumulation of intra-abdominal fat. Our group has shown, for example, a relationship between the increase of macrophages in visceral adipose tissue and hepatic ﬁbroinﬂammation [58]. In a recent study, expression levels of MCP-1 and colonystimulating factor-1 genes and proteins were also associated with this macrophage accumulation in obese subjects [85]. Since the visceral adipose tissue is drained by the

14.7 Adipokines and Weight Loss

portal system, it cannot be excluded that some adipokines delivered by adipose tissue could contribute to the alteration of hepatic function observed in obese subjects, the mechanism of which needs to be investigated. The relationships between the newly identiﬁed adipokines, like cathepsin S and the SAA, and the complications of obesity are still to be evaluated in clinical studies. Furthermore, many differences may exist between humans and rodents, notably whether some of these molecules have speciﬁc actions in the development of obesity complications.

14.7 Adipokines and Weight Loss

Even modest weight reduction is well known to improve the metabolic and cardiovascular risks associated with human obesity. Measures of endothelial stress also improve after weight reduction in women [86–88]. In this context, many studies showed that a decrease in food intake, and sometimes an increase in exercise, reduce global systemic inﬂammation. A reduction of numerous inﬂammatory molecules and endothelial risk factors, and an increase of adiponectin have been observed during weight-loss programs, and sometime relates to the improvement of insulin sensitivity [89]. This has been described for CRP [90], IL-6 [91], IL-18 [92], IL1-Ra [22], PAI-1 [93], SAA [30, 94], cathepsin S [95], matrix metalloproteinase (MMP)-9 [96], adhesion molecules (intercellular adhesion molecule (ICAM) and vascular cell adhesion molecule (VCAM)) [88], tissue factor [97], macrophage inhibitory factor (MIF) [98], MCP-1 [99], soluble receptor factors of TNF receptor-1, and eotaxin, an inﬂammatory factor implicated in asthma – another complication of obesity [100]. Weight loss induced by gastric bypass also induces a reduction in the circulating levels of visfatin [101] and this has also been discussed for TNF-a [102, 103]. Exercise does not always have a major effect on the modiﬁcation of these parameters [104]. The dynamics, amount of weight loss, and amelioration of insulin sensitivity can sometimes, but not always, modify these circulating parameters. We followed 60 obese patients during the course of weight loss induced by bariatric surgery leading to a reduction of 30% of initial weight. A signiﬁcant decrease in CRP, SAA, orosomucoid protein, IL-6, TNF-a, and ﬁbrinogen, and an increase in the level of circulating adiponectin was observed. After the surgery, the level of IL-6 decreased slowly while the level of SAA and/or CRP decreased very quickly. On the other hand, the variation of SAA was not associated with the rapid change in insulin sensitivity [105]. Are these systemic changes of inﬂammatory-related molecules associated with signiﬁcant modiﬁcations of inﬂammatory gene expression in adipose tissue? We observed a signiﬁcant modiﬁcation in the expression of inﬂammatory genes in the subcutaneous adipose tissue of obese women following a hypocaloric diet [11]. The expression of 100 genes linked to inﬂammation processes was modiﬁed after 4 weeks (41% increased and 59% decreased). These genes belong to at least 12 functional

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families, including cytokines, interleukins, the complement factor cascade, and acute-phase proteins of inﬂammation, and of certain molecules playing a role in cellular adhesion and in the remodeling of the extracellular matrix. The improvement of the inﬂammatory proﬁle is not only associated with decreased expression of the proinﬂammatory factors, but also with the increase in the expression of IL-10 or IL1Ra – molecules having anti-inﬂammatory properties. The modiﬁcation of the inﬂammatory gene expression proﬁle was very similar in subjects during bariatric surgery and was associated with a reduction of macrophage inﬁltration. In this study the protein expression of IL10 also increased, suggesting a possible M1/M2 (proinﬂammatory to anti-inﬂammatory) switch of macrophage phenotypes [27]. Overall, the improvement of the systemic inﬂammatory proﬁle observed during weight loss is associated with modiﬁcations of adipose tissue proﬁles in inﬂammatory gene expression. The eventual consequence of this phenomenon on the local biology of adipose tissue remains to be identiﬁed.

14.8 Conclusions

The discoveries of a chronic low-grade inﬂammatory state in obesity and the inﬁltration of macrophages in adipose tissue have opened new perspectives in the search for the physiopathologic mechanisms that can explain the development of the complications linked with obesity, their evolution, and their persistence. A novel objective would be the identiﬁcation of molecular mechanisms of adipose tissue with the goal of controlling the interaction between macrophages and adipocytes and other tissues. Eventually, other biomarkers must also be identiﬁed, produced speciﬁcally by adipose tissue during obesity, leading to comorbidities in the obese patient. Acknowledgments

The author would like to thank the Agence Nationale de la Recherche (Programs RIOMA and ObCat), lAssistance Publique Hôpitaux de Paris/Direction de la Recherche Clinique (PHRC, CRC), and lINSERM et lUniversite Pierre et Marie Curie Paris 6 (BQR Paris 6) and the European Commission (7th framework ADAPT) for project fundings related to inﬂammatory processes in obesity, and also Daniele Lacasa, Joan Tordjman, Raffaella Cancello, and Froogh Hajduch for their reading and contribution to the manuscript.

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Part Four Adipose Tissue and Disease

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15 Depot-Specific Biology of Adipose Tissues: Links to Fat Distribution and Metabolic Risk Mi-Jeong Lee and Susan K. Fried 15.1 Introduction

An abdominal or so-called central pattern of fat distribution of body fat is tightly correlated with complications of obesity, such as insulin resistance, dyslipidemia, hypertension, hypercoagulation, and chronic inﬂammation. The clinical and epidemiological literature show strong associations of abdominal obesity (high waist circumference) with adverse health outcomes, while lower-body fat stores typical of females are protective [1–3]. Many clinical studies that used computerized tomography (CT) or magnetic resonance imaging (MRI) to distinguish visceral and subcutaneous abdominal depots point to a strong negative effect of visceral fat mass on metabolic risk [4], while other studies show independent associations with abdominal sc fat [2]. The production of metabolic products (fatty acids, glycerol, lactate), adipokines, and other proteins secreted from the adipocyte varies among fat depots and may link regional fat to metabolic risk [5]. Thus, increasing research attention has focused on the physiological, biochemical, and molecular determinants of regional differences in adipose tissue development, and the metabolic and endocrine functions of different adipose depots. Body fat is mainly stored within the adipocytes (fat cells) of adipose tissues that collectively deﬁne the adipose organ (i.e., a group of tissues that performs a speciﬁc function or group of functions, as deﬁned in medical dictionaries) and includes numerous discrete anatomical depots [6]. Adipocytes are a highly specialized cell type that store energy in the form of triglyceride and releases it as required by the energy needs of other cells [6, 7]. The adipocyte is also an endocrine cell that secretes several hormones, most notably leptin and adiponectin, which modulate fuel metabolism, immunity, and reproduction [8]. Our objectives in this chapter are to summarize (i) the phenotypic characteristics of different fat depots (i.e., including morphological, physiologic, and metabolic characteristics), and (ii) the molecular underpinnings of depot differences in metabolic and endocrine function.

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15.2 Adipose Depots: Definitions

Subcutaneous adipose tissue, located just under the skin, stores more than 85% of the total body fat. Different locations within the subcutaneous depot are morphologically (e.g., fat cell size and number) and functionally heterogeneous. The most commonly deﬁned and studied subcutaneous depots are the abdominal, femoral, and gluteal. The fascia superﬁcialis (Scarpas fascia), a layer of connective tissue visible on MRI or CT, demarcates deep from superﬁcial subcutaneous fat [9]. The thickness of the deep subcutaneous layer is of particular interest, as it independently correlates with metabolic complications of obesity in both men and women. Intra-abdominal fat depots are associated with internal organs, and in humans are divided into intra- and retroperitoneal. Intraperitoneal depots are associated with digestive organs, and include the omental (hangs off the stomach), mesenteric (associated with the intestine), and epiploic (along the colon). These visceral fat depots are characterized by the presence of lymph nodes and milky spots, distinguishing them from other intra-abdominal depots such as the perirenal and retroperitoneal, as well as subcutaneous [10]. There are also numerous smaller depots such as epicardial that share an embryological origin with the other visceral depots and may subserve specialized functions [11, 12].

15.3 Physiological and Anatomical Differences among Depots may Drive Functional Heterogeneity 15.3.1 Depot Differences in Cellular Composition

Functional differences between depots appear to be related to the cellular composition as well as variations in blood ﬂow [13] and innervation [14]. Possibly as a result of these physiological differences, as well as possible intrinsic differences in fat cells from different depots [15, 16], the average fat cell size and distribution varies among depots [17]. Adipocyte size is of importance because it is related to adipocyte metabolic and endocrine function (e.g., leptin production) [18, 19]. Triglyceride turnover is higher in larger adipocytes – basal lipolysis and triglyceride synthesis (fatty acid esteriﬁcation) are both increased. As basal rates are higher, relative responses (fold change from basal) to the stimulatory effect of catecholamines on lipolysis and the insulin stimulation of glucose uptake and metabolism decline. In women, lower-body subcutaneous adipocytes tend to be larger than subcutaneous abdominal ones, which are in turn larger than omental adipocytes. Depot differences in adipocyte size are less apparent in men [20, 21]. In addition to adipocytes, adipose tissue is composed of ﬁbroblasts, preadipocytes, stem cells, mast cells, endothelial cells, and pericytes, as well as immunocytes (neutrophils, dendritic cells, T cells, lymphocytes, macrophages). Omental adipose

15.3 Physiological and Anatomical Differences among Depots may Drive Functional Heterogeneity

tissue also includes mesothelial cells [22]. The number of stromal vascular cells per gram of adipose tissue is greater in omental compared to subcutaneous adipose tissue [23, 24]. Depending on the level of obesity, adipose tissue includes substantial numbers of resident macrophages, especially in the omental depot of humans. These are often found in crown-like structures surrounding dead adipocytes during periods of tissue remodeling [25]. The higher number of macrophages in omental compared to subcutaneous adipose tissue is likely the major source for the higher production of cytokines from this depot [26–28], but other stromal cells may also contribute [29]. 15.3.2 Definition of Visceral Fat Depots

The literature on human adipose tissues frequently deﬁnes visceral depots as portally draining, as this anatomical relationship is thought to drive the metabolic complications associated with visceral obesity [30, 31]. Thus, this deﬁnition would include only omental and mesenteric depots. The rationale for distinguishing these depots based on their anatomical location is that the metabolic (fatty acids, glycerol, lactate) and secretory products (e.g., adipokines) of these depots are predicted to be high within the portal vein, and thereby have a large impact on hepatic function, including hepatic glucose production and very-low-density lipoprotein triglyceride secretion (Figure 15.1). However, the experimental basis for this assumption is considered weak [32]. A recent study found only a higher concentration of interleukin (IL)-6 in

Figure 15.1 Consequences of portal drainage of visceral adipose tissue. It has been hypothesized that fatty acids and adipokines released from visceral adipose tissues (omental and mesenteric) may directly reach the liver in high concentrations via the portal vein. As a result, higher production of IL-6, and perhaps other adipokines, may promote hepatic

production of very-low-density lipoprotein (VLDL) and hypertriglyceridemia, and impair insulin sensitivity, contributing to hyperglycemia. In contrast, free fatty acids (FFAs) released from subcutaneous fat enter the systemic circulation where they reach muscle and can be oxidized, reducing concentrations reaching the liver.

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the portal compared to peripheral blood of humans [33], and other studies calculate no difference in portal free fatty acid concentrations [34]. Interestingly, there was a strong association of IL-6 and circulating C-reactive protein (CRP) – a marker of chronic inﬂammation that is produced mainly in the liver. Thus, it seems likely that the disproportionate inﬂuence of portally draining adipose tissues is related to their impact on hepatic metabolism, although the mediator(s) are as yet undeﬁned. Pond [35] suggests that lymph node containing depots are functionally distinct and play a special role in immunity. As compared to subcutaneous depots, visceral depots within the peritoneal cavity of humans (omental, mesenteric) contain a greater abundance of milky spots where immunocytes accumulate and these undoubtedly contribute to the overexpression of immune-related proteins [36].

15.4 Heterogeneity in Adipocyte Function among Adipose Depots 15.4.1 Lipolysis

A number of studies conducted over the past 30 years indicated that omental adipocytes (as a typical visceral adipocyte) are more lipolytic compared to subcutaneous adipocytes and this idea has become ingrained in the literature [37–40]. Compared to abdominal subcutaneous adipocytes, omental adipocytes of women exhibit lower basal (spontaneous) lipolysis and higher incremental (as percentage or delta over baseline) responses to mixed adrenergic agonists, due mainly to the lower expression of antilipolytic a2-adrenergic receptors [40, 41]. However, maximally stimulated rates of lipolysis do not differ. In addition, omental adipocytes were reported to be less sensitive to the antilipolytic effect of insulin [42]. Most of these studies were carried out on a relatively small number of subjects who were not wellcharacterized with respect to level of visceral adiposity. Recently, Tchernof et al. analyzed the lipolytic function of omental compared to abdominal subcutaneous adipocytes of premenopausal women [21] and men [20] across a wide range of visceral adiposity. They found similarly sized adipocytes and no regional differences in basal lipolysis or in the absolute rates of b-adrenergic-stimulated lipolytic agonists in men, and this was consistent across subjects with low and high waist circumference. In their study of premenopausal women, they found smaller omental fat cells with lower basal lipolysis and no differences in rates of stimulated lipolysis, although due to the lower basal, the response to an adrenergic agonist was relatively increased (fold stimulation). Similarly, Reynisdottir et al. [43] found that the higher maximally stimulated lipolysis observed in abdominal subcutaneous rather than omental adipocytes is associated with higher hormone-sensitive lipase activity in subjects over a wide range of body mass index (BMI). However, the differences could be accounted for by the smaller size of omental adipocytes. These data agree with previous reports in morbidly obese subjects [44, 45].

15.4 Heterogeneity in Adipocyte Function among Adipose Depots

With regard to heterogeneity in lipolysis between femoral, gluteal, and abdominal subcutaneous depots, many but not all studies indicate that lower-body adipocytes are more responsive to the antilipolytic effects of a2-adrenergic agonists, resulting in lower lipolytic responses to mixed agonists that are consistently documented in in vitro studies [46]. A study of premenopausal women indicated that gluteal adipocytes were more sensitive to the antilipolytic effect of insulin, but that basal lipolysis did not differ [47]. However, our preliminary studies do not show this depot difference in postmenopausal women [48]. Thus, it is tempting to speculate that estrogen may differentially regulate adipocyte metabolism in gluteal/femoral compared to abdominal subcutaneous depots and the change in fat distribution after menopause [49]. Animal studies support the hypothesis that estrogen regulates regional adiposity [50, 51]. Further work on the mechanisms by which sex steroids regulate fat distribution are clearly needed. Fewstudies have addressed invivo factors that could affect adipocyte lipolytic activity. Differential sympathetic innervation of visceral depots may mediate the preferential mobilization of these depots under hypocaloric conditions [14], although this possibility has not been documented in humans. The rate of removal of lipolytic products can also affect the rate of lipolysis in vivo. Virtanen et al. [52] found that blood ﬂow per kilogram of fat tended to be higher in omental than abdominal subcutaneous or perirenal depots of both lean and obese humans, but differences were not statistically signiﬁcant with a relatively small sample size (10 in each group). In rats, mesenteric adipose tissue blood ﬂow is around 5-fold greater than other depots [53, 54]. These physiologic factors merit further study as determinants of adipocyte function. Recent studies point to the importance of the lipid droplet-associated proteins and lipases in the regulation of adipocyte lipolysis, but their role in depot differences in adipocyte metabolism are not yet clear [55, 56]. Both Wang et al. [57] and Ryden et al. [58] found lower mRNA, but not protein expression, of perilipin in human omental compared to abdominal subcutaneous adipose tissue. This result suggests that depot differences in perilipin mass are unlikely to explain the lower basal lipolysis in omental fat and are consistent with the lack of depot difference in maximally stimulated lipolysis. The expression of intracellular fatty acid-binding protein (FABP)-4 (adipocyte lipid-binding protein (ALBP); aP2) at the mRNA and protein levels is higher in subcutaneous than omental adipose tissue of obese subjects, while FABP-5 (keratinocyte lipid-binding protein (KLBP)) was lower in subcutaneous adipose tissue [59]. Additionally, the ALBP: KLBP ratio was higher in subcutaneous compared to omental of both lean and obese. ALBP levels have been associated with insulin resistance and lipolytic rates by some [60, 61], but not all studies [62], thus further analysis of the roles of these proteins in regulating depotspeciﬁc alterations in fatty acid ﬂux are warranted. 15.4.2 Triglyceride Deposition

A major mechanism regulating the rate of triglyceride synthesis and storage in adipocytes is the activity of the enzyme lipoprotein lipase (LPL). LPL hydrolyzes

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circulating triglycerides in chylomicra and very-low-density lipoproteins to fatty acids that can cross the endothelium, and be transported and esteriﬁed by adipocytes. Our early studies showed lower LPL activity per adipocyte in the omental depot of premenopausal severely obese subjects [63], and a recent study showed that this depot difference applies to women across a range of adiposity and fat distribution [21]. In contrast, LPL activity per fat cell is similar in omental and abdominal subcutaneous adipose tissue of severely obese men [63], while in nonobese or moderately obese subjects, omental fat cells are larger and LPL activity is higher (most markedly in men with intermediate waist circumference) [20]. Thus, in men, higher LPL may favor the preferential fat deposition in visceral depots [20]. Binding of the adipocyte secretory product acylation stimulation protein (ASP, C3a), which promotes triglyceride synthesis, is higher in subcutaneous than omental adipose tissue [64]. However, glucose transporter 4 (GLUT4) expression and glucose transport were also higher in the omental depot of severely obese women [65]. Higher glucose uptake would be expected to lead to higher rates of fatty acid esteriﬁcation to match a hypothesized higher rate of lipolysis. However, in vivo studies do not support this hypothesis. A recent study by Jensens group [66] analyzed regional meal fatty acid uptake in obese premenopausal women. They found that in women visceral fat accounted for only approximately 5% of meal fat disposal irrespective of visceral fat mass, while with increasing thigh fat mass there was a proportional increase in meal fatty acid uptake. These results strongly suggest that mechanisms that regulate fat deposition play an important role in regulating regional fat mass. They are also consistent with the conclusion that in obese women, visceral fat is not hyperlipolytic in vivo, but further studies in men are required [66].

15.4.3 Glucose Uptake and Insulin Action

Our early studies showed that on a per fat cell basis, there were no differences in the in vitro glucose uptake and conversion to triglyceride among omental, mesenteric, and abdominal subcutaneous adipose tissues of men, but lower uptake in omental adipose tissue (with smaller adipocytes) of women [67]. Recent in vivo [52] and in vitro [68] studies could document no depot differences in glucose uptake or insulin action in visceral compared to abdominal subcutaneous adipose tissue, while others ﬁnd evidence for insulin resistance in omental adipose tissue [42]. Surprisingly, considering the high levels of inﬂammatory cytokines and activation of mitogenactivated protein kinases (MAPKs) in omental adipose tissue, factors usually associated with insulin resistance, the insulin stimulation of Akt did not differ in omental and abdominal subcutaneous adipose tissue. Thus, we speculate that a unique function of the omental depot protects it from insulin resistance; high concentrations of the insulin-sensitizing visceral-speciﬁc adipokine omentin [69] being one possible mediator.

15.6 Search for Novel Adipokines with Depot-Specific Expression that Link Regional Adiposity

15.5 Regional Differences in Adipose Tissue Gene Expression and Protein Production: Relationship to the Metabolic Syndrome

It is becoming increasingly clear that adipocytes function to integrate nutrient sensing pathways, inﬂammation, and immunity [70]. Thus, a mechanism that probably evolved to promote survival and adapt to undernutrition may lead to chronic inﬂuence in the face of chronic overnutrition, with pathophysiological consequences on metabolism, ultimately leading to diabetes, stroke, atherogenesis, and premature death. In obese compared to lean, and most markedly in visceral compared to subcutaneous depots, levels of proinﬂammatory cytokines are elevated and levels of antiinﬂammatory cytokines such as adiponectin and IL-10 are diminished, as reviewed in several excellent recent papers [8, 71, 72]. Adipokines produced within adipose tissues have local paracrine and endocrine effects. Proinﬂammatory cytokines, especially tumor necrosis factor (TNF) and IL-6 [73, 74], can impair insulin action on target tissues, decreasing insulin effects on glucose uptake into skeletal muscle and hepatic glucose output, contributing to the development of insulin resistance and type 2 diabetes. Adipokines can also act in a paracrine fashion by interfering with insulin signaling in adipose tissue. Inﬂammatory cytokines increase expression levels of p38 MAPK, extracellular signal-regulated kinase, c-Jun kinase-1, and IkB kinase-b at the mRNA and protein (total and phosphorylation) levels in adipose tissue. Consistent with higher proinﬂammatory cytokine levels, these MAPKs levels were 1.5- to 2.5-fold higher in omental versus subcutaneous fat [68]. Increased MAPK activity is known to impair insulin signaling at the insulin receptor substrate level [75].

15.6 Search for Novel Adipokines with Depot-Specific Expression that Link Regional Adiposity to Metabolic Risk

To identify new factors that may explain the association between visceral adiposity and metabolic disease produced from excess visceral compared to subcutaneous adipose tissue, several groups have searched for genes or secretory protein products that are differentially expressed between visceral and subcutaneous. In addition to proinﬂammatory cytokines, visceral adipose tissues overexpress many molecules involved in innate immunity and the acute-phase response, including serum amyloid A (SAA), retinol-binding protein (RBP) and complement factors [76–78] (Table 15.1). Yang et al. recently identiﬁed omentin, a C-type lectin, as a visceral-speciﬁc adipokine that is secreted from visceral, but not subcutaneous, adipose tissue. Omentin increases insulin sensitivity in human adipocytes [69]. Furthermore, plasma levels and mRNA expression of omentin in visceral tissue are decreased in obese, insulin-resistant subjects as compared to nonobese [79]. Another visceral adipokine, visfatin (also known as pre-B cell colony-enhancing factor-1 and nicotinamide phosphoribosyltransferase) was initially identiﬁed as a visceral-speciﬁc adipokine [80]. However, subsequent studies showed similar expression levels between

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Table 15.1 Genes known to be differentially expressed between omental (om) and subcutaneous (sc) adipose tissues.

Gene

Adipose hormones/cytokines/chemokines leptin [120, 121] adiponectin [122] RBP4 [76, 77] angiotensinogen angiotensin II [121] TNF-a [123, 124] IL-6 [125] IL-8 [126] IP-10 [127] MCP-1 [128] macrophage inﬂammatory protein-1a a plasminogen activator inhibitor-1 [26] RANTESa GM-CSFa resistin [130] visfatin [80, 81] omentin [69, 131] TSP-1 [110] Growth factors VEGF [26] HGFa ﬁbroblast growth factor-1 and -9 [132] Extracellular matrix VIM [113] SPARC (osteonectin) [113] ANXA5 [113] Adipogenesis/organogenesis C/EBP-d [112] frizzled 7 [112] NR2F1 [112] HoxA5 [112] HoxC8 [112] Tbox-15 [112] HoxA10 [112] HoxC6 [112] C/EBP-a [112] PPAR-c [113, 129] Metabolism FABP-4 [113] Perilipin [113] CD36 [113] GLUT4 [65] glycogen synthase [113]

Differential expression

Role

om < sc om sc om > sc om > sc

energy/immune and so on energy/anti-inﬂammatory ? blood pressure

om > sc om > sc om > sc om sc om > sc om > sc om > sc om > sc om > sc om > sc om sc om sc om > sc

proinﬂammatory proinﬂammatory proinﬂammatory anti-inﬂammatory chemotactic proinﬂammatory coagulation proinﬂammatory proinﬂammatory proinﬂammatory insulin signaling? insulin signaling? angiogenesis

om > sc om > sc om > sc

angiogenesis angiogenesis ?

om < sc om < sc om < sc

? ? ?

om > sc om > sc om > sc om > sc om > sc om > sc om < sc om < sc om < sc om < sc

adipogenesis adipogenesis organogenesis organogenesis organogenesis organogenesis organogenesis organogenesis adipogenesis adipogenesis

om < sc om < sc om < sc om > sc om < sc

lipid metabolism lipid metabolism lipid metabolism glucose metabolism glucose metabolism (continued)

15.7 Importance of Adipose Tissue Macrophages and other Immunocytes Table 15.1

(Continued)

Gene

Differential expression

Role

cholesteryl ester transfer protein [121] Complement factors C2 [36] C3a, ASP [36, 121] C4 [36] C7 [36] Factor B [36] adipsin (Factor D) [36] Others carboxypeptidase E [110]

om < sc

lipid metabolism

om > sc om > sc om > sc om > sc om > sc om < sc

immune immune/lipid metabolism immune immune immune immune

om > sc

prohormone processing

a

Our unpublished observation.

depots [81, 82]. Intracellularly, visfatin regulates NAD þ levels and therefore works with sirtuins to regulate cell survival after nutritional or environmental stress [83]. Visfatin is also secreted through a nonclassical pathway [84] and is found in the circulation. The role of omentin and visfatin in obesity-related diseases remains largely unknown, although correlations between serum levels of these proteins and obesity and insulin resistance markers have been noted [79, 81]. Both omentin and visfatin are expressed in stromal vascular fraction and the cell type(s) expressing these proteins have not yet been identiﬁed. Adipose tissue, speciﬁcally the adipocyte, has been identiﬁed as a major producer of circulating SAA [78, 85] in individuals without acute infection. SAA is an acutephase reactant that is an independent risk factor for coronary artery disease [86]. SAA is primarily associated with high-density lipoprotein (HDL) and may inhibit reverse cholesterol transport [87–89]. Thus, SAA has been suggested as a link between obesity and coronary artery diseases. Furthermore, SAA is known to activate the chemoattractic formyl peptide receptor like-1, which results in migration of blood monocytes and neutrophils [90]. Adipose tissue has also been identiﬁed as a site of expression of the inﬂammatory marker CRP [91] and other proteins, including resistin (expressed in monocytes) and fasting-induced adipose factor (peroxisome proliferator-activated receptor (PPAR)-c angiopoietin related, expressed in fat cells), but their importance is as yet unclear. However, portally derived proinﬂammatory cytokines such as IL-6 can induce hepatic CRP production.

15.7 Importance of Adipose Tissue Macrophages and other Immunocytes in Regional Adipose Tissue Dysfunction

Recent research has focused on the importance of macrophage accumulation within obese adipose tissue as a mediator of inﬂammation and adipose tissue remodel-

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ing [92]. Two inﬂammatory cytokines that have been implicated in monocyte/ macrophage chemotaxis and activation, monocyte chemoattractant proteins (MCP)-1 and granulocyte-macrophage colony-stimulating factor (GM-CSF)-1 are expressed at higher levels in omental versus subcutaneous, leading macrophage inﬁltration into visceral adipose tissue. The involvement of MCP-1 and its receptor CCR2 in macrophage inﬁltration into adipose tissue has been demonstrated [93]. Macrophage inﬁltration is increased in omental compared to subcutaneous in all groups including lean controls and this difference is exaggerated in viscerally obese [94]. Preferential macrophage inﬁltration into omental was mainly observed in a subgroup in whom obesity was associated with impaired glucose tolerance (IGT). Further, there was a stronger association of macrophage inﬁltration into omental versus subcutaneous with obesity comorbidities, IGT, triglyceride, HDL-cholesterol, and blood pressure. They also showed that signiﬁcant association between age and percent omental macrophages. In ﬂammatory cytokines produced by nonadipocytes such as TNF-a, IL-6, and MCP-1 can impair insulin signaling [73, 75], decrease glucose metabolic capacity, increase lipolysis, and alter adipokine production [95–100]. These effects are likely to mediate the association of macrophage inﬁltration with systemic insulin resistance and, ultimately, risk of type 2 diabetes. Although most studies to date have focused on macrophages, increasing evidence points to the importance of other immunocytes within adipose tissue. RANTES (Regulated upon Activation of Normal T cells Expressed and Secreted) and its receptor CCR5 were signiﬁcantly increased in diet induced obese mouse and human obese adipose tissue [101]. Adipose conditioned medium, especially culture medium from adipose tissue of obese animals, induced T cell migration while monoclonal antibodies to block RANTES signiﬁcantly reduced T cell chemotaxis, suggesting RANTES/CCR5 may be important in leukocyte inﬁltration. Systemic RANTES concentration was higher in individuals with IGT [102] compared to normal subjects, and RANTES levels are higher in visceral than subcutaneous adipose tissue (mRNA and secretion; [101] and our unpublished observation), providing yet another potential link of visceral adiposity to systemic inﬂammation. In addition to factors involved in inﬂammation and immunity, an ever-growing and wide-ranging list of proteins that regulate lipid and lipoprotein metabolism, coagulation, and blood pressure are synthesized within adipose tissue and released systemically [8]. Also, bioactive factors released by adipose tissue include fatty acids, monobutyrin [103], prostaglandins, and steroids are either synthesized de novo or converted in adipose tissue and released into the blood stream [8, 104]. Adipose tissue secretes a number of angiogenic factors, including, hepatic growth factor (HGF), leptin, and vascular endothelial growth factor (VEGF), and these act in an autocrine or paracrine manner in adipose tissue, but may also act as endocrine hormones. Adipogenesis is closely related to angiogenesis; adipose tissue growth occurs with weight gain and requires the formation of new capillaries. Interestingly, angiogenic inhibitors are known to decrease obesity and ameliorate metabolic proﬁles [105, 106]. HGF expression is increased in obesity [107] and in vitro, omental adipose tissue secretes more HGF than subcutaneous adipose tissue (our unpublished observation). HGF has angiogenic, mitogenic, and antiapoptotic effects.

15.8 Gene Expression Profiles are Providing New Insights on Regional Adipose Growth

Circulating levels of HGF are associated with metabolic syndrome, cardiovascular, and cancer risk [108, 109]. Adipose tissue also secretes angiogenesis inhibitors, adiponectin, and thrombospondin (TSP)-1. Expression levels of TSP-1 are higher in omental than subcutaneous adipose tissue [110]. It is evident that interactions between different cell types within tissue modulate depot function. Studies directed at better deﬁning these networks are critical to understanding the depot-speciﬁc biology of adipose tissues.

15.8 Gene Expression Profiles are Providing New Insights on Regional Adipose Growth and Function

Characterization of differences in gene expression between human visceral and subcutaneous adipose tissue using microarrays provides strong evidence for genetic and developmental heterogeneity between depots. Genes involved in immunity and adipose tissue development are over-represented in the list of genes differentially expressed between visceral and subcutaneous adipose tissue [36, 111, 112]; 25% of 28 genes whose expression levels were higher in omental compared to subcutaneous were genes involved in innate immunity [36]. Complement components C2, C3, C4, C7, and factor B expression are higher in omental compared to subcutaneous adipose tissue, whereas expression levels of factor D (adipsin), C1QB, C1R, and C1S are similar between these depots. Vohl et al. [111] found that out of around 6000 genes expressed in adipose tissue, 409 transcripts corresponding to 347 encoded genes or expressed sequence tags were differentially expressed between omental and subcutaneous adipose tissue; 216 genes are expressed at higher levels in visceral, whereas 131 were expressed more abundantly in subcutaneous adipose tissue. Genes involved in cell differentiation and adipogenesis/organogenesis, such as CCAAT/enhancer-binding protein (C/ EBP)-d, Frizzled 7 (Wnt signaling), and NR2F1, are more abundant in omental adipose tissue, while homeo box gene, HoxA10 and HoxC6, C/EBP-a, and adipsin levels were higher in subcutaneous adipose tissue. Gesta et al. [112] conducted an analysis of adipose tissue gene expression in men and women of widely varying BMIs. They found clear depot differences in expression of developmental genes involved in anteroposterior or dorsoventral patterning in adipocytes from intra-abdominal and subcutaneous depots of mice and humans. Similar differences were found in a stromal fraction of the tissue that includes preadipocytes. The differences in gene expression pattern persist even after in vitro differentiation, suggesting the differences are independent of extrinsic factors. Kirklands group also demonstrated that gene expression proﬁles of preadipocytes/differentiated adipocytes are also different between depots [113]. Taken together, these ﬁndings suggest that different adipocyte precursors are responsible for a speciﬁc adipose depot development, and may participate later in the functional differences observed between intra-abdominal and subcutaneous adipose depots. Epididymal adipose tissue of rodents shows a human visceral-type pattern of gene expression as revealed by analysis of gene expression [112]. Developmental

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genes, most strikingly Nr2f1, Thbd, HoxA5, and HoxC8, were more highly expressed in both mouse epididymal and human omental adipose tissue, and applied to both fat cell and stromal fractions. Both these depots show a lower capacity for growth through fat cell number (preadipocyte recruitment) compared to subcutaneous adipose tissue and they both may exhibit higher rates of adipocyte death/tissue remodeling as reﬂected in an increased number of macrophages forming crown-like structures around necrotic/apoptotic adipocytes under the stress of obesity or high-fat feeding [25, 114]. This result was somewhat unexpected because the epididymal depot, unlike typical human visceral depots, it is not portally draining or rich in lymph nodes or milky spots. Humans do not have a large epididymal fat pad as do rodents, but like the omental depot of humans, this depot lies over the intestine in vivo, where it may serve some sort of immune defense function, as does the omental depot.

15.9 Depot Differences in Cell Proliferation and Differentiation Capacity

Given the association of excess intra-abdominal fat and metabolic disorders, it is important to know how the deposition of intra-abdominal versus subcutaneous adipose tissue is regulated. Accumulation of adipose tissue depends in part on the balance of new adipocyte formation, hypertrophy of existing adipocytes, and cell death. To understand the potential depot differences in new preadipocyte recruitment and differentiation, the isolated stromal vascular cellular fraction taken from different adipose depots (subcutaneous, omental, or mesenteric origin) has been used for in vitro studies. Several studies showed that there is no depot difference in preadipocyte differentiation and similar proliferative rates between omental versus subcutaneous stromal vascular cells [23, 24], while others show higher proliferation/ differentiation capacity in subcutaneous [16, 115]. Varying culture conditions, differentiation protocols used, subject characteristics (age, BMI, fat distribution), and contamination of other cell types during culture could explain discrepancies. Differences in innervation, vascularity, and overall cellular composition of each depot might be also relevant in vivo. Convincing studies using clonally derived preadipocytes strongly suggest that compared to omental preadipocytes, subcutaneous preadipocytes show greater replication potential and differentiation capacity [16, 115]. To avoid contamination of other cells and to detect intrinsic or cell autonomous differences in preadipocytes from different depots, Tchkonia et al. compared clonally derived cells from subcutaneous, mesenteric, and omental adipose tissue, and showed regional differences are still evident [16]. Expression levels of the adipogenic transcription factor C/EBP-a are higher in subcutaneous than omental cells, and subcutaneous preadipocytes proliferate more rapidly and differentiate to a greater extent. Furthermore, these differences persist for 40 population doublings in the human telomerase reverse transcriptase-expressing strains derived from a single cell from each depots [116]. Consistent with this observation, omental preadipocytes have longer telomeres than

15.10 Conclusions and Future Directions

subcutaneous preadipocytes [116], suggesting subcutaneous preadipocytes have a longer replication history. Further, there is a negative correlation between age and proliferation of subcutaneous stromal cells, but not of omental cells [23]. The greater capacity of subcutaneous preadipocytes to proliferate may underlie the great growth capacity of these tissues in response to overnutrition or treatment with PPAR-c agonists during therapy for diabetes. Upon differentiation in vitro, genes that associated with adipocyte maturation are more highly expressed in cultures derived from subcutaneous than omental depots (e.g., GLUT4, leptin, FABP-4) [113]. This fuller differentiation in subcutaneous depots may have been mediated by higher expression of the key adipocyte transcription factors C/EBP-a and PPAR-c [113]. Expression proﬁles of mesenteric preadipocytes are much more like subcutaneous than omental preadipocytes [113], and mesenteric preadipocytes replicate and differentiate faster than omental preadipocytes [16, 116]. These apparently intrinsic differences in adipocytes from different depots may mediate the depot differences in metabolic and endocrine function that were described earlier. Adipocyte cell number also depends on rates of cell loss as well as a recruitment of preadipocytes and differentiation into adipocytes. It is shown that human omental preadipocytes are more susceptible to apoptotic stimuli (serum deprivation or TNF) than subcutaneous preadipocytes [116, 117], suggesting the intrinsic differences in apoptosis between preadipocytes from two depots.

15.10 Conclusions and Future Directions

The mechanisms linking regional adiposity to metabolic complications are rapidly being elucidated, but there remain many key gaps in our knowledge. Many studies have measured expression and secretion of numerous pro- and anti-inﬂammatory cytokines in obese adipose tissues. As different cytokines can work additively, synergistically, or antagonistically, it will be important to understand how the balance of these factors affects regional adipose and systemic metabolism through paracrine and endocrine interactions. A number of new studies point to evidence for as-yet unindentiﬁed depot- and sexspeciﬁc factors. A recent study showed that removing similar and relatively small quantities of gonadal fat improved glucose tolerance in lean male and female mice, while with diet-induced obesity, only removal of parametrial fat in females was effective [118]. Kahns group [119] also showed that transplantation of epididymal fat adversely affected glucose homeostasis, while inguinal (subcutaneous) fat improved it. Changes in the levels of the known key adipokines were not explanatory, so further work with these models will be important. Most studies of regional differences in adipose function only sample fat from subjects at their usual weight and this may limit our ability to detect differential sensitivity or responsiveness among fat depots. For example, we recently demonstrated that culture with the glucocorticoid analog dexamethasone plus insulin

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(compared to insulin alone) increased 11b-hydroxysteroid dehydrogenase activity in the omental but not subcutaneous depot of obese subjects [133]. It seems likely that sampling of adipose tissues of men and women in response to stresses (e.g., over- or underfeeding or hormone administration) is likely to reveal important depot-speciﬁc responses. Research on the regulation of regional adipose tissue growth is receiving more attention as the molecular details of adipogenesis in cell culture models are being unraveled. Differences in the mass of different fat deposits may be more closely related to adipocyte number than size, particularly in the obese. Thus, further understanding of the mechanisms regulating preadipocyte recruitment and differentiation will be important in understanding sex- and age-related differences in fat distribution. Also, with better phenotyping and genome-wide associations, analysis of the genetic determinants of fat distribution will likely yield important insights into mechanisms. In summary, the mechanisms underlying the regulation of regional adiposity and the disproportionate effects of enlargements in visceral fat depots on health of men and women remain enigmatic. Our knowledge of physiologically important variations in the secretome of the adipose depots is rapidly emerging. Further research on the local and systemic consequences of these mediators holds promise for understanding the mechanisms linking adiposity to health and disease.

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depot-speciﬁc susceptibility to apoptosis. Diabetes, 47, 1365–1368. Shi, H., Strader, A., Woods, S.C., and Seeley, R.J. (2007) The effect of fat removal on glucose tolerance is depot speciﬁc in male and female mice. Am. J. Physiol. Endocrinol. Metab., 293, E1012–E1020. Tran, T.T., Yamamoto, Y., Gesta, S., and Kahn, C.R. (2007) Beneﬁcial effects of subcutaneous fat transplantation on metabolism. Cell Metab. 7, 410–420. Russell, C.D., Petersen, R.N., Rao, S.P., Ricci, M.R., Prasad, A., Zhang, Y., Brolin, R.E., and Fried, S.K. (1998) Leptin expression in adipose tissue from obese humans: depot-speciﬁc regulation by insulin and dexamethasone. Am. J. Physiol. Endocrinol. Metab., 275, E507–E515. Dusserre, E., Moulin, P., and Vidal, H. (2000) Differences in mRNA expression of the proteins secreted by the adipocytes in human subcutaneous and visceral adipose tissues. Biochim. Biophys. Acta, 1500, 88–96. Fisher, F.M., McTernan, P.G., Valsamakis, G., Chetty, R., Harte, A.L., Anwar, A.J., Starcynski, J., Crocker, J., Barnett, A.H., McTernan, C.L., and Kumar, S. (2002) Differences in adiponectin protein expression: effect of fat depots and type 2 diabetic status. Horm. Metab. Res., 34, 650–654. Hube, F., Birgel, M., Lee, Y.M., and Hauner, H. (1999) Expression pattern of tumour necrosis factor receptors in subcutaneous and omental human adipose tissue: role of obesity and noninsulin-dependent diabetes mellitus. Eur. J. Clin. Invest., 29, 672–678. Alessi, M.C., Bastelica, D., Morange, P., Berthet, B., Leduc, I., Verdier, M., Geel, O., and Juhan-Vague, I. (2000) Plasminogen activator inhibitor 1, transforming growth factor-beta1, and BMI are closely associated in human adipose tissue during morbid obesity. Diabetes, 49, 1374–1380. Fried, S.K., Bunkin, D.A., and Greenberg, A.S. (1998) Omental and subcutaneous adipose tissues of obese subjects release interleukin-6: depot difference and

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cell progenitors through distinct expression proﬁles and developmental gene patterns. Am. J. Physiol. Endocrinol. Metab., 292, 298–307. Fain, J.N., Cheema, P.S., Bahouth, S.W., and Lloyd, H.M. (2003) Resistin release by human adipose tissue explants in primary culture. Biochem. Biophys. Res. Commun., 300, 674–678. Schafﬂer, A., Neumeier, M., Herfarth, H., Furst, A., Scholmerich, J., and Buchler, C. (2005) Genomic structure of human omentin, a new adipocytokine expressed in omental adipose tissue. Biochim. Biophys. Acta, 1732, 96–102. Gabrielsson, B.G., Johansson, J.M., Jennische, E., Jernas, M., Itoh, Y., Peltonen, M., Olbers, T., Lonn, L., Lonroth, H., Sjostrom, L., Carlsson, B., Carlsson, L.M., and Lonn, M. (2002) Depot-speciﬁc expression of ﬁbroblast growth factors in human adipose tissue. Obes. Res., 10, 608–616. Lee, M.J., Fried, S.K., Mundt, S.S, Wang, Y., Sullivan, S., Stefanni, A., Daugherty, B.L, and Hermanovski-Vosatka, A. (2008) Depot-speciﬁc regulation of the conversion of cortisone to cortisol in human adipose tissue. Obesity, 16, 1178–1185.

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16 Viral Induction of Obesity and Adipogenesis Magdalena Pasarica, Rohan N. Dhurandhar, Nazar Mashtalir, and Nikhil V. Dhurandhar 16.1 Introduction

The World Health Organization has declared that the world is experiencing an obesity epidemic. As this epidemic continues unabated, infectobesity (obesity of infectious origin) has been receiving increasing attention in recent years [1–3]. In the last two decades, 10 obesity-promoting microbes have been reported [4–19]. During this period, the prevalence of obesity has increased by 30% in the United States. Several developed and developing countries echo this trend [20, 21]. Even in the best hands, obesity treatment yields merely marginal and transient weight loss. Considering our dismal record in ﬁghting this epidemic, new information about every aspect of obesity, from its etiology to its treatment, is urgently needed to devise new and more effective strategies. An accurate understanding of the varied etiological factors of obesity may point the way to treatments directed speciﬁcally towards the cause and, consequently, its successful management. This chapter presents information about a relatively novel and unusual cause of obesity – obesity of infectious origin. A multitude of factors contribute to the etiology of obesity [22] and a proportion of obesity may be caused by infections. If infection by certain pathogens promotes human obesity, recognizing and acknowledging their role is the ﬁrst important step in dealing with the infection and its pathogenesis. The adipogenic pathogens reported to date include human and nonhuman viruses, scrapie agents, bacteria, and gut microﬂora and parasites. This chapter describes various adipogenic pathogens reported since the ﬁrst of such reports in 1982 [23], their natural hosts, signs and symptoms, and pathogenesis (Table 16.1). Data for this chapter were identiﬁed by searches of MEDLINE and references from relevant articles. Numerous articles were ﬁrst identiﬁed by using search terms infectobesity, obesity and infection, viruses and obesity, fat and virus, and infection and weight gain. Search was further reﬁned by using search terms pertinent to the adipogenic pathogens reported in the literature. English and French language publications were searched. Publications including original articles, reviews, and editorials describing causation, association, and/or mechanism of

Adipogenic pathogens.

Pathogen

Animal model, age, and sex

Effect of infection on body weight

Effect of infection on biochemical parameters

Potential mechanism of action

Association with human obesity

CDV [23, 29, 32, 34, 37–39, 127–131]

Swiss albino mice, male and female, 4–5 weeks old

obesity developed 6–10 weeks postinfection and reaches a plateau after 16–20 weeks mean weight of infected obese animals was nearly twice compared to the uninfected group

" insulin

alters hypothalamic integrity

association with human multiple sclerosis

no signiﬁcant change in glucose

# leptin long receptor and " leptin

association with obesity not yet reported

" leptin levels # MCH, neuropeptide Y # catecholamine # neuropeptides

# MCH # neuropeptides

RAV-7 [7, 8, 44, 46, 132]

10-day-old embryos of white Leghorn chickens

stunting – 2 weeks posthatching

anemia

" visceral fat – 3 weeks of age

hyperlipidemia immune suppression

# catecholamine " cytokine production changes in lipid metabolism # thyroxine

not reported

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Table 16.1

BDV [16, 50, 54, 59, 133]

SMAM-1 [10, 14, 75]

Ad-36 [4, 12, 13, 15, 79–81]

4-week-old Lewis rats

3-week-old white Leghorn broilers

chickens, mice, male rhesus monkeys and marmosets, rats

" body weight – 14 days postinoculation

stunting

" visceral fat 3 weeks postinoculation " body weight

" amylase " insulin # glucose # thyroxine " triglycerides

moderate " blood glucose # cholesterol

possible hypothalamic damage due to inﬂammatory lesions and viral replication

transmission of BDV from animals to humans is possible; increase BDV seroprevalence in psychiatric and neurological disorders; association with obesity not yet reported

impaired liver function

yes; seropositive obese subjects heavier versus seronegative obese counterparts

# triglycerides

impaired lipogenesis glucagon deﬁciency

# cholesterol

" replication

(continued)

16.1 Introduction

yes; greater prevalence of seropositivity in obese versus nonobese subjects; seropositive subjects were heavier than seronegative counterparts

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Ad-37 [4, 89]

Animal model, age, and sex

3-week-old white Leghorn chickens

3-week-old female CD1 mice

Effect of infection on body weight

Effect of infection on biochemical parameters

Potential mechanism of action

" visceral, epididymal-inguinal, retroperitoneal fat onset 4 weeks in chickens, 6–7 months postinoculation in rodents and mammals

# triglycerides

" differentiation

" insulin sensitivity

" lipid accumulation in preadipocytes

# leptin expression and secretion # corticosterone and norepinephrine # triglycerides

" insulin sensitivity

no difference in body weight " visceral fat onset 4 weeks postinoculation " body weight

179% " whole-body adiposity

—

# leptin secretion and expression in fat cells " adipocyte differentiation " triglyceride accumulation

increased preadipocyte differentiation – hypothesized anti-inﬂammatory response determined by infection – hypothesized

Association with human obesity

not reported

not reported

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(Continued)

Pathogen

Ad-5 [18]

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Table 16.1

C. pneumoniae [9, 92, 134–136]

humans

Microbiota [5, 19, 100, 137]

8- to 10-week-old germ-free male B6/ NMRI mice

onset 22–23 weeks postinoculation signiﬁcantly higher BMI

association with coronary heart disease

no difference in body weight

insulin resistance, " glucose, " insulin

57% " whole-body fat

" metabolic rate with no alteration in tissue high-energy phosphate stores 2.3-fold " hepatic triglyceride content " expression of de novo fatty acid synthesis pathway (acetyl-CoA carboxylase-1 and fatty acid synthase)) # expression FIAF

69% " epididymal fat 7% # lean mass

onset after 14 days of colonization

mechanism unknown; preponderance of seropositivity in obese subjects is not a result of obesity " processing of polysaccharides

" inefﬁcient metabolism

association with increased BMI

obesity correlates with abundance of Firmicutes in the gut ﬂora weight loss decreases Firmicutes

" hepatic triglyceride production " FIAF, " incorporation of triglycerides into adipocytes by LPL

(continued)

16.1 Introduction

" leptin, " LPL activity in epididymal fat and heart

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(Continued)

Pathogen

Animal model, age, and sex

Effect of infection on body weight

Effect of infection on biochemical parameters

Potential mechanism of action

Association with human obesity

Gut parasites [101, 102]

adult male dragonﬂies (L. pulchella)

26% more thoracic lipid

no change in hemolyph lipid content

mechanism unknown; decline in ﬂight-muscle power related to changes in parasite size, maturity, and length of infection

not reported

brain function alteration

not reported

Scrapie agents [17, 107, 109, 110, 117, 138]

6- to 9-week-old female mice SJL, C57NL, A2G, SAMP8, SAMR1, AKR, 22L

" body weight due to fat accumulation

12 weeks postinfection

Reproduced with permission from [139].

" hemolyph carbohydrate levels " insulin resistance " chronic p38 mitogen-activated protein kinase activation # ﬂight-muscle power output " blood glucose

# GLUT1 density in certain brain areas

# GLUT1 density, # glucose tolerance " adrenal weight disturbs hypothalamic–pituitary–adrenal axis

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Table 16.1

16.2 Viruses

microorganisms with obesity, and/or weight gain in animals and humans were reviewed.

16.2 Viruses 16.2.1 Canine Distemper Virus

Canine distemper virus (CDV) was the ﬁrst obesity-promoting pathogen reported [23]. It is a lymphotropic and neurotropic negative-stranded RNA virus from the genus morbillivirus and paramyxovirus subgroup [24–26]. CDV causes a frequently fatal neurodegenerative disease in dogs and a wide range of mammals [23, 27–30]. The virus is antigenically related to the measles virus [31] and although CDV may be associated with other human diseases [29], it is not considered a human pathogen. CDV induced obesity in 26% of Swiss Albino mice after intracerebral viral inoculation and in 16% of animals after an intraperitoneal inoculation [23]. These results have since been reconﬁrmed [32–34]. CDV increased fat cell number and size, and body and fat pad weights in mice surviving the infection [23]. CDV inoculation by the intranasal, footpad, or subcutaneous route did not produce obesity, and vaccination with CDV surface antigens partially protected against acute encephalitis and obesity [35, 36]. In another study, the CDV-infected obese group showed 6 times greater insulin levels without differences in glucose levels compared to control animals [34]. Reduced catecholamine levels [23], decrease in hypothalamic expression of longform leptin receptor and the consequent leptin resistance [37], and downregulation of melanin-concentration hormone (MCH) precursor mRNA (ppMCH) are considered to contribute to CDV-induced obesity. Leptin, an adipokine, modulates food intake and energy metabolism, and MCH is a potent anorectic peptide involved in the hypothalamic regulation of feeding behavior. These ﬁndings suggest a role for food intake disregulation in CDV-infected animals. The food intake of CDV-infected animals was greater, but did not attain statistical signiﬁcance [23]. The additional energy intake, however small, over several weeks may eventually contribute to biologically signiﬁcant weight gain. Recently, Verlaeten et al. [38] suggested that CDV alters hypothalamic integrity and subsequently modiﬁes the homeostasis of the brain – a condition known to lead to obesity. Although viral products are expressed at high levels in the acute stages of infection, surprisingly, by the time the obesity develops, CDV is no longer detectable in the hypothalamus [32, 39]. Therefore, Bernard et al. [37] speculated that the initial impact of CDV on the hypothalamus initiates changes that continue to promote obesity months after the acute infection, suggesting a hit-and-run effect similar to that described by Oldstone et al. [40]. No association of CDV with human obesity has been reported.

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16.2.2 Rous-Associated Virus-7

Rous-associated virus (RAV)-7 was reported to cause obesity as well as stunting and hyperlipidemia in chickens [8]. It is an avian leucosis retrovirus and the most common avian retrovirus associated with neoplastic disease in domesticated poultry [41]. RAV-7 belongs to subgroup C of the ﬁve subgroups of the virus based on envelope glycoproteins, A–F [42]. It shows the least antigenic cross-reactivity among the other subgroup C viruses [43] and shows similar pathogenesis, but also produce obesity [44]. RAV-7-induced obesity is characterized by stunting, fat deposition around crop and abdominal fat pads, and hyperlipidemia [8]. While there was no evidence of neoplastic transformation, stunted growth was the most impressive effect of RAV-7. Obesity was observed in birds infected as 10-day-old embryos, but not when inoculated as 1-day-old hatched chickens [8]. Difference in food intake was observed as early as 3 weeks postinfection, when stunting and obesity developed [7]. Fifty days after hatching, the RAV-7-infected chickens weighed 194 g compared to 515 g in the control group. The infected chickens developed fatty, yellow-colored livers, hepatomegaly, anemia, immune suppression, and striking hyperlipidemia. Serum triglyceride levels in the infected group were 1500 mg/dl compared with a range of 68.2–90.5 mg/dl in uninfected chickens. Pancreatic damage was indicated by histological changes, increased serum amylase levels, and reduced serum glucose and glucagon levels. Glucagon levels dipped 20 and 30 days posthatching, and as expected in poultry [45], the drop was accompanied with higher mortality. Insulin levels, on the other hand, remained at, or even above, normal. RAV-7-induced thyroiditis, and marked reductions in T3 and T4 levels [46], which may explain the adiposity and hyperlipidemia [46, 47]. The virally induced syndrome was ameliorated by thyroid hormone replacement [46], demonstrating the role of hypothyroidism in development of RAV-7-induced adiposity. 16.2.3 Borna Disease Virus

Borna disease virus (BDV) is an enveloped, nonsegmented, negative-stranded RNA virus belonging to the Mononegavirales order [48]. Although it can produce a strong immune response and can be fatal, it can persist in the nervous system with no major symptoms [49]. It causes encephalopathy in a broad range of animals, like horses, sheep, cattle, cats, and dogs [50]. Experimental infection with BDV has been reported in chicken, mice, rat, rabbit, hamster and rhesus monkey models [51, 52]. Infected rats show behavior changes and learning deﬁcits [50]. In 1991, Gosztonyi et al. [53] described an obesity syndrome induced by BDV in rats. The induction of obesity was dependent on the age and genetic characteristics of the animals and the virus strain [16, 53]. Weanling or adult rats developed acute encephalitis 1–4 months postinfection. Two months later, surviving animals developed marked obesity. Obese rats had massive visceral fat deposition, with increased

16.2 Viruses

triglycerides and hyperplasia of islets of Langerhans, and a moderate increase in blood glucose [16]. It is believed that BDV-induced obesity may be due to inﬂammatory lesions and viral antigen expression in brain, especially in the hypothalamus, which is known to regulate body weight and food intake [54]. BDV received considerable attention when de la Torre demonstrated that it can infect the human brain, by detecting a BDV-speciﬁc antigen in four autopsied human brains [55]. Ludwig et al. [50] have since postulated that a cross-species transmission of BDV from animals to humans could occur. Although its pathogenesis is not yet clear, BDV as a pathogen once thought to only infect animals, indeed infects humans as well. BDV incidence in humans is reported worldwide: in the United Kingdom [56], Japan [57], China [58], Germany [59], and in the United States [60]. Seroprevalence in humans varies from 0 and 48% and viral prevalence from 0 to 82% [61]. Common human mental disorders like depression and schizophrenia are associated with the prevalence of BDV-speciﬁc antibodies [57–59, 62–64] and the presence of BDV RNA in the peripheral blood [65]. Chen et al. [62] provided some evidence for a possible human-to-human transmission of BDV infection by showing that the family members of psychiatric patients and mental health workers had a higher prevalence of BDVantibodies than the control population. However, Ferszt et al. [63] postulated that the higher prevalence of BDV antibodies in depressive patients may be a nonspeciﬁc aspect of immunosuppression. An association between obesity and mental depression is well documented [66–68], and some recent reports show that depression can predict weight gain [66, 69]. Although an association between BDV and human obesity has not been established, the role of BDV in depressed obese humans appears worth investigating. 16.2.4 Adenoviruses

Adenoviruses are nonenveloped DNA viruses with icosahedral symmetry and a diameter of 65–80 nm [70]. They were ﬁrst isolated in 1953 by investigators trying to establish cell lines from the adenoidal tissue of children after tonsillectomies [71]. In nature, adenoviruses are widespread; infecting birds, mammals and humans. In humans, adenoviruses are frequently associated with acute upper respiratory tract infections, enteritis, or conjunctivitis. The presence of serum antibodies to various adenoviruses is common in the general population [72]. Adenoviral DNA has been detected in adult human lymphocytes and the number of these lymphocytes increases in correlation with the age of the person [73]. There are six major subgroups (A–F) among the 50 human adenoviruses maintained by the American Type Culture Collection. Each subgroup has a number of speciﬁc serotypes. The following is a brief description of four adenoviruses reported to cause obesity in animal models. 16.2.4.1 SMAM-1 The avian adenovirus SMAM-1 was discovered during a poultry epidemic in the early 1980s [74]. It is serologically related to the CELO (chick embryo lethal orphan) avian

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adenovirus. SMAM-1 causes adiposity in chickens. The peculiar features of its syndrome include increased adiposity and relative hypolipidemia. Chickens intranasally inoculated with SMAM-1 and their uninfected cagemates showed significantly greater adiposity, particularly in visceral fat depots (compared to separately housed control animals) [10, 14]. The development of adiposity in the uninfected cagemates of infected chickens adiposity suggested a horizontal transmission of the virus. Interestingly, the virally induced adiposity was accompanied by paradoxically lower serum cholesterol and triglycerides levels. The food intake of the chickens could not explain the adiposity [10, 14]. Subsequently, in 1997, Dhurandhar et al. [75] reported an association between SMAM-1 seropositivity and human obesity. Thus, it was the ﬁrst virus reported to be associated with human obesity. Twenty percent of obese subjects screened had antibodies to SMAM-1. Seropositive subjects had a signiﬁcantly greater body weight (95.1 2.1 versus 80.1 0.6 kg; P < 0.02), but 15% lower serum cholesterol and 60% lower serum triglycerides [75]. Avian adenoviruses, being serologically different from their human counterparts, have been thought to be incapable of infecting across species [76, 77]. Humans showing antibodies to an avian adenovirus challenges that view. It is also possible that the subjects of this study had antibodies to a human adenovirus that had cross-reacted with SMAM-1. Prevalence of SMAM-1 seropositivity in general population has not been reported. 16.2.4.2 Ad-36 Ad-36 was ﬁrst isolated in 1978 in Germany from the feces of a 6-year-old girl with diabetes and enteritis [78]. Ad-36 infection was thought to be rare, but recent work indicates that 11–30% of the individuals screened in the United States are seropositive for Ad-36 antibodies [4]. In separate experiments, chicken, mice, rats, and marmosets (nonhuman primates) inoculated with human adenovirus Ad-36 developed a syndrome of increased adipose tissue, and paradoxically low levels of serum cholesterol and triglycerides [12, 13, 15, 79], without a signiﬁcant change in food intake. The adipogenic effect was most pronounced in visceral fat depots. The amount of Ad-36 DNA present in visceral fat correlated with the amount of total visceral fat in the infected animals [12] (r ¼ 0.41, P < 0.03). When obesity was deﬁned as body fat greater than or equal to the 85th percentile of that in the control group, 64–72% of the Ad-36-infected animals were considered obese versus only 18% in the uninfected controls Sections of the brain and hypothalamus of inoculated animals did not show any overt histopathological changes [13]. Subsequently, the avian adenovirus CELO, and two other human adenoviruses, Ad-2 and Ad-31, were shown to be nonadipogenic in animal models [4, 13]. Thus, the potential to promote obesity is not common to all adenoviruses. Post hoc screening of stored plasma samples from adult male rhesus monkeys housed at the Regional Primate Research Center in Wisconsin showed natural infection of animals with Ad-36 as indicated by development of neutralizing antibodies [15]. This spontaneous appearance of Ad-36 antibodies was followed by signiﬁcant gain in weight and reduction in serum cholesterol in the year following

16.2 Viruses

the seroconversion compared to the year preceding it. This association with seroconversion is suggestive of the adipogenic consequence of natural Ad-36 infection. Interestingly, the human adenovirus Ad-36 appears to infect across a wide range of nonhuman hosts including chicken, mice, rats, marmosets, and rhesus monkeys [12, 13, 15, 79]. Although it showed causation or association with adiposity in these models, its effect on body weight of the host varied with the species (Table 16.1). Ad36-infected rats and mice showed 7% greater body weight compared to the uninfected controls, whereas infected chickens did not show greater body weight. The difference in body weights was greater for marmosets and rhesus monkeys (12 and 15%, respectively), but was the highest for humans (20%). The adipogenic mechanism of Ad-36 is unclear, but several in vitro and in vivo studies provide some insight about its molecular effects on adipose tissue metabolism. Recent studies show that Ad-36 has effects similar to those of thiazolidinediones (TZDs) – a class of antidiabetic drugs. Like TZDs, Ad-36 increases fat cell replication [80], differentiation [81], lipid accumulation [81], and insulin sensitivity [81, 82], and reduces leptin secretion and expression [82]. These effects were not observed with Ad-2 – a nonadipogenic human adenovirus. When infected, 3T3-L1 preadipocytes (mouse embryo-derived preadipocytes) and human primary preadipocytes show increased differentiation to adipocytes [81]. Ad-36 enhances lipid accumulation induced by differentiation inducers (Figure 16.1) and also induces it spontaneously even in the absence of inducers (Figure 16.2). Figure 16.2 shows that only Ad-36-infected human primary adipose tissue-derived stem cells (hASCs), as evident from the presence of viral proteins and bulging nucleus, show adipose differentiation-related protein (ADRP) – an early marker of adipose differentiation program and lipid presence [83]. The cells were not exposed to adipocyte inducers. Hence, as expected, the uninfected cells remained undifferentiated. After viral

Figure 16.1 Lipid accumulation in 3T3-L1 preadipocytes infected with Ad-36 in the presence of a cocktail of methylisobutylxanthine, dexamethasone, and insulin (MDI). (a) Mock infected. (b) Ad-36 infected. Confluent 3T3-L1 preadipocytes were infected with Ad-36, or mock

infected, and differentiation was induced by exposure to MDI. Cells were stained with lipidspecific fluorescent dye BODIPY (493/503). The figure shows a much greater number of lipidbearing cells in the Ad-36-infected group. (Reprinted with permission from [81].)

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Figure 16.2 Cells from the stromal vascular fraction of human adipose tissue were infected with Ad-36 with a multiplicity of infection of 3.8. Cells were fixed on day 9 postinfection in 4% paraformaldehyde and processed for immunofluorescence. Slides were mounted in ProLong Gold antifade reagent with DAPI (blue) (Invitrogen). Lipid droplets were identified using mouse monoclonal anti-ADRP antibodies (Research Diagnostics Inc.). AD-36 viral

particles were identified using anti-Ad-36 rabbit polyclonal antibodies. Secondary antibodies were anti-mouse Alexafluor 488 (green) (Invitrogen) and anti-rabbit Alexafluor 594 (red) (Invitrogen), respectively. (a) Cell nuclei stained with DAPI. (b) ADRP indicating lipid accumulation. (c) Viral proteins in and around a cell nucleus. (d) Overlay slide shows ADRP only in the cell that is infected with Ad-36, but not in the surrounding cells without virus.

inoculation, not all cells in a plate get infected with Ad-36 immediately. The experiment described in Figure 16.2 provided an excellent opportunity to compare onset of lipogenesis in uninfected and infected cells in the same plate. These data show that within the infected group of hASCs, lipogenesis is initiated as a direct consequence of Ad-36 infection. Although multiple pathways may explain the adipogenic effect of Ad-36, its prodifferentiation effect on preadipocytes appears to be a promising explanation. Association with Human Obesity Recently, prevalence of Ad-36 antibodies in 360 obese (body mass index (BMI) 30 kg/m2) and 142 non-obese (BMI < 30 kg/m2) subjects from Wisconsin, Florida, and New York was reported. Antibodies were more prevalent in obese subjects (30%) than in nonobese subjects (11%). The effect of Ad36 seropositivity on the risk of obesity was highly signiﬁcant, and independent of age, sex, and the data collection site [4]. Interestingly, seropositive subjects, both obese and nonobese, had signiﬁcantly greater BMI compared to their respective seronegative counterparts. Similarly, between human twins – who are generally believed to have

16.2 Viruses

similar body weight and fatness – Ad-36-seropositive twins had signiﬁcantly greater body fat and BMI compared to their antibody-negative cotwins [4]. Globally speaking, the prevalence of Ad-36 antibodies in the human population appears to vary geographically. Only 5% of subjects screened in Denmark had antibodies to Ad-36 [84]. Ad-36 is a human virus that causes adiposity in several animal models and shows association with human obesity. This makes Ad-36 perhaps a leading candidate pathogen for determining its causative role in human obesity. Ethically, it is impossible to infect humans with the virus, which precludes an unequivocal demonstration of its causative role in human obesity. Therefore, such evidence must come from indirect approaches, such as determining the viral mechanism of action using tissue culture and animal models, and then applying that information to humans. 16.2.4.3 Ad-5 Ad-5 is a human adenovirus that causes respiratory tract infections in humans [85]. Recently, So et al. [18] showed that Ad-5 infection induces adiposity in mice, independent of their food intake. The infected group had a 2.8-fold increase in percentage of body fat measured by in vivo magnetic resonance spectroscopy. About 67% of the infected animals developed obesity against only 17% in the control group. This incidence of obesity was very similar to the 64–72% rate of obesity development observed for Ad-36 [12]. Although the authors suggested that increased preadipocyte differentiation (as in the case of Ad-36) may be responsible for the increase observed in body fat, no data to support this speciﬁc mechanism are available. Ad-5 is extensively used for gene therapy [86], because as a vector it is safe, efﬁcient, and can accommodate large antigens encoding structure [87]. Although replicationdeﬁcient virus is used as a vector, adipogenic potential of a replication deﬁcient Ad-5 have not been reported. Such veriﬁcation would greatly beneﬁt researchers using Ad5 vectors to avoid potentially confounding adipogenic effects of the virus. 16.2.4.4 Ad-37 The human Ad-37 was discovered by de Jong in 1981 [88]. It causes keratoconjunctivitis and genitourinary tract infections in humans [88]. Whigham et al. [89] showed that Ad-37 causes adiposity in chickens, by an unknown mechanism. The visceral fat pads of infected chickens were 3 times heavier than those of the uninfected control group. Unlike Ad-36, it did not reduce serum cholesterol levels. Food intake could not explain the adiposity. 16.2.4.5 Adipogenic Potential of other Adenoviruses The adipogenic potential is not common to all adenoviruses. Thus far, reports show that human adenoviruses Ad-5, Ad-36, and Ad-37 are adipogenic, and Ad-2 and Ad-31 and avian adenovirus CELO are nonadipogenic [4, 13, 18, 89]. Unlike Ad-36, Ad-2 and Ad-31 did not show association with human obesity; the antibody prevalence between obese and nonobese subjects was similar (76 versus 81% for Ad-2 and 70 versus 80% for Ad-31). In contrast to the ﬁndings with Ad-36, BMIs and serum lipids were

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virtually identical for Ad-2 or Ad-31 regardless of the respective antibody status [4]. These ﬁndings are important – the almost equal distribution of seropositivity of Ad-2 and 31 between obese and nonobese subjects alike suggests that the greater prevalence of Ad-36 antibodies in obese subjects is most likely a cause, not result, of obesity. The adipogenic potential of the other serotypes is unknown. More information about the mode of action and elucidating viral mechanisms of adipogenesis may help in screening other serotypes for their adipogenic effects. Such a screening is important for the adenovirus class of viruses due to their high presence in the population and the use as vectors in research. 16.3 Chlamydia pneumoniae

Chlamydia are eubacteria that cause infection at some time in life for almost all humans [90]. It results in 10% of community acquired pneumonia, and 5% of bronchitis and sinusitis cases [91]. Chlamydia (Chlamydophila) pneumoniae is also the ﬁrst bacteria reported to be associated with increased BMI in humans. Some studies have reported an association between C. pneumoniae infection and increased BMI [9, 92], while others have not found any relation [93–95]. Subjects positive for immunoglobulin G to C. pneumoniae had a signiﬁcantly higher mean BMI compared to the seronegative subjects [9]. Ekesbo [92] reported that a group of subjects with a combined serology for C. pneumoniae and Helicobacter pylori had signiﬁcantly higher BMI compared to the control group (27.3 versus 25.8 kg/m2). Greater age, lower socioeconomic status, and greater fasting levels of insulin also characterized the antibody-positive subjects. The authors of the study suggested that obesity might be a marker not only for lower social class but also for greater than normal susceptibility to such infections [92]. Obesity is associated with impaired immunity [96] and, as suggested by Ekesbo et al. [92], might be the indicator for increased susceptibility to H. pylori and C. pneumoniae infection [92]. However, Dart et al. [9] found no preponderance of antibodies to other types of Chlamydia (Chlamydia trachomatis and Chlamydia pssittaci) in the same subject population that showed an association of BMI with C. pneumoniae antibodies. This indicated that an increased BMI did not necessarily predispose subjects to catch a C. pneumoniae infection. Considering its widespread prevalence, to further determine the role of C. pneumoniae in obesity, experiments with animal models designed to investigate the adipogenic potential of C. pneumoniae as well as human sero-epidemeological data are needed. 16.4 Gut Microbiota

Gut microbiota is the collection of microorganisms that grow in the intestines. It is mostly composed of anaerobic bacteria, which are essential for processing dietary

16.5 Gut Parasites

polysaccharides. These bacteria act in symbiosis with the gut – consuming and distributing energy by processing nutrients otherwise inaccessible to humans [6]. Moreover, they fortify the mucosal barrier and stimulate angiogenesis [97–99]. Introduction of normal gut microbiota into the guts of 8- to 10-week-old germ-free mice caused insulin resistance, a 57% increase in total body fat with a 61% increase in epididymal fat, and a 7% decrease in lean mass [5] of the germ-free recipients. Serum lipid proﬁles did not change after microbial colonization. The age at intervention or the duration of colonization of the microbiota did not increase the effect on adipose tissue. Microbiota colonization resulted in a 27% decrease in metabolic rate of the animals and upregulation of the de novo fatty acid synthesis pathway, which may explain the observed weight gain without increased food intake. Microbial colonization of the gut suppressed fasting-induced adipocyte factor (FIAF), which is an inhibitor of lipoprotein lipase. Thus, suppression of FIAF was required to promote triglyceride deposition in adipocytes. Effects, if any, of gut microbiota on human body weight gain are unknown. However, the ﬁndings of Backhed et al. [5] demonstrate the ability of some nonpathogenic microbes to alter body composition and metabolism in a proadipogenic manner. In 2006, Turnbaugh et al. [19] demonstrated that the microbiota of the distal gut of genetically obese (ob/ob) mice have an increased ability to harvest energy from the diet. When the microbiota of these genetically obese mice were transmitted to colonize the gut of germ-free mice, the newly infected mice had a signiﬁcantly increased level of total body fat than their germ-free counterparts colonized with microbiota from genetically lean mice. Moreover, the obese microbiota had a relative abundance of the Firmicutes versus the Bacteroidetes division of bacteria and the reverse was true with microbiota of the lean mice. In a companion publication, the same group [100] also reported that obese human subjects had a lesser proportion of Bacteroidetes than lean subjects. Weight loss on a low calorie diet increased the proportion of Bacteroidetes – a lean-type microbiota. This, according to the authors, suggests possibilities for microbe-based therapeutic intervention for obesity. Although these studies collectively suggest a novel role for gut microbiota in regulation of adipose tissue mass, the accompanying Editorial cautions about statistically signiﬁcant, but biologically small gains in body fat of the mice receiving obese microﬂora [19]. Furthermore, it is inexplicable why the gut ﬂora of obese subjects after weight loss change to those of lean subjects. It appears counterintuitive to see the change of the gut ﬂora upon weight loss to less-efﬁcient harvester of energy. Further research is required to answer these questions.

16.5 Gut Parasites

Dragonﬂies (Libellula pulchella) infected with a common, noninvasive gregarine gut parasite (Apicomplexa: Eugregarinorida) showed symptoms similar to the human metabolic syndrome [19]. Infected dragonﬂies developed signiﬁcantly higher thoracic lipid accumulation, an inability to oxidize fatty acids in muscle tissue, 2-fold

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higher hemolyph carbohydrate concentrations than uninfected insects, increased insulin resistance, and elevated markers of a chronic inﬂammatory state. To isolate the effects of the gregarine parasite itself, healthy dragonﬂies were fed water with small amounts of excretory-secretory products (ESP s) from live gregarines. Two days of exposure to gregarine ESP s caused an elevation in hemolyph carbohydrate concentrations compared with control dragonﬂies (receiving regular water), comparable to the difference between healthy and gregarine infected dragonﬂies. This indicated that factors secreted by the parasites in the midgut in infected insects were responsible for the changes in hemolyph glucose concentrations [101]. The sister taxon of gregarine parasites, Cryptosporidium, shows similar symptoms in AIDS-infected humans. HIV-seropositive males with AIDS-deﬁning diagnoses of Cryptosporidium infections had reduced resting energy expenditure and fat oxidation, and increased carbohydrate oxidation and nonprotein respiratory quotient in comparison with subjects with other AIDS-deﬁning diagnoses (microsporidiosis, Pneumocystis carinii pneumonia, cytomegalovirus enteritis, and Mycobacterium avium-intracellulare) [102]. While the speciﬁc relation of cryptosporidiosis and obesity is not clear, it is of relevance that a parasite analogous to gregarine can result in similar metabolic changes in humans.

16.6 Scrapie Agents

Scrapie is a neurodegenerative disease with a long incubation period, known to occur in sheep and goats, and it can also infect mice and other small rodents [103]. Although the main manifestation of scrapie infections is abnormal behavior and motor dysfunction, certain scrapie strains induce obesity in experimentally infected mice. Weight increase in scrapie-infected animals was observed as a preclinical manifestation as early as 1968 by Pattison [104], and was followed by observations by Outram and Markovits in 1972 and 1981, respectively [105, 106]. The weight increase was due to fat accumulation and not edema [106, 107]. Regardless of the mouse strain, the scrapie strain ME7 injected in the hypothalamus induced obesity 12 weeks postinfection [17, 107–109]. This effect was also observed for the 22L [17, 107], but not for 139A [17, 107–109] or 22A scrapie strains [107]. Weight gain in ME7-infected mice paralleled increased food consumption [109]. The weight gain appeared 67 days postinfection and continued throughout the preclinical phase of the disease [109]. Unexpectedly, there was also an increase in food consumption in nonadipogenic scrapie strain 139A-infected mice, which did not become obese, suggesting possibly separate mechanisms for the increases in food consumption and weight [109]. Involvement of the brain is strongly implicated in scrapie-induced obesity. Kim et al. [17] speculated that scrapie-induced obesity is related to changes in central nervous system and neuroendocrine dysfunction. Role of brain function alterations was further supported by the ﬁndings of Vorbrodt et al. [110], who reported a decrease in glucose transporter 1 (GLUT1) density in certain brain regions, and reduced

16.7 Interaction of Pathogens and Adipose Tissue

glucose tolerance and hyperglycemia in ME7-infected mice. Glucose is of critical importance for normal functioning of the nervous system and impaired glucose transport may lead to impaired brain function. The authors recommend further studies to elucidate the regional difference in GLUT1 expression and the effect on brain function [110]. In addition to the role of higher centers, a role for a hypothalamic–pituitary–adrenal axis-mediated mechanism for scrapie-induced obesity has been suggested [109]. The adrenals in scrapie-induced obese mice were signiﬁcantly larger than controls [109] and adrenalectomy performed before ME7 injection prevented weight gain in mice, which supports the hypothesis. A role, if any, of scrapie agents in human obesity has not been reported.

16.7 Interaction of Pathogens and Adipose Tissue

Although a causative role of certain infections in obesity is a relatively novel concept, the involvement of adipose tissue with modulators and mediators of immune response is well documented. For instance, Cousin et al. [111] showed that preadipocytes function like macrophages, and possess phagocytic and microbicidal activity. Leptin, an adipocyte-secreted hormone involved in body weight regulation, also enhances the proliferation and activation of human circulating T lymphocytes and stimulates cytokine production [112]. In addition, adipocytes themselves secrete various cytokines [113, 114], and, in turn, preadipocytes and adipocytes are subject to cytokine-directed modulations [115, 116]. Combs et al. [117] showed that adipocytes provide a chronic reservoir for Trypanosoma cruzi parasites, which attracts inﬂux of macrophages to adipose tissue and also modulates adipokine secretion. With such an extensive interaction between the immune system and adipose tissue, the expansion of the latter in response to particular infections is certainly conceivable. For instance, macrophage colony-stimulating factor (M-CSF), which promotes the production of macrophages, is also secreted by adipocytes and, when overexpressed in vivo, induces signiﬁcant adipose tissue hyperplasia [118]. It is unknown if obesity promoting pathogens stimulate M-CSF production leading to the growth of adipose tissue. M-CSF is an example of a proinﬂammatory cytokine. The relationship of infections with inﬂammation is well known. A recent body of evidence shows an association of obesity with cytokines and markers of inﬂammation. Elevated levels of interleukin6 [119] and C-reactive proteins (CRPs) [120] are observed in obese individuals. Interestingly, Duncan et al. [121] showed that markers of inﬂammation can predict weight gain in middle-aged adults. It remains to be determined if inﬂammation is a cause or effect of obesity, and whether a pathogenic infection leads to inﬂammation and consequent increase in adiposity. Recently, Fernandez-Real et al. [122] showed that in healthy middle-aged men, burden of infection was positively correlated with fat mass. Total seropositivity for herpes simplex virus 1 and 2, C. pneumoniae, and enteroviruses contributed to 9% of fat mass variance, independent of age and CRP. The authors concluded that either the subjects with greater fat mass were more

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susceptible to contracting multiple infections, or the exposure to multiple infections increases fat mass [122]. In a closely related study, the same team of researchers also showed that in healthy middle-aged men, the burden of infection due to these same pathogens was independently and negatively related to insulin sensitivity [123]. The authors hypothesized that exposure to multiple infections may cause a chronic lowgrade inﬂammation and result in insulin resistance.

16.8 Adipogenic Pathogens and Humans

Although some of the aforementioned adipogenic pathogens are conventionally considered to be nonhuman pathogens, the possible use of a human host by these pathogens cannot be ruled out. Such was the case with BDV, which was originally considered to be a virus of horses and sheep, but now shows clear evidence of human infections [16, 50, 55, 61]. Similarly, humans show antibodies to SMAM-1, the adipogenic avian adenovirus [75]. Moreover, human adenovirus Ad-36 is not fastidious about a host species, and infects chickens, mice, nonhuman primates, rats, and hamsters [12–14, 79]. A change of host by pathogens has been well documented. For instance, CDV changed the host to infect feline species and considerably reduced the lion population in the Serengeti [124]. Therefore, nonhuman adipogenic pathogens cannot be guaranteed to be nonpathogenic for humans. Although several pathogens have been unequivocally shown to cause obesity in animal models, determining their contribution to human obesity is of great significance – and a challenge. Unlike animal models, humans cannot be experimentally infected with these pathogens due to ethical reasons. This limitation precludes any direct demonstration of a cause-and-effect relationship between a pathogen and adiposity, and the evidence of such must be only indirect and circumstantial. The insidious onset of obesity makes it difﬁcult to link to a particular episode of infection experienced in the past. Moreover, if an adipogenic pathogen uses a hit-and-run mechanism to induce obesity, it may be undetectable in the body by the time the adiposity is noticed. Weight of mounting indirect evidence will be required to convincingly establish such a relationship. Using animal and tissue culture models to elucidate the adipogenic mechanism of a pathogen may help considerably in understanding its role in human obesity.

16.9 Conclusions

The rapid increase in obesity, its comorbidities, and associated healthcare costs have prompted a search for better approaches for its prevention and management [125]. Such efforts may be facilitated by better understanding the etiology of obesity. Of its many etiological factors [22], the viral etiology of obesity was ﬁrst reported in 1982 [23], followed by several reports of adipogenic pathogens. Some of these

References

pathogens are associated with human obesity [4, 9, 126], but their causative role in human obesity has not been established. In addition to treatment, prevention could be a long-term goal of researchers investigating infectobesity. Vaccines against adipogenic pathogens could be expected to provide protection speciﬁcally against obesity due to speciﬁc pathogens. In summary, infectobesity (obesity of infectious origin) would be a novel, yet signiﬁcant concept, if it is shown to be relevant to humans. An adequate understanding of infectobesity is needed if we are to understand the implications of adipogenic pathogens and, thereof, treatment for the better management of obesity. A new perspective on the infectious etiology of obesity may stimulate additional research to assess the contribution of hitherto unknown pathogens in human obesity and possibly to prevent or treat obesity of infectious origins.

Acknowledgments

This work was partly supported by NIH grant 1R01 DK066164-01 awarded to N.V.D.

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17 Adipose Tissue Cachexia Michael John Tisdale 17.1 Introduction

Patients with cancer cachexia show a progressive loss of body weight due to selective depletion of skeletal muscle and stores of adipose tissue. Studies on the body composition of cachectic cancer patients show that even in severe wasting a considerable proportion of body fat is retained (the maximum fat loss observed was 50%) and loss of body weight was predominantly due to loss of skeletal muscle, while nonmuscle tissue, including the visceral fraction, did not change [1]. However, body fat is lost more rapidly than lean tissue in progressive cancer cachexia [2]. Plasma levels of unesteriﬁed fatty acids have been shown to be higher in patients with neoplastic disease [3], and cancer patients with weight loss have increased glycerol and fatty acid turnover compared with normal subjects or cancer patients without weight loss [4]. Fasting levels of plasma glycerol have also been shown to be higher in weight-losing cancer patients compared with weight-stable individuals, providing evidence for an increase in lipolysis [5]. Studies in nude mice show that depletion of carcass lipid is a function of tumor type and not tumor burden, suggesting that tumors produce substances that induce lipid depletion in cachexia [6]. This would explain why glucose administration in normal individuals suppresses lipid mobilization, while in patients with malignant diseases there is impaired suppression and continued oxidation of fatty acids [7].

17.2 Changes in Adipose Tissue in Cachexia

Most of these studies have been carried out in mice with cachexia, where there is extensive loss of white adipose tissue (WAT). Such tissue contains shrunken adipocytes, which are heterogeneous in size, and there is increased ﬁbrosis [8]. The small adipocytes show severe delipidation and modiﬁcations in cell membrane conformation. The mitochondria differ from typical WATmitochondria, are electron

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Table 17.1 Changes in mRNA and protein levels of adipogenic transcription factors and other proteins in cachexia.

Decreases

Increases

C/EBP-a and -b PPAR-c SREBP-1c acetyl-CoA carboxylase glycerol-3-phosphate acyl transferase GLUT4 leptin inhibitory GTP-binding protein (Gai) UCP-2 stimulatory GTP-binding protein (Gas)

dense, and have increased cristae. There are major reductions in both mRNA and protein levels of adipogenic transcription factors including CCAAT/enhancer-binding protein (C/EBP)-a and -b, peroxisome proliferator-activated receptor (PPAR)-c, and sterol regulatory element-binding protein (SREBP)-1c. Levels of mRNA for acetyl-CoA carboxylase, stearoyl-CoA desaturase-1, and glycerol-3-phosphatase acyl transferase, targets of SREBP-1c, also fell, as did glucose transporter 4 (GLUT4) and leptin (Table 17.1). Levels of uncoupling protein (UCP)-2 increased. These changes point to an impairment not only in the lipid storage capacity of adipocytes in cancer cachexia, but also the ability to differentiate. Adipocyte differentiation is controlled by a myriad of transcription factors that are activated in a sequential manner [9]. Initially, C/EBP-b and -d are transiently activated, and these then activate the expression of PPAR-c, which in turn stimulates C/EBP-a expression, which synergizes with PPARc in controlling terminal differentiation. Differentiation is enhanced by SREBP-1c, which activates transcription of PPAR-c [10]. C/EBP-a deﬁciency in mice leads to greatly reduced body fat, and adipocytes lacking C/EBP-a accumulate less lipid, cannot induce PPAR-c expression, and there is also a complete absence of insulinstimulated glucose transport [11]. Thus, the decreased expression of adipogenic transcription factors in cancer cachexia would disrupt recruitment and maintenance of the adipocyte phenotype. The decrease in GLUT4 mRNA would suggest an altered glucose metabolism and adipose tissue of cachectic mice has been shown to use glucose at only one-third of the rate of noncachectic animals [12]. Glucose consumption by the brain is also decreased in response to the increased glucose utilization by the tumor and the brain adapts to use ketone bodies derived from fat metabolism, as in starvation. There is evidence that adipocytes from cachetic mice as well as humans with cancer cachexia have an increased response to lipolytic stimuli [13]. Patients with weight loss show an increased sensitivity to adrenaline compared with weight-stable subjects, and also show increased plasma and urinary catecholamines, elevated heart rate, and increased oxidation of fatty acids [5]. The effect has been attributed to an enhanced stimulation of adenylyl cyclase, due to an increased expression of the stimulatory GTP-binding protein (Gas) and a decrease in the inhibitory form (Gai) [13]. Similar changes were noted in both mice and humans with cachexia. The changes in

17.3 Energy Expenditure in Cancer Patients

G-protein expression were induced in WAT from normal mice by a tumor-produced lipid-mobilizing factor (LMF) (see Section 17.7), suggesting that such factors not only directly stimulate lipolysis, but also sensitizes adipocytes to other lipolytic stimuli. There may be changes in lipid metabolism in cancer patients, even when there is no overt cachexia. Thus, in the sera of patients with gynecological cancer, not generally considered to be associated with cachexia, there is an elevated lipolysis promoting activity, which increased the level of hormone-sensitive lipase (HSL) in normal adipocytes [14]. The increased mobilization and use of fat in cancer patients may be due to the increased metabolic demands of the tumor. Fat constitutes 90% of the energy reserves in birds and mammals, because of the high caloriﬁc value of lipid (39.1 kJ/g), compared with carbohydrate (15.4–17.5 kJ/g). Also, in contrast to carbohydrate, triacylglycerols can be stored with very little associated water.

17.3 Energy Expenditure in Cancer Patients

In normal subjects, the basal metabolic rate (BMR) decreases in response to a decrease in food intake (e.g., in chronic starvation). Thus, in cachectic cancer patients with a reduced food intake, the BMR would be expected to be also reduced. However, a number of studies have reported that cancer patients have an increased resting energy expenditure (REE) compared with either weight-losing or weight-stable controls [15]. The major determinant of an increased energy expenditure appears to be the type of tumor [16]. Thus, while patients with pancreatic [17] and lung cancer [16] showed an increase in REE, compared with healthy controls, patients with gastric and colorectal cancer showed no such elevation in REE [16]. Although REE has been shown to be increased in patients with pancreatic cancer, both the total energy expenditure (TEE) and physical activity level (PAL) have been shown to be reduced [18], reﬂecting the patients lack of ability to move to any signiﬁcant extent, and a poor quality of life. Also in patients with pancreatic cancer the REE was signiﬁcantly greater in patients with an acute-phase response [17], possibly reﬂecting an increased metabolic demand on the liver for the synthesis of acute-phase proteins, as well as for reactions due to proinﬂammatory cytokines. There are also increased metabolic demands due to futile cycling in cancer patients. Warburg was the ﬁrst to observe that tumors have an increased demand for glucose for energy production. This is probably due to the tumor microenvironment, where oxygen levels are low and insufﬁcient for mitochondrial oxidative phosphorylation to operate effectively. In this case tumors switch to an anaerobic conversion of glucose to lactate, which is highly energy inefﬁcient yielding only 2 mol ATP/mol glucose consumed, compared with 38 mol ATP in the presence of oxygen [19]. The lactate is released from the tumor and transported to the liver where it is converted back into glucose (Cori cycle). This is also an energy consuming process requiring 6 mol ATP to convert 2 mol lactate back into glucose. The Cori cycle has been calculated to account for an additional loss of energy in the order of 300 kcal/ day [20]. Other substrates also contribute to the increased gluconeogenesis seen in

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cancer patients including glycerol, which is released from adipose tissue during lipolysis, and some amino acids, which are released from skeletal muscle due to an increase in protein degradation. Weight-losing cancer patients show a 40% increase in hepatic glucose production compared with controls, in contrast with patients with anorexia nervosa who show reduced levels [21]. There are also other metabolic changes leading to an increased energy utilization in cachexia. Thus, cachectic mice show an elevation of the triacylglycerol/fatty acid substrate cycle [22]. In the process of lipolysis one molecule of triacylglycerol is hydrolyzed to three molecules of nonesteriﬁed fatty acids (NEFAs) and one molecule of glycerol. However, in mice bearing a cachexia-inducing tumor a high proportion of the NEFA is immediately re-esteriﬁed, contributing to an increased energy demand. Energy may also be lost as heat through uncoupling of respiration from phosphorylation of ADP. The UCPs achieve this by facilitating proton leakage across the inner mitochondrial membrane. In mice, the major UCP involved in thermoregulation is UCP-1 found in BAT, although adult humans have been reported to have little, if any, BAT. However, a single study [23] that examined samples of periadrenal tissue by light microscopy revealed the presence of BAT in 80% of the samples from cachectic cancer patients, compared with 13% in age-matched controls. In humans another uncoupling protein, UCP-3, found in skeletal muscle is likely to play an equivalent role to UCP-1 in energy balance and lipid metabolism. In transgenic mice overexpressing UCP-3 in skeletal muscle there was a large reduction in adipose tissue mass, accompanied by a reduction in body weight, even though they were hyperphagic [24]. Cancer patients with weight loss have been reported to have a 5-fold increase in the levels of mRNA for UCP-3 in skeletal muscle compared with controls and cancer patients that had not lost weight [25]. This increase in UCP-3 would be expected to enhance energy expenditure, and may account for nightly sweating [19]. In experimental mice bearing a cachexia-inducing tumor levels of mRNA for UCP-1 in BAT and UCP-3 in skeletal muscle were also signiﬁcantly increased [26]. The higher rate of energy expenditure in such animals was shown by a 2-fold increase in oxidation of infused lipid to CO2, in comparison with animals bearing a tumor of the same type, which did not induce cachexia, as well as non-tumor-bearing controls [27].

17.4 Factors Governing Adipose Tissue Mass

The mass of adipose tissue is governed by the rate of synthesis of triacylglycerol from exogenous and endogenous NEFAs, and the rate of hydrolysis back to NEFAs and glycerol. Synthesis of NEFAs can arise from excess carbohydrate in the diet and they are also transported from the liver to adipose tissue as triglyceride-rich lipoproteins, such as chylomicrons and very-low-density lipoprotein. The enzyme lipoprotein lipase (LPL) is responsible for the movement of fatty acids from the blood into the adipocytes by hydrolyzing the triglyceride-rich lipoproteins, so making them available for triacylglycerol synthesis. Leptin is a neuropeptide secreted from adipocytes that modulates food intake and energy balance through its actions at speciﬁc

17.5 Mechanism of Loss of Adipose Tissue in Cachexia

receptors in the hypothalamus. Cytokines such as tumor necrosis factor (TNF)-a and interleukin (IL)-1 and -6 increase ob gene expression and leptin secretion and decrease food intake resulting in a decrease in adipose mass [28]. Leptin causes loss of fat without a rise in NEFAs or ketones. It does this by decreasing expression of fatty acid synthase, while increasing PPAR-a and the enzymes of fatty acid oxidation, and stimulates release of glycerol without a proportional release of NEFAs [29]. Lipolysis is regulated by a number of hormonal and paracrine and/or autocrine signals. The key rate-limiting step in this reaction is the hydrolysis of triacylglycerol by the enzyme HSL. Another enzyme, adipose triglyceride lipase (ATGL), removes the ﬁrst fatty acid from the triacylglycerol molecule generating NEFA and a diacylglycerol, while HSL degrades diacylglycerol into monoacylglycerol and NEFA. Thus, ATGL and HSL act coordinately in hydrolyzing triacylglycerol stores, and are responsible for more than 95% of the triacylglycerol hydrolase activity present in WAT [30]. HSL is activated by phosphorylation through protein kinase A (PKA), which in turn is activated by cAMP. Hormone production of cAMP is stimulated as a consequence of G-protein-coupled receptors acting through adenylyl cyclase. G-protein-coupled receptors can also activate mitogen-activated protein kinase (MAPK) and extracellular signal-regulated kinase (ERK) pathways. Activated ERK also increases lipolysis by phosphorylating HSL at Ser600 – one of the sites phosphorylated by PKA [31]. TNF-a has been shown to stimulate lipolysis in human adipocytes through stimulation of MAPK and ERK [32]. However, it also decreased the expression of cAMP phosphodiesterase by 50%, suggesting a mechanism by which it could increase cAMP. In contrast to HSL, ATGL is not a target for PKAmediated phosphorylation, but is regulated by an activator protein known as a/bhydrolase domain containing-5, also known as comparative gene identiﬁcation (CGI)-58. CGI-58 is bound to perilipin A, which protects triacylglycerol stores from lipolysis. After lipolytic stimulation CGI-58 is released from perilipin A and becomes available for binding to ATGL, causing an increase in triacylglycerol hydrolysis.

17.5 Mechanism of Loss of Adipose Tissue in Cachexia

The primary mechanism for the depletion of triacylglycerol in adipose tissue appears to arise from an increase in HSL [33]. Although some studies have reported a decrease in plasma LPL in patients with cancer cachexia [34], other studies [33] have reported no change in either the total LPL or the relative level of mRNA for LPL in adipose tissue of cancer patients compared with controls. Lipogenesis from glucose in WATwas increased 6-fold in the tumor-bearing state, but this was not related to the development of cachexia [35]. Most studies report an increase in serum levels of triacylglycerol and NEFA, despite evidence of an increased lipid utilization, since the rate of removal of infused lipids from the blood is increased in cachectic cancer patients [36]. The increased lipolysis in cachectic cancer patients has been suggested to arise in part from an increased b-adrenoreceptor activity [37]. Thus oral administration of the speciﬁc b1 receptor blocker atenolol, as well as the nonspeciﬁc b1/b2-

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adrenoreceptor blocker propanolol, was found to signiﬁcantly reduce REE, wholebody oxygen uptake, and CO2 production in weight-losing cancer patients. Such patients also show an increased sensitivity to adrenaline, and have increased plasma and urinary catecholamines, elevated heart rate, and increased fat oxidation [5]. These results suggest that stimulation of HSL in the adipocytes of cachectic cancer patients may be mediated through a b-adrenoreceptor. In rats with experimentally induced cachexia, anorexia is not related to changes in leptin, since both serum leptin levels and leptin mRNA in adipose tissue decreased prior to a drop in food intake [38]. In both mice and humans, leptin levels are generally proportional to the body fat content and decrease with loss of adipose mass. However, in weight-losing patients undergoing surgery for colorectal cancer omental WAT leptin mRNA levels were found to be decreased, while subcutaneous WAT showed higher levels of leptin mRNA than the same tissue of weight-stable patients [39].

17.6 Requirements of Tumor-Bearing Animals for Lipids

As already discussed, the energy demands in the tumor-bearing state are increased, resulting in an increased oxidation of lipids to CO2. It is unlikely that the tumor oxidizes lipid to any great extent, because of the low oxygen tension, but there may be a need for NEFAs, primarily polyunsaturated fatty acids (PUFAs), as growth signals to promote tumor growth. The subcutaneous adipose tissue of patients with ovarian and endometrial tumors contains a lower concentration of linoleic acid than that found in cancer-free subjects, suggesting selective loss of this PUFAs in the cancerbearing state [40]. Linoleic acid has been shown to stimulate tumor growth, both in vitro and in vivo, when the level exceeded 3.8% of the total calorie intake [41]. The stimulatory effect was attenuated by inhibitors of both cyclooxygenase and lipoxygenase (LOX). Arachidonic acid has also been shown to stimulate tumor growth [41] and malignant prostatic tissue has been found to contain lower levels of this PUFAs than benign tissue, which may be due to an increase in metabolism [42]. The critical metabolic pathway seems to occur by transformation through the 12- or 15-LOX to 12 (S)- or 15(S)-hydroxyeicosatetraneoic acids (HETEs), since LOX inhibitors induce apoptotic death in tumor cells, which can be partially inhibited by exogenous 12(S)- or 15(S)-HETEs [43]. The LOX inhibitor CV-6504 has been shown to have profound antitumor activity against murine tumors, which are generally refractory to cytotoxic agents [44].

17.7 Fat-Mobilizing Substances in Cancer Cachexia

The tumor and the hosts need for increased lipid utilization would favor those tumors capable of mobilizing triacylglycerol from adipose tissue. The ﬁrst report

17.7 Fat-Mobilizing Substances in Cancer Cachexia

indicating that nonviable preparations of Krebs-2 carcinoma were able to induce fat depletion in vivo, in a manner similar to that of viable preparations, came in 1962 [45]. This suggested that tumor metabolism alone was not responsible for the atrophy of adipose tissue. Since then substances capable of inducing lipolysis in adipose tissue have been found to be produced by the murine tumors, Ehrlich ascites carcinoma [46], a thymic lymphoma [47], sarcoma 180 [48], MAC16 adenocarcinoma [49], and rabbit VX2 carcinoma [50]. Such LMFs are not conﬁned to animal tumors. A LMF has been isolated from human A375 melanoma cells [51], in the ascites ﬂuid of patients with hepatoma [52], in the sera and ascites ﬂuid of patients with ovarian cancer [14], and in the serum and urine of cancer patients with weight loss [53]. A linear relationship was obtained between LMF activity and weight loss. In addition, LMF activity was reduced in cancer patients responding to therapy, suggesting a tumor origin [54]. It is not clear whether the LMFs from the different sources are the same material or a group of related substances, since the isolated materials differ in molecular weight. All, however, have been shown to have an overall negative charge – a unique feature of LMFs, since the natural lipolytic hormones such as adrenaline and glucagon all have positive charges. The best-characterized LMF is that produced by the MAC16 tumor, which appears to be identical with a LMF isolated from the urine of cachectic cancer patients [55]. Both materials have an apparent Mr of 43 kDa and sequence analysis showed them to be identical to a known protein, zinc a2-glycoprotein (ZAG), the function of which was previously unknown. Expression of ZAG in mouse tumors was found to correlate with the depletion of carcass lipids when they were transplanted into mice. Pure ZAG was found to stimulate lipolysis at the same concentrations as LMF, through stimulation of adenylate cyclase in a GTP-dependent process [55]. Both LMF [55] and ZAG [56] produced weight loss when administered to obese or ex-breeder mice, without a reduction in food and water intake. Loss of body weight was entirely due to loss of adipose tissue with no effect on lean body mass. There was a 3-fold increase in oxygen consumption by interscapular BAT, suggesting an increase in thermogenesis [55]. This was accompanied by an increase in expression of UCP-1. The increase in plasma NEFA may contribute to the increased expression of UCP-1 in BAT. However, in vitro experiments showed that ZAG induced a concentration-dependent increase in the expression of UCP-1 in primary cultures of BAT [57]. In addition, there were increases of between 2.5- and 3.5-fold in the expression of UCP-3 and UCP-2 in murine myotubes. These results conﬁrm that ZAG is not only capable of stimulating lipolysis through activation of HSL, but also increases the utilization of the released NEFA through an increased level of UCPs in BAT and skeletal muscle. Evidence has been presented that both LMF [58] and ZAG [56] induce lipolysis in adipocytes through interaction with a b3-adrenoreceptor. This observation correlates with the clinical observation that lipolysis occurs through a b-adrenoreceptor [57]. Induction of UCP-3 in skeletal muscle occurs through a MAPK pathway rather than through a b3-adrenoreceptor [57]. Other studies have also shown that chronic treatment of mice with a speciﬁc b3-adrenoreceptor agonist markedly increased the expression of UCP-1 in BAT [59]. b-Agonists are known to stimulate skeletal muscle hypertrophy in animals, and studies with both LMF [55] and ZAG [56] show a

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tendency to increase lean body mass despite the loss of adipose tissue. LMF has been shown to stimulate protein synthesis in murine myotubes in vitro through an increase in intracellular cAMP, since it was attenuated by inhibition of adenylate cyclase and by an antagonist of the b3-adrenoreceptor [60]. There was also a decrease in protein catabolism mediated through the ubiquitin–proteasome proteolytic pathway. LMF was shown to increase the energy utilization of mice [61], producing a signiﬁcant increase in oxidation of both glucose and triolein to CO2. Thus, the increased oxidation of lipid in cachectic cancer patients probably arises from the production of LMF. Although ZAG was known to be produced by the liver and cachexia-inducing tumors, recent evidence [62] suggests that it is also expressed in WATand BAT. There was a 10-fold increase in mRNA levels of ZAG in WATof mice with cachexia, that had lost 61% of their fat mass. ZAG mRNA levels in BATwere also increased 3-fold, while liver ZAG mRNA was increased 2-fold in comparison with non-tumor-bearing controls. In contrast, leptin mRNA was suppressed 33-fold, while adiponectin mRNA levels remained unchanged. Expression of ZAG mRNA levels in 3T3-L1 adipocytes was enhanced 3.3-fold with a b3-adrenoreceptor agonist and 55-fold with the synthetic glucocorticoid dexamethasone. Further studies have shown ZAG to increase its own expression through a b3-adrenoreceptor-mediated pathway [63]. Upregulation of ZAG expression during cachexia is probably due to increased cortisol production, since in a murine cachexia model the increase in ZAG expression in WAT could be blocked by the glucocorticoid receptor antagonist RU38 486 [63]. ZAG was also shown to upregulate its own expression, and this was attenuated by the b3-adrenoreceptor antagonist SR59 230A, suggesting that it was mediated through a b3-adrenoreceptor. Eicosapentaenoic acid (EPA), which has been shown to preserve adipose tissue in cachectic mice, caused downregulation of ZAG expression through interference with glucocorticoid signaling [64]. EPA has also been shown to attenuate lipolysis in WAT, by preventing stimulation of adenylyl cyclase, through an inhibitory guanine nucleotide-binding protein [65]. In home-living cachectic patients, with advanced pancreatic cancer, receiving an energy- and protein-dense oral supplement containing EPA, there was an increase in TEE and PAL [18]. The increased PAL may reﬂect an improved quality of life, but this most likely arises from an ability of EPA to preserve lean body mass. Patients receiving megestrol acetate gain weight through an increase in adipose tissue and water, but there is no effect on fat-free mass [66]. As a result, such patients show no improvement in the Karnofsky index – a measure of the activity level. This suggests that agents only attenuating the increased lipolysis may not be sufﬁcient for the treatment of cancer cachexia.

17.8 Conclusion

Mobilization of triacylglycerol from adipose tissue in the cachectic state is indicative of the increased energy demands imposed by the tumor. The ability of tumors to produce LMF may be a general phenomenon and not restricted to those where gross

17.8 Conclusion

loss of adipose tissue is evident. LMF caused upregulation of UCP-2 in tumor cells, which is thought to be involved in the detoxiﬁcation of free radicals and reduced the growth-inhibitory effect of free radical generators [67]. During the past 15 years evidence has been provided that in addition to its lipid storage capacity, WAT is capable of releasing a growing number of products into the blood stream. It now seems that in addition to production of LMF by certain tumors, WAT is a key player in this process, producing a speciﬁc LMF, ZAG, which in turn is able to stimulate

Figure 17.1 Interactions between tumor, WAT, BAT, skeletal muscle, and liver in the cachetic state. TG-LP, triglyceride-rich lipoprotein; AR, adrenoreceptor; other abbreviations are given in the text [69].

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lipolysis. It thus joins a growing list of adipokines that signal changes in the mass of WAT and energy status to other organs that control fuel usage. These also include TNF-a and IL-1b and -6, which are potent inhibitors of LPL, while TNF-a also suppresses key enzymes that contribute to lipogenesis and opposes the recruitment/ differentiation of mature adipocytes [68]. The interaction between these factors is not known, but the combined effect is likely to lead to extensive depletion of ZAG in WAT in the cachectic state. An overview of this process is provided in Figure 17.1 [69].

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18 Obesity and Diabetes: Lipotoxicity Christopher J. Lelliott, Matej Orešic, and Antonio J. Vidal-Puig 18.1 Introduction

Probably the most important emergent health crisis is the increase in prevalence of obesity. Obesity causes mechanical and psychological disorders, but more importantly it causes severe metabolic complications. No one doubts that the development of obesity requires a chronic state of positive energy balance. However, it is intuitively less clear why expansion of adipose tissue should be associated with insulin resistance, diabetes, dyslipidemia, fatty liver, and/or cardiovascular complications, collectively known as the metabolic syndrome [1, 2]. Current research on the pathogenesis of obesity has predominately focused on aspects related to food intake and energy expenditure as major determinants of energy balance. It has been incorrectly assumed that while there is a state of positive energy balance the adipose tissue would simply expand to meet storage demands. However, it is clear that the current epidemic of obesity poses unprecedented stress on the adipose tissue to reach previously unknown levels of expansion. The processes of adipogenesis and adipocyte growth are complex, and require the coordinated sequential activation of speciﬁc genetic pathways. Disturbance of speciﬁc components of these pathways, or of their sequential activation, results in defective adipose tissue development or function. This compromises not only the lipid buffer capacity of the adipose tissue, but also results in changes in synthesis and secretion of speciﬁc adipokines with important roles controlling insulin sensitivity. In this chapter, we review the evidence supporting the hypothesis that the link between obesity and these metabolic complications may be related to organ-speciﬁc toxic effects induced by excess of nutrients in the form of reactive lipid species. The concept of lipotoxicity describes a situation where ectopic deposition of reactive lipid species in organs different from adipose tissue induces a toxic reaction leading to insulin resistance, oxidative stress, endothelium reticulum stress, and ultimately dysfunction and cellular death. In this context, the most commonly studied tissues undergoing lipotoxicity include skeletal muscle, heart, liver, and pancreas. In the

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context of obesity we consider exhausted capacity for adipose tissue expandability as an important determinant leading to lipid leakage into other organs.

18.2 White Adipose Tissue at the Center of Lipid Homeostasis and Delivery

White adipose tissue (WAT) senses and responds to the nutritional condition of the organism. In the postprandial condition, adipocytes are stimulated by insulin to take up glucose and fatty acids from the circulation, and to store them as triacylglycerol. Insulin also inhibits the release of fatty acids (lipolysis) from the fat cell [3]. Conversely, the reduction of circulating insulin together with stimulation from other peripheral factors, such as catecholamines, induces the release of fatty acids from the adipocyte for delivery to other tissues. Whilst this regulation is tightly controlled, in the obese patient as adipose tissue expands it becomes progressively resistant to the action of insulin. Under these conditions insulin-stimulated glucose and fatty acid uptake is impaired, and lipolysis is incompletely suppressed. Thus, fatty acids remain in the circulation longer and are now available to the rest of the body. In fact, the amount of stored fat may not be the main determinant of insulin resistance. It is not uncommon to identify morbid obese individuals that maintain their insulin sensitivity. The converse is also true – there are patients that could be considered inappropriately too insulin-resistant for their degree of obesity. Therefore, the question is to determine the key difference between these two scenarios. Our hypothesis is that intrinsic differences in the degree of adipose tissue expandability and functionality may account for the different susceptibility to obesity associated metabolic complications.

18.3 Insulin Resistance in Adipocytes Disrupts the Balance between Lipid Storage and Secretion

The adipocyte is a cell speciﬁcally adapted for the storage and release of energy in the form of lipids. Impaired insulin action in the adipocyte results in reduced postprandial uptake of circulating lipids and glucose, together with an incomplete shut-off of lipolysis. The net result is a delay in the clearance of circulating lipids and glucose after a meal and continued exposure of other peripheral tissues to these elevated substrates. The pathology of lipotoxicity is thought to arise from the ectopic deposition of lipids into these tissues that are not adapted to safely handling or storing large amounts of lipid molecules.

18.4 Scenarios that may Result in Ectopic Fat Deposition

We consider that there are a number of different scenarios that could result in lipotoxicity in peripheral tissues, as shown in Figure 18.1.

18.4 Scenarios that may Result in Ectopic Fat Deposition

Figure 18.1 Defects in adipose tissue expansion pattern drives the development of lipotoxicity in peripheral tissues. The adipose tissue must adapt to be able to handle increased amounts of energy delivered to it. The balance of existing adipocyte expansion and the recruitment of new adipocytes determines the metabolic outcome for peripheral tissues and the whole organism. Adipose tissue expansion that is the result of new adipocyte recruitment, hyperplasty, allows the adipose tissue to store more lipids, without detrimental effects on insulin sensitivity or cytokine output (top right). Adipose tissue expansion due mostly to the

enlargement of existing adipocytes adversely affects insulin responses to substrate storage and to the inhibition of lipolysis. In addition, the cytokine output pattern from the adipose tissue is deranged. Thus, lipids are available in circulation for uptake by peripheral tissues not capable of handling them in large amounts. The combination of ectopic lipid deposition with a detrimental hormonal pattern results in lipotoxicity (bottom left). If adipose tissue storage capacity is below storage demands, as in lipodystrophies, the excess lipids are also ectopically deposited in peripheral tissues (bottom right).

18.4.1 Altered Plasticity of the Adipose Tissue: A Shift in Expansion Towards Hypertrophy

Efﬁcient expansion of the adipose tissue in response to overnutrition requires a titrated process that requires a tight balance between adipocyte enlargement, hypertrophy, and recruitment of new preadipocytes – hyperplasty. This process requires close contact between mature adipocytes and preadipocytes to communicate when a speciﬁc maximum threshold of adipocyte growth has been reached and differentiation of new storage units is required. Changes in the mechanisms controlling this balance may result in delayed recruitment of new adipocytes or hypertrophic adipose tissue, whereas early recruitment of preadipocytes may lead to a hyperplastic state [4]. Based on our research, our interpretation is that hypertrophic

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adipose tissue reﬂects a failure of new preadipocytes to differentiate, resulting in overstretched mature adipocytes attempting to store the excess of nutrients [5, 6]. Current thinking suggests that a critical difference between the two types of adipose is due to the size of the adipocyte. Typically hypertrophic adipocytes are associated with altered gene expression, insulin resistance (impaired glucose and fatty acid accumulation, incomplete suppression of lipolysis) [7, 8], and worsened adipokine proﬁle [9]. The adipocyte is known to produce at least 20 factors (known as adipokines) that are able to inﬂuence diverse aspects of metabolic homeostasis. Hypertrophic adipocytes have an altered pattern of adipokines that overall has a negative affect on peripheral metabolism (Table 18.1) [4, 9], which promotes lipid deposition, but not the mechanisms to dispose of the lipids. Considering that adipose tissue is associated in signiﬁcant amounts with skeletal muscle, heart, and liver, a paracrine effect of these adipokines on other peripheral tissues must be considered. The speciﬁc reason why an enlarged adipocyte has a worse proﬁle is unclear, although a number of hypotheses have been proposed. These include changes in membrane mechanical stress due to expansion of the adipocyte, distorted properties of lipids and cholesterol in the plasma membrane or lipid droplets, and changes in the interaction of the adipocyte with the extracellular matrix. In contrast, hyperplastic adipose tissue, with increased proliferation of smaller adipocytes with an improved metabolic proﬁle, may be beneﬁcial, as seen with treatment with certain thiazolidinedione-class peroxisome proliferator-activated receptor (PPAR)-c agonists [10]. Hyperplastic adipose tissue may be particularly beneﬁcial since it has an increased amount of total membrane and therefore an increased buffer capacity to store lipids. 18.4.2 Impaired Fat Deposition Capacity in Adipose Tissue

Lipotoxicity in peripheral tissues may develop since adipocytes may be unable to store the huge amount of lipid as demanded and so they remain in circulation, available for deposition in other tissues. Hypertrophic, insulin-resistant adipocytes have an altered balance of lipolysis and storage and would contribute to a higher maintenance of fatty acids in the circulation. There are extreme examples showing that defects in proteins needed for lipid deposition – such humans with defective LPL enzyme activity suffer from chylomicronemia and hypertriglyceridemia [11]. However, we think that in the current obesogenic climate, metabolic disturbances are more likely due to subtle genetic defects that previously did not cause metabolic problems, but now produce an adipose tissue with an inappropriately limited maximal expansion insufﬁcient for the modern-day storage demands. In this context, adipose tissue may be incapable of expanding sufﬁciently by recruitment and differentiation of preadipocytes, or by apoptosis of existing adipocytes. The range of conditions, known as lipodystrophies where bodily adipose tissue is less than necessary, provides a dramatic illustration of whole body metabolism with insufﬁcient adipose storage. Various forms of lipodystrophy are associated with

Table 18.1

Summary of known adipose tissue-generated hormones that may impact upon the development of insulin resistance and lipotoxicity Circulating levels in obesity

Central effect

Effect on adipocyte glucose/fatty acid flux

Liver

Metabolic actions on muscle

b-Cell survival/insulin production

Net effect of change in obesity for development of lipotoxicity

Leptin

elevated, but leptin resistance may occur

anorexigenic

antiadipogenic in vitro, promotes lipolysis

increases insulin sensitivity and b-oxidation via AMPK

enhances insulin sensitivity in muscle and b-oxidation via AMPK; decreases IMCL

inhibits insulin release; possibly antiapoptotic

Adiponectin

reduced

orexigenic

promotes lipogenesis in vitro

increases insulin sensitivity and b-oxidation via AMPK

increases b-oxidation via AMPK

may be beneﬁcial for insulin secretion in insulin-resistant models

TNF-a

elevated

anorexigenic

antiadipogenic, lipolytic, inhibits glucose uptake

insulin resistance, although no effect in healthy humans infused with TNF-a; stimulates fatty acid and triacylglycerol biosynthesis

reduced insulinstimulated glucose uptake via multiple pathways; inhibits AMPK and promotes diacylglycerol and ceramide formation

IL-6

elevated

anorexigenic

insulin resistance, lipolytic

insulin resistance, in part by inhibition of insulin signaling via SOCS-3; stimulates fatty acid and cholesterol biosynthesis

in obesity, thought to reduce glucose uptake, but IL-6 is elevated in exercise and related here to enhanced insulin sensitivity

in vitro, impairs glucose-stimulated insulin secretion, proapoptotic; unclear role in vivo since transgenic expression of TNF-a in pancreas results in insulitis, not diabetes islet-speciﬁc transgenic expression of IL-6 in pancreas results in b-cell hyperplasia and insulitis

redirection of lipids from adipose to nonadipose for oxidative disposal –although some evidence it may be proinﬂammatory redirection of lipids from adipose to nonadipose, but reduced oxidative capacity results in lipotoxicity redirection of lipids from adipose to nonadipose, but reduced oxidative capacity results in lipotoxicity

redirection of lipids from adipose to nonadipose; may result in lipotoxicity

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(Continued)

18.4 Scenarios that may Result in Ectopic Fat Deposition

Adipokine

(Continued)

Adipokine

Circulating levels in obesity

Central effect

Effect on adipocyte glucose/fatty acid flux

Liver

Metabolic actions on muscle

b-Cell survival/insulin production

Net effect of change in obesity for development of lipotoxicity

Resistin

elevated

inhibits glucose uptake

Reduced insulin sensitivity, increased hepatic glucose output

reduced glucose uptake, fatty acid metabolism

Retinol binding protein-4

elevated

evidence for anorexigenic effect with intracerebroventricular administration unknown

reduced insulin sensitivity

elevated

unknown

increases gluconeogenesis, reduces insulin sensitivity; promotes lipid accumulation reduces glucose output, independent of insulin, in hepatocytes

redirection of lipids from adipose to nonadipose, but reduced oxidative capacity results in lipotoxicity to be fully determined, but appears to be prolipotoxic

Pre-B cell colonyenhancing factor

promotes insulin resistance, but not defects in glucose uptake in cell cultures promotes lipogenesis, glucose uptake

impaired glucosestimulated insulin secretion in vitro, but at low concentrations may be a survival factor unknown

may be upregulated in obesity to enhance metabolic status and prevent lipotoxicity

Omentin-1

reduced

unknown

not known

not clear, although omentin is expressed predominantly in omental adipose tissue

increased insulin sensitivity, glucose uptake

may be a b-cell survival factor since circulating visfatin is increased with progressive b-cell deterioration unknown

AMPK, AMP-activated protein kinase.

enhances glucose uptake

unclear but may be beneﬁcial

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Table 18.1

18.4 Scenarios that may Result in Ectopic Fat Deposition

a number of mutations in genes including PPARG, LMNA/C, BCSL, and AGPAT2, as well as with the use of antiretroviral therapy for HIV [12]. In all of these disorders, the lipodystrophy is associated with glucose intolerance, insulin resistance, and dyslipidemia. Whilst the mechanisms behind the loss of adipose tissue are diverse, a diminished ability of the adipose tissue to deposit lipids means that excess glucose and lipids remain in circulation, and accumulate in other peripheral tissues. In addition, the lack of adipose tissue means a reduced output of protective adipokines such as leptin and adiponectin, which would normally contribute to normal metabolic homeostasis. Interestingly, in mouse models for lipodystrophy where mice are genetically modiﬁed to lack adipose tissue, the metabolic disorders can be recovered by transplantation of normal adipose tissue [13], which demonstrates that defects in adipose tissue storage capacity and adipokine output are a primary source for the development of lipotoxicity in other peripheral tissues. With respect to obesity and insulin resistance, we consider that there are strong parallels with lipodystrophy. In our opinion, obesity-associated metabolic effects may occur when there is a mismatch between adipose tissue expandability and adipose tissue storage requirements. In short, metabolic problems associated to obesity may occur when the obese patient cannot become more obese because their adipose tissue has reached its maximal expansion. 18.4.3 Inappropriate Balance of Substrate Uptake and Oxidative Capacity in Peripheral Tissue

When the adipose tissue fails to correctly store lipids, circulating fuel is ﬁnally deposited in other peripheral tissues such as skeletal muscle, pancreas and liver. These tissues are not passive in the handling of excess lipids, but attempt to adapt to the incoming fuel load. Even relatively short periods of high-fat feeding increases the amount of fatty acid clearance and oxidation into muscle [14, 15]. Disposal of fatty acids by oxidation would be considered the best alternative when dealing with lipids in a tissue that is unable to store safely large amounts of them (Figure 18.2). However, prolonged exposure of muscle to excess of fatty acids results in a diminished fatty acid oxidation capability, elevated reactive lipid deposition, insulin resistance and impaired glucose uptake. The accumulation of lipids in nonadipose peripheral tissues may also be a consequence of a primary defect in this peripheral tissue. For example studies with healthy relatives of type 2 diabetic sufferers have revealed that oxidative metabolism gene expression and mitochondrial function in the skeletal muscle of this group appears lower than in subjects without a family history of type 2 diabetes [16–19]. This suggests a genetic predisposition to lower skeletal muscle oxidative capacity, which is required when the tissue is threatened with lipotoxicity. A second factor that would contribute to increased peripheral lipid deposition is the uptake capacity of the tissue. Using lipoprotein lipase (LPL) as an example, it has been suggested that the relative level of LPL activity, and thus fatty acid supply, between different tissues could be an important determinant for the peripheral

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Figure 18.2 Ectopic fat deposition may be enhanced by a mismatch in uptake and disposal rates. The accumulation of lipids in a tissue can be thought of as a balance between lipid uptake capacity and oxidation. In situations where excess lipids can be more rapidly oxidized for disposal than accumulated by the tissue, insulin sensitivity can be maintained since toxic lipid products are unable to be built up and

metabolized further (bottom left). If, however, oxidative capacity is lower than the rate of lipid intake, a build-up of lipid occurs and the resulting pools of lipid may be subject to the generation of lipotoxic products (bottom right). Familial studies may show that a lack of peripheral tissue oxidative capacity may predispose to insulin resistance and type 2 diabetes.

localization of lipid deposition [20]. An increased activity in WAT versus the muscle would be protective for muscle lipotoxicity, at the expense of increased adipose tissue expansion. Conversely, a predisposition towards a higher ﬂux of substrates to the muscle by elevated LPL expression may also result in increased muscle lipid deposition and, consequently, lipotoxicity. The role of substrate ﬂux has been tested using transgenic mice. Overexpression of LPL in skeletal muscle results in an elevation in muscle triacylglycerol content with concomitant decrease in insulinstimulated glucose uptake [21]. Similarly, hepatic overexpression of LPL also raises liver triacylglycerol content and blunts insulins ability to suppress glucose output [21]. Interestingly, levels of muscle LPL are increased by exercise training, together with increases in carnitine palmitoyl transferase (CPT)-1, PPAR-c coactivator (peroxisome proliferator-activated receptor-c coactivatorPGC)-1a, and mitochondrial number and activity [22, 23], thus suggesting that the balance between lipid uptake and oxidative capacity reﬂects the outcome of insulin sensitivity. The converse situation states that impaired fatty acid uptake in the muscle is beneﬁcial to muscle insulin sensitivity. Mice lacking fatty acid transporter CD36/fatty acid

18.5 Mechanisms Contributing to the Lipotoxicity in the Peripheral Organs

translocase (FAT) on a chow diet are more insulin-sensitive than wild-type and are partially protected from a high-fat diet [24], with insulin-sensitive muscles despite hypertriglyceridemia.

18.5 Mechanisms Contributing to the Lipotoxicity in the Peripheral Organs 18.5.1 Lipotoxicity in Skeletal Muscle

Obesity is associated with insulin resistance in skeletal muscle and reduces its insulin-stimulated glucose uptake capacity. In addition, oxidative capacity is decreased in skeletal muscle from insulin-resistant humans, offering a mechanism for further weight gain. Importantly, the development of insulin resistance in muscle is associated with an accumulation of intramyocellular lipids (IMCL). Of note, increased IMCL levels are found in lean, glucose-tolerant offspring of parents who both have type 2 diabetes, when compared to those with no family history of type 2 diabetes [25]. Also, its oxidative capacity appears to be impaired in obese and type 2 diabetes sufferers related to markedly reduced mitochondrial size, even when related to ﬁber type [26]. In addition, there is a deﬁciency of sarcolemmal-located mitochondria in the skeletal muscle in both obese and type 2 diabetes sufferers [27]. This is potentially relevant to insulin resistance since these mitochondria are thought to provide energy for processes occurring at the cell surface, such as ion transport, glucose uptake, and signal transduction. An apparent paradox exists here since well-trained humans, despite being insulin-sensitive, also have signiﬁcant stores of IMCL [25], and endurance training elevates IMCL content [28]. Here, the IMCL presumably acts as a readily accessible store of fatty acids for oxidation. This suggests that more important than the accumulation of fat is the type of fat accumulated. We have shown that accumulation of triacylglycerols in peripheral organs in the context of overnutrition may be a physiological protective adaptation that allows the storage of fatty acids as safe triacylglycerols [5]. Conversely, accumulation of nutrients as reactive lipid species is a more direct pathogenic mechanism linking overnutrition and insulin resistance. 18.5.1.1 Randle Hypothesis and its Successors The original hypothesis for why excess fatty acids in skeletal muscle causes insulin resistance was proposed by Randle in 1963 [29]. The essence of the hypothesis is that glucose and fatty acid oxidation are in competition with each other, and that glucose uptake and utilization is limited in tissues where fatty acid oxidation can be used for cellular energy requirements. Thus, an overload of fatty acids to a tissue such as muscle will impair glucose utilization – an event that has been shown both in vitro and in vivo. At the molecular level, the interplay of glucose and fatty acid metabolism is well established, since both glucose and fatty acid synthetic and oxidation pathways share a number of molecules from one pathway that are able to

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feed in to or cross-regulate reactions and processes in the alternative pathway (Figure 18.3). At the pathway level, Randle hypothesized that there are a number of key points for cross- and counter-regulation between the glucose and fatty acid oxidation pathways, so fatty acid overload results in a build-up of glucose-6-phosphate, which inhibits hexokinase and generates an increase in intracellular glucose to prevent glucose uptake. Since then, the hypothesis has identiﬁed signaling molecules as alternate effectors of fatty acid-induced insulin resistance. A number of studies using in vivo imaging techniques have suggested that in humans, increased plasma free fatty acid concentrations result in decreased glucose-6-phosphate levels that precede a drop in muscle glycogen synthesis – a ﬁnding also seen in type 2 diabetic humans [30–32]. From this, impaired glucose transport and therefore disposal has been highlighted as the primary defect in insulin resistance in skeletal muscle [33]. In addition, elevated plasma free fatty acid concentration causes a decrease of glucose concentrations, suggesting a direct inhibition in glucose transport and a loss of insulinstimulated phosphoinositide-3-kinase (PI3K) activity [34]. Thus, the hypothesis has emerged that triggers of insulin resistance, including free fatty acids, affect the very earliest steps of glucose disposal and the insulin signaling pathway required to control this. 18.5.2 Molecular Mechanisms for the Generation of Muscle Lipotoxicity

Fatty acids enter the cell via a variety of mechanisms including ﬂip-ﬂop diffusion across the plasma membrane in combination with transport mediated by the CD36/FAT transporters. Upon entry into the cell, fatty acids are rapidly converted to fatty acyl-CoA derivatives by acyl-CoA synthases and fatty acid transporter protein, and transported by fatty acid-binding proteins. From here, fatty acyl-CoAs then either enter the mitochondria for b-oxidation or are converted to stored triacylglycerols via diacylglycerol intermediates. Here, the adipose tissue plays an important role in the supply of lipids to other peripheral tissues in times of fasting. The conversion of the muscle from a glucose-oxidative fed state to a fatty acid-oxidative fasting state requires that the adipose tissue is able to deliver or retain fatty acids at the appropriate times. The idea of metabolic ﬂexibility has been proposed as a central hypothesis by which the muscle, or other important metabolic tissues, is able to adapt to changing energy demand with alternate fuel supplies [35]. In the context of insulin resistance, this metabolic ﬂexibility is impaired or inﬂexible due to inappropriate fuel supply from the adipose tissue and incomplete switching between metabolic pathways. Thus, with time, ectopic deposition of reactive lipid species occurs due to the inequality between fatty acid supply and utilization. 18.5.2.1 Type of Lipids is More Important than the Amount of Fat Deposited Following the ﬁnding that free fatty acids and reactive lipid species can interfere directly with insulin signaling, there has been much speculation about which molecule or molecules are responsible for this. The most-studied candidates include

18.5 Mechanisms Contributing to the Lipotoxicity in the Peripheral Organs

Figure 18.3 Randle hypothesis and interplay of glucose and fatty acid metabolism. In the original Randle hypothesis, it was thought that exposure of tissues to high levels of fatty acid, such as palmitate, would cause a switch in metabolism towards a preferential oxidation of the fatty acid. b-Oxidation of fatty acid would elevate acetyl-CoA/CoA and NADH/NAD þ ratios, which would inactivate pyruvate dehydrogenase (PDH). Citrate levels would increase and in turn inactivate phosphofructokinase (PFK)-1. A build-up of PFK substrate fructose 6-phosphate then inactivates hexokinase (HK) and in turn blocks glucose

uptake. Interestingly, fatty acid metabolism and glucose metabolism are coordinately regulated and intricately connected by common intermediated. Citrate can be diverted from the mitochondria and converted to acetyl-CoA for channeling into de novo fatty acid synthesis, via acetyl-CoA carboxylase (ACC) and fatty acid synthase (FAS). Flux through this pathway increases the concentration of malonyl-CoA, a potent inhibitor of CPT-1. Thus, glucose and fatty acid metabolism are able to regulate each other to switch between different sets of metabolic pathways.

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fatty acids or fatty acyl-CoA derivatives themselves, or alternatively metabolites deriving from fatty acids, including triacylglycerols, diacylglycerols, sphingolipids, and ceramides. It has been argued that elevated triacylglycerols in muscle of insulinresistant humans is merely a marker for more deleterious lipid species derived from fatty acids [36]. The type of fatty acids that are most consistently linked to metabolically deleterious consequences in humans and rodents are saturated fatty acids. In many insulin and metabolically related systems, such as insulin signaling and glucose utilization, saturated fatty acids show a greater impairment of the given system than unsaturated fatty acids. Unsaturated fatty acids are preferably oxidized and disposed of, suggesting that saturated fatty acids are more likely to accumulate. Finally, unsaturated fatty acids are thought to be metabolized to ligands for a number of transcription factors that have a beneﬁcial metabolic effect, such as PPAR-a, so a diet proportionally rich in saturated fatty acids will essentially dilute out these lipids making them less detectable by the lipid sensor mechanisms. 18.5.2.2 Diacylglycerols and Insulin Resistance Current evidence suggests that saturated fatty acids and their derivatives play the roles in stimulating insulin resistance and lipotoxicity in the muscle to varying degrees (Figure 18.4). Long-chain fatty acyl-CoA accumulation, either by an increase in exogenous delivery or decreased mitochondrial oxidative capacity, may be converted to diacylglycerol moieties. Notably in both, diacylglycerol and long-chain fatty acyl-CoA levels are increased in insulin-resistant muscle [37, 38]. Diacylglycerols may also accumulate in hyperglycemia conditions via de novo synthesis from phosphatidic acid and through an elevated turnover of phosphatidylcholine. Diacylglycerols are cofactors of conventional and novel protein kinase C (PKC) isoforms. In addition, diacylglycerol-mediated PKC activation appears to be augmented by nonesteriﬁed fatty acids and CoA derivatives, enhancing the effect of an elevated fatty acid inﬂux. The bulk of current data suggests that novel PKCs PKCe and PKCq are more likely to be involved in insulin signaling inhibition in skeletal muscle and liver [39]. Diacylglycerol-activated PKC phosphorylates insulin receptor substrate (IRS)-1 on serine residues that impede activating tyrosine phosphorylations by an activated insulin receptor. Since insulin signaling is impaired at the level of IRS-1, the activities of other critical signaling pathways downstream of IRS-1 such as the PI3K/Akt pathway are also blunted. This may be especially important in the muscle since both insulin-stimulated glucose uptake and glycogen synthesis is controlled via the IRS/PI3K/Akt pathway, and so diacylglycerols will impede muscle glucose disposal. 18.5.2.3 Ceramides and Insulin Resistance Evidence is accumulating for ceramide (and its derivatives) being a key mediator of fatty acid-mediated insulin resistance (Figure 18.3). Ceramide generation occurs both via the hydrolysis of sphingomyelin as a second messenger or de novo synthesis occurs via a four-enzyme pathway. This pathway is largely dependent on the supply of long-chain saturated fatty acids, with the key rate-limiting step serine-palmitoyl transferase providing the fatty acid chain speciﬁcity for the ﬁrst assembled fatty

18.5 Mechanisms Contributing to the Lipotoxicity in the Peripheral Organs

Figure 18.4 Accumulation of lipid species interferes with molecular processes and cell metabolism. Fatty acid uptake via flip-flop mechanisms or protein-based translocation results in the generation of a number of lipid species. Among those known to be involved in the development of lipotoxicity include fatty acid-CoAs themselves, diacylglycerols (diacylglycerols), which in turn are converted to triacylglycerols, and ceramides, generated by the serine palmitoyltransferase pathway, for which saturated fatty acids are a prerequisite. Fatty acid-CoAs, diacylglycerols and ceramides are able to interfere with various components of the insulin signaling pathway via intermediatory protein kinases such as PKCs (to catalyze inhibitory serine phosphorylations) and protein

phosphatases like PP2A. Inhibition of the insulin signaling pathway results in an insulin resistance, with blockade of glucose uptake and disposal in tissues such as muscle. Ceramides are also thought to impact directly on mitochondrial function by inhibition of the ETC that induces ROS and may trigger apoptosis via cytochrome c (CytC) export. Fatty acids, via ceramides, also activate pathways that alter nuclear gene expression such as the JNK/c-Jun and IkB/NFkB pathways. These pathways are also shared in inflammatory responses, and enhance the link between obesity and low-grade chronic inflammation. Finally, genetic factors may alter metabolic capabilities that predispose to insulin resistance.

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acid moiety [40]. Due to this requirement of saturated fatty acids, there is an obvious link between saturated lipid oversupply to the muscle and generation of ceramide. The putative role for ceramides in the development of peripheral insulin resistance has been proposed based on a number of lines of evidence. These include an increase in ceramide levels in tissue from insulin-resistant rodents and humans, the ability exogenously applied palmitate and ceramide to induce insulin resistance in cell culture, and that pharmacological inhibition of ceramide synthesis improves metabolic status in both cell culture and diabetic rat models [38, 40–42]. The mechanism by which ceramides are able to inhibit insulin signaling is at present unclear, but ceramide treatments are known to inhibit several of the signal transduction components downstream of the insulin receptor [40, 43]. There are reports that ceramideinducing inhibitory serine phosphorylations of IRS-1, inhibition of PI3K activity and blockade of Akt/protein kinase B (PKB) action, probably via a combination of dephosphorylation by protein phosphatase 2A (PP2A) and inhibitory phosphorylation by atypical (diacylglycerol-independent) PKCz. Of these, the action on Akt/PKB has the most supportive data. There is also evidence for ceramides activating c-Jun N-terminal kinase (JNK) and IkB kinase (IKK)-b pathways. Both of these signaling pathways have been implicated in the development of insulin resistance from the perspective of inﬂammation, which may impact upon muscle metabolism both at the levels of signaling components such as IRS-1 and via the transcriptional activity of c-Jun and nuclear factor-kB. However, the full details of these pathways are still to be elucidated. Consistent with the mitochondrial dysfunction seen in the insulinresistant subjects, ceramides appear to also have direct effects on mitochondria, especially since many of the enzymes required for ceramide synthesis are localized here, in addition to the endoplasmic reticulum [40]. Ceramide analogs inhibit the electron transport chain (ETC) at complex III and promote reactive oxygen species (ROS) generation – an additional deleterious factor [44, 45]. There is also some evidence that ceramides may also trigger apoptosis by increasing mitochondrial permeability to cytochrome c via Bcl-2 proteins.

18.6 Impaired Oxidation as a Trigger for Lipotoxicity

Whilst increased ﬂux of lipids into nonadipose tissue organs is one possible way for intracellular lipids to accumulate, there is also increasing evidence that impaired disposal of these lipids is also a key factor. Increased oxidation of accumulated lipids by the mitochondria is seen as one mechanism by which skeletal muscle insulin resistance could be reversed pharmacologically. Indeed various studies have shown that in obese or insulin-resistant rodents and humans, mitochondrial oxidative capacity is reduced at both gene expression and functional levels [17–19]. This reduction in mitochondrial activity has also been suggested as a primary mechanism that predisposes certain individuals to the development of insulin resistance and obesity since offspring of subjects with type 2 diabetes show a reduction in

18.6 Impaired Oxidation as a Trigger for Lipotoxicity

mitochondrial enzymes or in key transcriptional regulators of mitochondrial action, such as PGC-1. This implies that in pre-insulin-resistant states, the oxidative capacity of mitochondria is already compromised and less able to cope when insulted with increased lipid loads. Accumulated lipids may additionally become deleterious through the generation of ROS. This can occur through the deposition and conversion of lipid moieties to reactive peroxides or through the generation of mitochondrial ROS via impaired ETC activity. Both PGC-1a and -1b are important regulators of mitochondrial activity [46, 47]. In addition, PGC-1a is essential for the generation of an adequate ROS defense system [48], so in insulin-resistant states where PGC-1 action is inhibited, the double-whammy of reduced oxidative capacity and increased ROS production occurs. 18.6.1 Adipocytokines Proinflammatory Activity Contributes to Lipotoxicty in Skeletal Muscle

Hypertrophic adipose tissue not only has a dysregulation of lipid storage which contributes to elevated nonesteriﬁed fatty acid levels and lipotoxicity, but also has an altered pattern of adipokine output that impacts on whole-body energy homeostasis. The typical pattern of adipokines and related inﬂammatory markers release from hypertrophic adipose tissue is thought to have a dual effect on other nonadipose tissue peripheral tissues. (i) The pattern of cytokines points towards a reduced insulin sensitivity, decreased glucose disposal, and inhibition of insulin-suppressed glucose output. (ii) Peripheral oxidation capacity is reduced, by decreased adiponectin and increased tumor necrosis factor (TNF)-a. This results in an increase in intracellular lipids and the associated hazards concerning lipotoxicity. It is worth noting that adipocytes are not the only cell type in adipose tissue that produce cytokines or other factors that can impact on peripheral metabolism. 18.6.1.1 Adipose Tissue Macrophages as Key Players for Lipotoxicity Adipocytes are not the only contributors to the cytokine proﬁle of the WAT. Many proinﬂammatory cytokines, such as interleukin (IL)-6, IL-1 and CCL2, and antiinﬂammatory cytokines, such as adiponectin, have been identiﬁed and shown to be dysregulated in WAT from obese subjects [49, 50]. Accumulating evidence shows that activated macrophages occurring in the WAT from obese rodents and humans can contribute to secretion of adipokines [51, 52] and adipocyte turnover, especially in obese tissue. As yet, it is not clear whether these activated macrophages are recruited from the circulation during the induction of obesogenic development or lie dormant in the tissue awaiting activation by engorged adipocytes [53]. Nonetheless, they appear to play an important part of a cycle of cytokine release that results in an unfavorable pattern of adipokines released from the adipocyte. 18.6.1.2 Signaling Effector Pathways in Lipotoxicty Inﬂammatory signals and lipids share some similar pathways that are known to mediate insulin resistance. Inﬂammatory cytokines such as TNF-a and IL-6 are

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known to increase IRS-1 serine phosphorylation and increase suppressor of cytokine signaling (SOCS)-3 expression. Serine kinases known to perform this phosphorylation include JNK, IKK, and PKCq. These pathways terminate at transcription factors such as AP-1 and c-Jun, which may also adversely impact on metabolic status. Both JNK and IKK are known players in a wide range of immune responses, and are also increased in liver and adipose in obese states [4, 49, 54–56]. In addition, in vivo and in vitro manipulation of these pathways, either by genetic modiﬁcation or pharmacological intervention, can improve (with inhibition of pathway) or worsen (with activation of pathway) the metabolic state of the tissue or organism [55]. JNK1 appears to be the more important of the JNK isoforms in the context of metabolic dysregulation and insulin resistance. PKCq, which is activated by diacylglycerols, links lipid inﬁltration and inﬂammatory signal transduction pathways through IKK, and so may also mediate IRS-1 serine phosphorylation-mediated insulin resistance via this mechanism. 18.6.2 Lipotoxicity and Insulin Resistance Affecting Liver Metabolism

It has been demonstrated that there is a clear link between abdominal fat expansion and the risk of developing the metabolic syndrome. Omental adipose tissue has greater uptake of fatty acids, higher sensitivity to catecholamine-stimulated lipolysis, and is less sensitive to inhibition of lipolysis by insulin than the subcutaneous adipose tissue. The liver directly receives the ﬁrst pass of both fatty acids (from higher basal and stimulated lipolysis in this adipose tissue) and cytokines produced by the visceral adipose tissue, which may be especially lipotoxic with hypertrophic adipose tissue. Expansion of the visceral adipose tissue with its enhanced lipid leakage facilitates the development of fatty liver. The speciﬁc reasons for why the mesenteric adipose tissue is more prolipotoxic to the liver are unclear, but a number of hypotheses have been postulated. These include (i) bacterial-induced inﬂammation from the intestine, (ii) more readily expanded adipocytes due to the relative hyperavailability/overload of the nutrients from the gut, and (iii) enhanced inﬂammatory proﬁle of the mesenteric fat due to adipocyte insulin resistance, altered cytokine proﬁle, macrophage inﬁltration, anoxia, and consequently apoptosis of adipocytes. 18.6.2.1 Hepatic Lipotoxicity A potential mechanism linking insulin resistance and hepatic steatosis involves dysregulation of IRS proteins and sterol regulatory element-binding protein (SREBP)-1 – the key transcription factor for de novo lipogenesis and triacylglycerol synthesis. SREBP-1 is activated both by saturated fatty acids and by proinﬂammatory signals, as well as in mice with insulin resistance [57]. In liver, the IRS proteins are linked to different aspects of metabolism. IRS-1 is primarily concerned with glucose homeostasis and control of glucose output, whereas IRS-2 is involved with lipid metabolism [58]. IRS-2 and SREBP-1 appear to be reciprocally regulated [59, 60]. Yet, IRS-2 expression is repressed in conditions of hyperinsulinemia, thereby allowing

18.8 New Analytical and Computational Methods

SREBP-1 action and, hence, de novo lipogenesis to occur. The ﬂux via de novo lipogenesis increases the intermediate malonyl-CoA. Since malonyl-CoA is a potent inhibitor of CPT-1 [61], the oxidative capacity of the liver becomes limited, facilitating hepatic lipid accumulation from de novo generated lipids and exogenously derived lipids from the adipose tissue.

18.7 Pancreatic b-Cell as a Target for Lipotoxicity

The transition from an insulin resistant to a diabetic state is marked by the failure of the insulin-producing b-cell to provide enough insulin to stimulate other tissues to contribute to glucose and fatty acid disposal. In metabolically healthy individuals, postprandial hyperglycemia is matched by a temporary hyperinsulinemia to enable peripheral substrate disposal. However, with time, as insulins effectiveness at stimulating glucose disposal decreases due to peripheral insulin resistance, insulin output increases until a point when the b-cells fail and insulin secretion no longer matches the requirements for substrate disposal. Fatty acids have been implicated in the development of the failing b-cell. Acute fatty acid treatment of b-cells actually stimulates insulin secretion, whereas chronic treatment inhibits insulin output. Treatment of isolated islets from humans and rodent models with palmitate prevents proliferation and triggers apoptosis [62, 63]. The combination of glucose and fatty acids, as occurring in a hyperglycemic, hyperlipidemic subject, may be additionally deleterious since glucose oxidation is preferred to b-oxidation and causes the accumulation of long-chain acyl-CoA units in the cell. It remains unclear whether fatty acids themselves or a speciﬁc metabolite are the main deleterious species. However, similar to the situation in muscle, it is likely that a combination of fatty acid derivatives including diacylglycerol or ceramide are likely to be involved [40]. In addition, deleterious contributions of fatty acids to mitochondrial function via enhanced uncoupling protein action, direct effects on the ion channels required for insulin secretion and on insulin synthesis and glucose metabolism have been proposed. It is also interesting to note that chronic high levels of glucose also trigger b-cell apoptosis, so it is likely that the combination of high circulating glucose and free fatty acids is especially deleterious for b-cells. As shown in Table 18.1, b-cell function is likely to be impaired by hypertrophic WAT adipokine output.

18.8 New Analytical and Computational Methods to Identify Lipotoxicity-Related Metabolic Networks

Given the impossibility of measuring the levels of all metabolites in a biological sample simultaneously with a single analytical platform, it was previously difﬁcult to identify a particular lipid species as being responsible for the metabolic abnormalities.

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The reason for this is that metabolites are chemically diverse and can cover a dynamic range of over 10 orders of magnitude in concentration, so a single extraction and detection method for all metabolites from biological matrices is unfeasible. Additionally, the complexity of biological samples may also affect the efﬁciency and reliability of detection, such as with ion suppression effects in mass spectrometrybased approaches. Multiple analytical platforms coupled to different extraction methods for speciﬁc groups of metabolites are commonly applied in parallel to cover broad ranges of metabolites. Analytical technologies based on gas chromatography coupled to mass spectrometry (MS), liquid chromatography/MS), capillary electrophoresis coupled to MS, as well as nuclear magnetic resonance have most commonly been applied. With the enhanced capabilities of modern MS instruments and interfaces, there has been rapid development of global lipid analytical methods, either using liquid chromatography/MS-based methods focusing on sensitive analyzes of total lipid extracts or on speciﬁc classes of metabolites, or direct MSn scanning driven by datadependent acquisition without the need for chromatographic separation. Due to the structural characteristics of lipids, their identiﬁcation from fragment mass spectra is generally easier than for other molecular components and todays typical global lipid proﬁling analyses allow identiﬁcation of several hundred lipid molecular species in parallel. Advances in analytical methods, along with improved data processing software solutions, demand development of comprehensive lipid libraries and bioinformatics solutions to allow system-level identiﬁcation, discovery, and subsequent study of lipids. Integrative studies combining multi-tissue lipidomic proﬁles with other levels of biological information such as gene expression and proteomics have been made possible due to such capabilities. However, reconstruction of lipid pathways from high-dimensional omics datasets remains a challenge. For example, current available pathway-level representations of lipids in databases such as KEGG is limited to pathway representation of generic lipid classes – including mainly the head group information and not including the fatty acid side-chain information. Therefore, these lipid databases lack the level of detail that is becoming available by modern lipidomics approaches. We have proposed a strategy to address this challenge that combines multivariate statistical analysis in order to extract the clusters of coregulated lipids, followed by instantiation of lipid pathways for individual lipids in the cluster based on their molecular structure and known biochemical reactions. Using our new methodology for lipid pathway reconstruction from lipidomics and transcriptomics data, we have recently reconstructed the ceramide synthesis pathways in fatty livers of obese mice. We have found that two ceramide synthesis pathways are upregulated in fatty livers: (i) de novo ceramide synthesis pathways due to increased ﬂux of fatty acids into the cells, and (ii) glucosylceramidase and galactosylceramidase pathways, leading to release of ceramide from the membrane glycosphingolipids. Reassuringly, in a recent study it was established that pharmacological inhibition of glucosylceramide synthase enhances insulin sensitivity in the obese mouse model.

References

18.9 Lessons from Lipotoxicity – Potential Antilipotoxic Therapeutic Strategies

While the best strategies to prevent lipotoxicity are to prevent or to cure obesity, it is clear that current therapeutic strategies are largely failing. Applying a more pragmatic approach, alternatives strategies are needed to prevent obesity-associated complications. As indicated above, there is a convincing link between excess or paucity of adipose tissue, deposition of lipids in non-adipose tissue organs, and the development of insulin resistance. In addition, imbalances in adipose expansion and in substrate uptake and disposal are likely to contribute to the pathology arising from excess circulating lipids. From the hypotheses generated here in this chapter, we propose a number of approaches to be considered for combating both lipotoxicity and insulin resistance: i. Modulation of adipose tissue plasticity to increase its lipid-buffering capacity by promotion of preadipocyte differentiation in an attempt to facilitate the development of hyperplastic instead of hypertrophic adipose tissue. An alternative for this is enhancement of adipose tissue metabolic capacity by stimulation of WAT towards a pro-oxidative brown adipose tissue phenotype. This has the advantage that is potentially a treatment for obesity by facilitating energy dissipation. ii. Repartitioning of nutrients between organs, by diverting fat away from organs with increased susceptibility to the deleterious effects of lipids and towards organs with increased capacity to deal with the excess of fat. Similarly, it may be conceivable to modulate the lipid enzymatic pathways to divert the excess of nutrients from the most toxic pathways and so generate less-reactive species. However, this approach demands acquiring complete knowledge about the organ-speciﬁc lipid metabolic networks. iii. Increase the oxidative capacity of peripheral organs to eliminate excess toxic lipids through either mitochondrial uncoupling and/or modulation of mitochondrial biogenesis. This strategy requires that elevation in oxidative capacity is matched by anappropriate substrateuptakecapability toensurethat substratesareavailable for use, but do not overwhelm the oxidative capacity of the organ. This strategy may partially recapitulate aspects related to the beneﬁcial effects of exercise. These ideas provide the framework for identifying druggable targets that can inﬂuence one or more of the aspects outlined above.

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19 Obesity and Cancer Andrew G. Renehan 19.1 Introduction

There is increasing body of epidemiological and clinical evidence implicating excess body weight both in increased risk of cancer incidence and cancer mortality. In this chapter, the evidence supporting these observations is summarized. The bulk of the chapter then lists a number of possible mechanisms underpinning the epidemiological associations, focusing on the three most studied – insulin and insulin-like growth factor, sex steroids, and adipokine systems. The chapter concludes in summarizing some alternative mechanisms and areas of new research.

19.2 Epidemiology 19.2.1 Excess Body Weight and Cancer Risk

Excess body weight, deﬁned as overweight (body mass index (BMI) 25–29.9 kg/m2) or obesity (BMI 30 kg/m2), has long been recognized as an important risk factor for cardiovascular disease and type 2 diabetes. It was not until 2002 that the International Agency for Research into Cancer [1] concluded that excess body weight is also associated with an elevated risk of developing of a number of cancers. More recently, the World Cancer Research Fund [2] reviewed a large body of literature and concluded that there was convincing evidence that body fatness (generally expressed as BMI) is associated with an increased risk of esophageal adenocarcinoma, and cancers of the pancreas, colorectal, postmenopausal breast, endometrium and kidney, and evidence of a probable association with risk of gallbladder cancer (Table 19.1).

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Table 19.1 Approximate estimates of association per incremental 5 kg/m2 BMI increase.

Colorectal cancer colon rectum Esophageal adenocarcinoma Gallbladder Pancreatic cancer Renal cancer Prostate cancer Breast cancer premenopausal postmenopausal Endometrial cancer Ovarian cancer

Men

Women

1.3 1.1 1.5 1.2 1.1 1.3 1.0

1.1 1.0 1.5 1.4 1.1 1.4

0.9 1.1 1.6 1.1

Estimates are approximations taken from large studies such as the Million Women Study [3] and meta-analyses reported in the World Cancer Research Fund review [2]. Direct comparisons were not possible as study selection criteria differed between cancer sites; hence, approximations are given.

Simultaneously, the Million Women Study [3] reported on a cohort of over 1 million women residing in the UK with a greater than 5 years of follow-up and listed the following cancers as at increased risk: endometrial cancer, adenocarcinoma of the esophagus, kidney cancer, leukemia, multiple myeloma, pancreatic cancer, nonHodgkins lymphoma, ovarian cancer, breast cancer in postmenopausal women, and colorectal cancer in premenopausal women. In many examples, associations were greater among never-smokers, suggesting interactions, in terms of risk, between smoking and BMI. Additionally, the effect of BMI on risk differed signiﬁcantly according to menopausal status (and, by extrapolation, hormonal inﬂuences) as follows: increased risk in premenopausal women for colorectal cancer and malignant melanoma, while increased risk in postmenopausal women for breast and endometrial cancers. A number of meta-analyses (albeit adopting different selection criteria) add to the evidence (complete references listed elsewhere [4]). Associations of increased risk with obesity or an incremental increase in BMI have been reported for colon (men and women) and rectal (men only), gallbladder, liver, pancreatic, ovarian, and prostate cancers, and leukemia, non-Hodgkins lymphoma, and multiple myeloma. 19.2.2 Excess Body Weight and Cancer Mortality

Given that BMI is consistently associated with cancer risk at several sites, it is not surprising that increased adiposity may have a negative effect on treatment outcome and ultimate survival. However, these data have only emerged in the past 5 years. There is now substantial evidence that obesity is a negative prognostic factor in breast

19.3 Biological Mechanisms

cancer [5]. Additionally, increased BMI has a negative impact on outcome for colon, endometrial, and ovarian cancers, and, paradoxically, increased BMI may be associated with improved outcome in renal cell carcinoma [4]. For prostate cancer, increased BMI is associated with a shorter time to biochemical progression and increased cancer-related mortality. These observations in human studies are supported by parallel observations in diet-induced obese animal models [6] and by the well recognized observation that energy restriction in animals is a potent inhibitor of progression of several tumor types [7]. Further evidence that obesity may be an unfavorable prognostic factor in patients diagnosed with cancer comes from cohort studies in which the relative risks associated with increasing BMI are substantially greater for mortality outcome than those predicted by incidence outcome [3, 8].

19.3 Biological Mechanisms 19.3.1 Candidate Mechanisms

Table 19.2 lists a number of possible mechanisms linking excess body weight and cancer risk. Among the biological mechanisms, there are three most-studied candidate systems; (i) insulin and insulin-like growth factors (IGFs), (ii) sex steroids, and (iii) adipokines. Insulin resistance is at the heart of many candidate systems and, as will be illustrated below, there are several examples where there are interactions between systems. With such diversity of obesity-related cancers, it is unlikely that there is a one system ﬁts all mechanism.

Table 19.2

Excess body weight and cancer – plausible biological mechanisms.

Biological mechanisms most studied systems insulin and IGFs sex steroids and sex steroid-binding globulin Adipokines (e.g., adiponectin and leptin) other mechanisms obesity-related inﬂammatory cytokines NF-kB system altered immune response oxidative stress peroxidation and renal cancer Mechanical mechanisms hypertension and renal cancer acid reﬂux and esophageal adenocarcinoma increased iodine uptake and thyroid cancer

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19.3.2 Insulin and IGFs 19.3.2.1 Insulin-Cancer Hypothesis For over a decade, it has been appreciated that there are similarities between risk factors for Westernized cancers and those for insulin resistance, suggesting that hyperinsulinemia may contribute to cancer development through the growth-promoting effect of elevated levels of insulin. In its simplest form, the insulin-cancer hypothesis postulates that prolonged hyperinsulinemia reduces IGF-binding protein (insulin-like growth factor-binding proteinIGFBP)-1 and -2 production (which normally bind IGF-I and inhibit its action), with resultant increases in the levels of free or bioactive IGF-I, and concomitant changes in the cellular environment favoring tumor development [9, 10]. A major limitation of the hypothesis is the lack of proof that links components of the IGF system in the circulation with cellular function. Additionally, and importantly, insulin resistance is not a simple direct model; instead, it interacts and regulates many systems that are relevant to tumorigenesis. Accordingly, we have used the terminology simple and complex insulincancer hypotheses (Figure 19.1) in previous publications [9]. 19.3.2.2 Insulin and C-Peptide Insulin is a two-chain peptide speciﬁcally secreted from the pancreatic b-cells. Insulin activation of the insulin receptor triggers intracellular signaling cascades in both the extracellular signal-regulated kinase (ERK) and phosphoinositide-3kinase (PI3K) pathways, and thus insulin signaling has the machinery to be mitogenic and antiapoptotic. Conventionally, however, it is felt that insulin is mitogenic only at supraphysiological levels and its main proliferative effects are probably mediated through IGF-I receptors [9]. Insulin analogs may be more mitogenic than insulin as they have a greater afﬁnity for the IGF-I receptor [11]. On the other hand, it is increasingly recognized that insulin has other biological properties (such as priming cells to the effects of other growth factors (including IGF-I) by inﬂuencing farnesylation of ras in vitro and in vivo, activation ERK and PI3K signal transduction pathways, and stimulation of b-catenin – an early signaling pathway in many cancers) that may be relevant to tumor development [4]. In cancer epidemiological studies, measurement of serum insulin is highly dependent on the state of fasting, assay characteristics, and genetic factors, and accordingly, surrogates of insulin secretion (e.g., C-peptide) or of insulin resistance (e.g., homeostasis model assessment (HOMA)) are used. Several epidemiological studies (Table 19.3) have independently linked high circulating levels of serum C-peptide with increased risk of postmenopausal breast cancer, colorectal cancer, and endometrial cancer, but inconsistent associations with premenopausal breast cancer, consistent with observations of cancers associated with obesity (a hyperinsulinemic state). Furthermore, insulin resistance is an adverse prognostic factor for breast, colorectal, and prostate cancer-related mortality (complete list of references listed elsewhere [4]).

19.3 Biological Mechanisms

Excess body weight

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Chronic sub-clinical inflammatory state • C-reactive protein↑

NF-κB system • IKKβ • PPARs

Metabolic syndrome

Insulin resistance Insulin-sex hormone axis • Peripheral aromatase activity↑ • Estrogen bio-availability↑ • SHBG production↓ • Ovarian androgen↑ • Progesterone↓

GH-GHBP axis • GH secretion↓ • Hepatic IGF-I and IGFBP-3 production↓ • Hepatic GH receptor↑ and GHBP↑ • Relative IGFI/IGFBP-3↑ Figure 19.1 Schematic representation of the complex insulin-cancer hypothesis. This figure demonstrates the complex interrelationships between obesity, insulin resistance, and pathways that may favor tumor development. Within these complexities, the simple insulin-cancer hypothesis is indicated

• Central obesity • Impaired glucose intolerance • Hypertension • Dyslipidemia

Hyperinsulinemia Type 2 diabetes mellitus

Insulin-IGF-I axis • IGFBP-1↓ • IGFBP-2↓ • Bio-availability IGF-I↑

Leptin↑ Adiponectin↓

Simple hypothesis by the dotted lined box and is only a small part of the overall emerging network of systems linking obesity with cancer. Abbreviations not defined in text: FFA, free fatty acids; GHBP, growth hormone-binding protein. ": increase in circulating concentrations; #: decrease in circulating concentrations.

As a clinical example, there is accumulating evidence that type 2 diabetes is associated, independent of obesity, with cancers of the colorectal, pancreas, kidney and endometrium, and variably with postmenopausal breast cancer. Associations are often strongest where the diagnosis of diabetes has been recent, consistent with the observation that type 2 diabetes is generally characterized by compensatory hyperinsulinemia in its early course. A further dimension is that, at least in the case of colorectal cancer, there is one study in which the use of therapeutic insulin increased the cancer association with type 2 diabetes [12].

Associations of serum insulin and C-peptide and various cancers.

Researchers (for complete references, see [4])

All breast cancer Mink et al., 2002 Premenopausal breast cancer Bruning et al., 1992 Del Giudice et al., 1998 Toniolo et al., 2000 Yang et al., 2001 Hirose et al., 2003 Postmenopausal breast cancer Bruning et al., 1992 Toniolo et al., 2000 Yang et al., 2001 Muti et al., 2002 Hirose et al., 2003 Keinan-Boker et al., 2003 Schairer et al., 2004 Colorectal cancer Schoen et al., 1999 Saydah et al., 2003 Kaaks et al., 2000 Ma et al., 2004 Wei et al., 2005

Country

Study design

Cases/controls

Serum insulin odds ratioa) (95% confidence intervals)

Serum C-peptide odds ratiod) (95% confidence intervals)

USA

cohort

187/7984

1.01 (0.55, 1.86)

—

Netherlands USA USA China Japan

case-control case-control nested case-control nested case-control case-control

79/129 99/99 172/486 45/45 88/79

— 2.83 (1.22, 6.58) — — 0.55 (0.22, 1.36)

6.8 (0.8, 60.6) — 0.76 (0.44, 1.31) 3.1 (0.7, 14.2) 0.85 (0.32, 2.23)

Netherlands USA China Italy Japan Netherlands USA

case-control nested case-control nested case-control nested case-control case-control nested case-control case-control

144/312 115/220 98/98 64/238 99/111 149/333 185/159

— — — 0.85 (0.36, 2.00) 2.43 (1.06, 5.58) — —

7.8 (2.2, 27.4) 1.24 (0.66, 2.34) 2.9 (1.1, 8.0) — 2.00 (0.89, 4.52) 1.3 (0.7, 2.7) 1.5 (0.7, 3.0)

USA USA USA USA USA

cohort case-control nested case-control nested case-control nested case-control

102/5747 men and women 173/346 men and women 102/200 women 176/294 men 182/350 women

1.2 (0.7, 2.1) 0.78 (0.45, 1.35) — — —

— — 2.92 (1.26, 6.75) 2.5 (1.2, 5.6) 1.17 (0.63, 2.20)

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Table 19.3

Prostate cancer Hsing et al., 2003 Chen et al., 2005 Endometrial cancer Weiderspass et al., 2003

Lukanova et al., 2004 Ovarian cancer Lukanova et al., 2003

China USA

case-control case-control

128/306 174/174

2.78 (1.63, 4.72)b) 0.89 (0.45, 1.76)

— —

Sweden

case-control

0.11 (0.03, 0.49)

—

2.44 (0.85, 7.07)

—

Sweden/USA/Italy

nested case-control

ever hormonal replacement therapy 140/109 never hormonal replacement therapy 120/187 166/315

Sweden/USA/Italy

nested case-control

132/261

—

4.40 (1.65, 11.7)

—

0.89 (0.44, 1.83)

Nested case-control: a case-control study design nested within a prospective cohort study. a) Uppermost versus lowermost category (e.g., upper quartile versus lowest quartile). Values taken for most adjusted model. b) HOMA insulin resistance ¼ fasting insulin fasting glucose.

19.3 Biological Mechanisms

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Table 19.4 IGF-I actions and tumor development.

Mitogenic and antiapoptotic Proangiogenic, induction of hypoxia-inducible factor-1-mediated vascular endothelial growth factor production Induction of tumor-related lymphangiogenesis Increased cell migration (in the majority of in vitro systems) Regulate cell differentiation, cell size, and organization of cellular cytoskeleton Potentiate the effects of other cell growth stimulants (e.g., estrogens) Regulation the location, stability, and transcriptional activity of b-catenin

19.3.2.3 IGFs The IGF system is a complex molecular network that includes two ligands (IGF-I and IGF-II), two receptors (IGF-I and IGF-II receptors), six high-afﬁnity binding proteins (IGFBP-1–6), and several binding protein proteases. IGF-I, IGF-II, and the IGFBPs occur in large quantities in the circulation and are readily measured. The IGF ligands bind IGFBP-3, the main circulating binding protein, to full saturation together with an acid-labile subunit to form a very stable ternary complex, which does not readily cross compartmental barriers. In terms of cancer risk, studies to date have focused mainly on circulating total IGF-I and IGFBP-3, both of which are growth hormonedependent, but are also inﬂuenced by age, gender, and nutritional status. IGF-I activates the IGF-I receptor to induce a variety of biological actions that may favor tumor growth (Table 19.4) – these are discussed in detail in a recent review [13]. Studies in the late 1990s suggested that circulating total IGF-I levels were positively associated, whereas total IGFBP-3 levels were negatively associated, with risk of common malignancies, including premenopausal breast, colorectal, prostate, and lung cancers. Subsequent studies and meta-analyses support a relationship between total IGF-I levels and risk of the ﬁrst three aforementioned malignancies, but reported relationships with IGFBP-3 have been inconsistent [14]. Acromegaly is an endocrine disorder characterized by sustained hypersecretion of growth hormone with concomitant elevation of IGF-I and is associated with a 2-fold increased risk of colorectal neoplasia [15]. Possible mechanisms underlying this increased risk include direct actions as a consequence of elevated levels of circulating growth hormone and IGF-I and/or other perturbations within the IGF system. 19.3.3 Sex Steroids 19.3.3.1 Estrogen and Breast Cancer For postmenopausal breast cancer, the increase in risk observed in obese women is generally explained by the higher rates of conversion of androgenic precursors to estradiol through increased aromatase enzyme activity in adipose tissue [16]. In support if this, there is abundant experimental evidence from in vitro and animal models that estrogens are mitogenic in normal and neoplastic mammary tissues [17]. At a molecular level, the estrogen receptor may act as a ligand-dependent

19.3 Biological Mechanisms

transcription factor that binds estrogen in a reversible fashion and with high afﬁnity to the hormone-binding domain. The possible role of estrogens as mutagens in the initiation of breast tumorigenesis has also been evaluated as estrogens may induce direct or indirect free radical-mediated DNA damage, genetic instability, and mutations in cells in culture and in vivo. However, even if estrogens can induce genetic damage, the data overall suggest that proliferative effects are likely to be the most important mechanism. Most established risk factors for breast cancer probably act through estrogenrelated pathways, supporting the unopposed estrogen hypothesis. Moreover, there is consistent and strong evidence that increased concentrations of circulating estrogens increase the risk of breast cancer in postmenopausal women. The Endogenous Hormones and Breast Cancer Collaborative Group (EHBCCG), in a pooled analysis of nine prospective studies, showed that postmenopausal breast cancer risk is increased (typically 2-fold for upper versus lowest quintiles) among women with higher concentrations of circulating sex steroids, including dehydroepiandrosterone (DHEA), its sulfate (DHEAS), D4-androstenedione, testosterone, estrone, and total estradiol, and decreased concentrations of sex hormone-binding globulin. Indeed, the EHBCCG analysis demonstrated that the association of BMI with postmenopausal breast cancer risk was almost entirely attributed to the increasing blood levels of estradiol with increasing BMI. The consistency of these associations is borne out in the large European Prospective Investigation into Cancer and Nutrition (EPIC) study (complete references listed elsewhere [16]). 19.3.3.2 Androgens and Breast Cancer Adiposity is inversely related to testosterone concentrations in men, but positively related in women. In addition, the EHBCCG and EPIC analyses demonstrate that elevated blood concentrations of androgens are associated with increased risk of breast cancer in both pre- and postmenopausal women, and thus androgens may be potential candidates linking obesity and breast cancer. However, there is conﬂicting experimental evidence regarding the role of androgens in breast cancer development. The conventional wisdom is that androgens inhibit breast growth [16]. However, the treatment of animals and cultured cells with androgens may have either inhibitory or stimulatory effects on the proliferation of mammary epithelia, and cancer cells; the mechanisms for these dual functions are unclear and are discussed elsewhere [18]. Experiments in rodents show that simultaneous treatment of androgen and estrogen synergizes for mammary gland carcinogenesis, and similar synergistic effects are observed for carcinogenesis of the uterine myometrium of female animals and for carcinogenesis of the prostate in males. It is postulated that concomitant elevation in both androgens and estrogens may confer a greater risk for tumorigenesis of the mammary gland, and probably other female reproductive tissues, than an elevation of each hormone alone [19]. 19.3.3.3 Sex Steroids and Endometrial Cancer For endometrial cancer, it is likely that there are several overlapping biological systems involved in the link between obesity and cancer predisposition. Increased

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estradiol levels not only increase endometrial cell proliferation and inhibit apoptosis, but also stimulate the local synthesis of IGF-I in endometrial tissue [10]. During the follicular phase of the menstrual cycle, when the ovaries produce estradiol but virtually no progesterone, the epithelial tissue and stromal ﬁbroblasts in the upper two-thirds of the endometrium proliferate. High proliferative rates continue until ovulation, when plasma estradiol levels reach a nadir and then decline rapidly during the luteal phase of the menstrual cycle, because of the increase in levels of progesterone, which antagonizes the proliferative actions of estrogen. In large part, the proliferative actions of estradiol on endometrial tissue are mediated by an increase in the local production (mostly in uterine tissue) of IGF-I. Progesterone diminishes estrogenic action in the endometrium by stimulating metabolism of estradiol and inducing the synthesis of IGFBP-1, which inhibits IGF-I action [20]. Similar to breast cancer risk, most established risk factors for endometrial cancer (e.g., early menarche, late menopause, obesity) probably act through pathways reﬂecting greater lifetime exposure to estrogens. Likewise, epidemiological studies have shown that higher levels of plasma estrone and estradiol are associated with increased endometrial cancer risk in postmenopausal women [20]. These relationships support an unopposed estrogen hypothesis for endometrial cancer, but ovarian hyperandrogenism may also be a central mechanism linking obesity to endometrial cancer risk. Furthermore, epidemiological studies report that elevated plasma androstenedione and testosterone concentrations predict for increased endometrial cancer risk in both pre- and postmenopausal women [20]. As a clinical example, polycystic ovary syndrome and obesity are associated with increased risk of endometrial cancer in pre- and postmenopausal women, respectively, and share mechanistic pathways that overlap between the estrogen, progesterone, androgen, and IGF systems [16]. 19.3.4 Adipokines 19.3.4.1 Overview Over the past decade, investigators have appreciated that adipose tissue is a highly active and large source of endocrine and metabolic activity. Polypeptide hormones derived from adipose tissue are known as adipokines. With regard to cancer development, the two most widely studied adipokines are leptin (and its soluble receptor) and adiponectin. 19.3.4.2 Leptin and Cancer in Humans Leptin, a 167-amino-acid product of the ob gene, is positively correlated with obesity and is intrinsically associated with insulin – insulin acts as a positive feedback (IGF-I is a negative regulator) on leptin gene expression, to signal suppression of appetite. Leptin-deﬁcient ob/ob mice overfeed, and rapidly become hyperinsulinemic and diabetic. Leptin binds to the ubiquitous Ob receptor and activates PI3K and Janus kinase–signal transducer and activator of transcription signaling pathways, critical for cell survival, proliferation, and differentiation. Leptin is also proangiogenic in

19.3 Biological Mechanisms

both in vivo and in vitro models. Furthermore, in vivo experiments show that calorie restriction signiﬁcantly decreases tumor growth while concomitantly decreasing circulating leptin levels in rat colon cancer models; however, unexpectedly, leptin appears not to promote the growth of colon cancer xenografts in nude mice or intestinal tumorigenesis in the ApcMin/ þ model. Readers are referred elsewhere for more details [21]. Epidemiological studies relating serum leptin with prostate cancer risk have reported inconsistent ﬁndings – positive in two studies, no association in one, and no associations with breast cancer. However, two prospective studies of colorectal cancer risk demonstrated signiﬁcant associations. Stattin et al. [22] measured leptin in prediagnostic plasma from subjects in a Swedish cohort and demonstrated increased colorectal cancer risk among men, but not women. The same investigators subsequently showed in a larger study of men only an approximate 3-fold increase in colon cancer risk for the uppermost leptin quartile compared with the lowermost. Furthermore, a recently published colonoscopy study in men has shown that those in the highest tertile of leptin concentrations had a 3-fold increased adenoma risk compared with those in the lowest tertile, but there were no associations between leptin concentrations and adenoma risk in women (complete references listed elsewhere [4]). 19.3.4.3 Adiponectin and Cancer Risk in Humans Adiponectin, also termed Acrp30 (adipocyte complement-related protein of 30 kDa), ADIPOQ, apM1 (adipose most abundant gene transcript-1), or GBP28 (gelatinbinding protein of 28 kDa), a 247-amino-acid peptide, is the most abundant adipokine, secreted mainly from visceral adipose tissue, and is inversely related to BMI. It is an important insulin-sensitizing agent, as adiponectin-deﬁcient mice are both insulin-resistant and diabetes-prone. Of importance to tumor development, adiponectin is a negative regulator of angiogenesis (an essential component of tumor growth and metastasis) through the induction of apoptosis in vascular endothelial cells and inhibition of cell migration. In mouse models, adiponectin is associated with inhibition of primary tumor growth (for review, see [21]). Two medium-sized case-control studies demonstrated signiﬁcant inverse associations between circulating concentrations of adiponectin and endometrial cancer – effects that appear strongest in younger women [4]. A further two case-control studies concurrently reported links between low adiponectin concentrations in the circulation and breast cancer – one study demonstrating signiﬁcant associations for pre- and postmenopausal cases, the other for postmenopausal women only. A small study of men with prostate cancer reported an inverse association with plasma adiponectin levels and prostate cancer – a negative association that was strengthened in the presence of advanced disease. For colorectal cancer, data from the Health Professionals Follow-up Study demonstrated that men with low plasma adiponectin levels had a higher risk of colorectal cancer than men with higher levels [23], but this has not been repeated in analyses from a Swedish cohort [24]. Inconsistencies in associations between genders and across populations may be partly explained by the wellrecognized differences in the relationships of circulating adiponectin with BMI by gender (Figure 19.2).

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men women

Men: r = -0.27, p < 0.001 Women: r = -0.44, p < 0.001

30

20

10

0 Adiponectin (mg/l)

20

25

30

35

40

BMI (kg/m2)

Figure 19.2 Relationship of BMI to circulating adiponectin by gender. Data from healthy volunteers aged 55–64 years on fasted samples (146 men: 79 women). There was a significant

difference (P < 0.001) between men and women in a regression model of adiponectin concentrations, BMI, and gender. r, Pearsons correlation coefficient.

19.3.4.4 Adipokines, Animal Models, and Cancer Risk While human data suggest that the link between obesity and cancer is driven by white adipose tissue (the source of adipokines), recent studies in fatless A-Zip/F1 mice, which have undetectable adipokine levels but display accelerated tumor formation, suggest that adipokines are not required for enhanced tumor development. The A-Zip/F1 mice are also diabetic and display elevated circulating levels of other factors frequently associated with obesity (insulin, IGF-I, and proinﬂammatory cytokines) and activation of several signaling pathways associated with carcinogenesis. In view of this observation, it is postulated that the pathways associated with insulin resistance and inﬂammation, rather than adipocyte-derived factors, may represent the key mechanisms underlying the obesity–cancer link [25].

19.4 Other Biological Candidates 19.4.1 Obesity-Related Inflammatory Markers

The concentrations of several circulating cytokines are increased in obesity, including interleukin (IL)-6, tumor necrosis factor (TNF)-a, soluble TNFreceptors, and C-reactive protein (CRP) – creating a subclinical inﬂammatory state. One prospective study has

19.5 Mechanical Mechanisms

shown that plasma CRP concentrations are elevated among both men and women who subsequently develop colorectal cancer, but a second study failed to substantiate this ﬁnding in a cohort limited to women (complete references listed elsewhere [4]). 19.4.2 Nuclear Factor-kB System

Conditions that promote insulin resistance include inﬂammation and cytokine production [26]. One possible mechanism through which these factors elicit their effects is activation of IkB kinase (IKK)-b, an upstream activator of nuclear factor-kB (NF-kB), and it is therefore conceivable that mitogenic and antiapoptotic effects of insulin are mediated through this pathway [27]. This hypothesis is supported by work demonstrating that high doses of salicylates, which inhibit IKK-b activation, reverse hyperglycemia, hyperinsulinemia, and dyslipidemia in obese rodents through sensitizing insulin signaling. Peroxisome proliferator-activated receptors (PPARs) are members of a superfamily of nuclear hormone receptors, which antagonize the activities of NF-kB, and may be important in tumor development. Pioglitazone, a PPAR agonist, which is antiproliferative, proapoptotic, and anti-inﬂammatory, has recently been shown to reduce macrovascular complications in type 2 diabetes [28], but failed to demonstrate a protective effect on cancer risk. 19.4.3 Oxidative Stresses

There is increasing evidence that obesity, even in the absence of smoking, diabetes mellitus, hyperlipidemia, and renal or liver diseases, may decrease the activities of bodys protective antioxidants and enhance the systemic oxidative stress. This may be particularly relevant to obesity-related renal cancer development [29]. By contrast, available evidence implicates increased lipid peroxidation in the anticarcinogenic effect of suspected protective factors for breast cancer, including soy, marine n-3 fatty acids, green tea, isothiocyanates, and vitamin D and calcium. In the genetically obese ob/ob mouse, chronic exposure to ultraviolet radiation results in greater oxidative stress in the skin of obese mice in terms of higher levels of hydrogen peroxide and nitric oxide production, photo-oxidative damage of lipids and proteins, and greater depletion of antioxidant defense enzymes, like glutathione, glutathione peroxidase, and catalase [30]. In turn, these ultraviolet-induced oxidative stresses may mediate activation of mitogen-activated protein kinase and NF-kB signaling pathways, favoring tumor growth. These novel observations may be important in the link between obesity and malignant melanoma.

19.5 Mechanical Mechanisms

Hypertension is a well-recognized risk factor for the development of renal cell carcinoma. While obesity is a risk factor for hypertension, there are several studies

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demonstrating that increased BMI is associated with increased risk of kidney cancer independent of hypertension. Additionally, obesity may be independently associated with increased intrarenal tubular pressures. Chronic acid reﬂux may be relevant for the development of esophageal adenocarcinoma through the formation of squamous metaplasia and (Barretts) dysplasia. However, in a study that adjusted for acid reﬂux, the association of BMI and increased risk of esophageal adenocarcinoma remained [31]. Obesity is associated with increased uptake of iodine by the thyroid gland and this may be relevant to the increased risk of thyroid carcinoma observed in obesity [32].

19.6 New Research Areas

New opportunities to explore the mechanisms underpinning the link between obesity and cancer risk have been made possible through microarray analyzes and more detailed analyses of genes that are up- and downregulated. It appears that obesity may modulate the expression of genes that also are relevant for tumorigenesis. Work from the laboratory of Henry Thompson (Colorado) demonstrates that dietary energy restriction is associated with an upregulated pattern of expression in genes involved in glycolysis, whereas gluconeogenesis was suppressed, and that these changes differ between normal mammary tissue and breast adenocarcinoma [33]. A further example is the use of fatty acid synthetase inhibition. Two such compounds, C75 and C247, have been shown to delayed mammary tumor development in the transgenic neu-N mouse model [34]. These new ﬁndings provide a rationale for the combination of approaches – from hormonal to basic gene expression – in the prevention of many common cancers.

References 1 IARC (2002) IARC Handbooks of Cancer

Prevention 6: Weight Control and Physical Activity (eds H. Vainio and F. Bianchini), IARC Press, Lyon. 2 WCRF (2007) Food, Nutrition, Physical Activity, and the Prevention of Cancer: A Global Perspective, 2nd edn, American Institute for Cancer Research, Washington, DC. 3 Reeves, G.K., Pirie, K., Beral, V., Green, J., Spencer, E., and Bull, D. (2007) Cancer incidence and mortality in relation to body mass index in the Million Women Study: cohort study. Br. Med. J., 335, 1134–1140. 4 Renehan, A.G., Roberts, D.L., and Dive, C. (2008) Obesity and cancer:

pathophysiological and biological mechanisms. Arch. Physiol. Biochem., 114, 71–83. 5 Carmichael, A.R. (2006) Obesity and prognosis of breast cancer. Obes. Rev., 7, 333–340. 6 Yakar, S., Nunez, N.P., Pennisi, P., Brodt, P., Sun, H., Fallavollita, L., Zhao, H., Scavo, L., Novosyadlyy, R., Kurshan, N., Stannard, B., East-Palmer, J., Smith, N.C., Perkins, S.N., Fuchs-Young, R., Barrett, J.C., Hursting, S.D., and LeRoith, D. (2006) Increased tumor growth in mice with diet-induced obesity: impact of ovarian hormones. Endocrinology, 147, 5826–5834.

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and carcinogenesis of the mammary gland. J. Steroid. Biochem. Mol. Biol., 80, 175–189. Roberts, D.L., Dive, G., Renehan, A.G. (2009) Biological mechanisms linking obesity and cancer risk: new perspectives. Annu. Rev. Med., (epub). Kaaks, R., Lukanova, A., and Kurzer, M.S. (2002) Obesity, endogenous hormones, and endometrial cancer risk: a synthetic review. Cancer Epidemiol. Biomarkers Prev., 11, 1531–1543. Rose, D.P., Komninou, D., and Stephenson, G.D. (2004) Obesity, adipocytokines, and insulin resistance in breast cancer. Obes. Rev., 5, 153–165. Stattin, P., Palmqvist, R., Soderberg, S., Biessy, C., Ardnor, B., Hallmans, G., Kaaks, R., and Olsson, T. (2003) Plasma leptin and colorectal cancer risk: a prospective study in Northern Sweden. Oncol. Rep., 10, 2015–2021. Wei, E.K., Giovannucci, E., Fuchs, C.S., Willett, W.C., and Mantzoros, C.S. (2005) Low plasma adiponectin levels and risk of colorectal cancer in men: a prospective study. J. Natl. Cancer Inst., 97, 1688–1694. Lukanova, A., Soderberg, S., Kaaks, R., Jellum, E., and Stattin, P. (2006) Serum adiponectin is not associated with risk of colorectal cancer. Cancer Epidemiol. Biomarkers Prev., 15, 401–402. Hursting, S.D., Nunez, N.P., Varticovski, L., and Vinson, C. (2007) The obesity–cancer link: lessons learned from a fatless mouse. Cancer Res., 67, 2391–2393. Bruce, W.R., Wolever, T.M., and Giacca, A. (2000) Mechanisms linking diet and colorectal cancer: the possible role of insulin resistance. Nutr. Cancer, 37, 19–26. Komninou, D., Ayonote, A., Richie, J.P. Jr., and Rigas, B. (2003) Insulin resistance and its contribution to colon carcinogenesis. Exp. Biol. Med., 228, 396–405. Dormandy, J.A., Charbonnel, B., Eckland, D.J., Erdmann, E., Massi-Benedetti, M., Moules, I.K., Skene, A.M., Tan, M.H., Lefebvre, P.J., Murray, G.D., Standl, E., Wilcox, R.G., Wilhelmsen, L., Betteridge, J., Birkeland, K., Golay, A., Heine, R.J., Koranyi, L., Laakso, M., Mokan, M., Norkus, A., Pirags, V., Podar, T., Scheen,

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A., Scherbaum, W., Schernthaner, G., Schmitz, O., Skrha, J., Smith, U., and Taton, J. (2005) Secondary prevention of macrovascular events in patients with type 2 diabetes in the PROactive Study (PROspective pioglitAzone Clinical Trial In macroVascular Events): a randomised controlled trial. Lancet, 366, 1279–1289. 29 Gago-Dominguez, M., Castelao, J.E., Yuan, J.M., Ross, R.K., and Yu, M.C. (2002) Lipid peroxidation: a novel and unifying concept of the etiology of renal cell carcinoma (United States). Cancer Causes Control, 13, 287–293. 30 Katiyar, S.K. and Meeran, S.M. (2007) Obesity increases the risk of UV radiationinduced oxidative stress and activation of MAPK and NF-kappaB signalling. Free Radic. Biol. Med., 42, 299–310. 31 Lindblad, M., Rodriguez, L.A., and Lagergren, J. (2005) Body mass, tobacco and alcohol and risk of esophageal, gastric cardia, and gastric non-cardia

adenocarcinoma among men and women in a nested case-control study. Cancer Causes Control, 16, 285–294. 32 Dal Maso, L., La Vecchia, C., Franceschi, S., Preston-Martin, S., Ron, E., Levi, F., Mack, W., Mark, S.D., McTiernan, A., Kolonel, L., Mabuchi, K., Jin, F., Wingren, G., Galanti, M.R., Hallquist, A., Glattre, E., Lund, E., Linos, D., and Negri, E. (2000) A pooled analysis of thyroid cancer studies. V. Anthropometric factors. Cancer Causes Control, 11, 137–144. 33 Zhu, Z., Jiang, W., McGinley, J.N., Price, J.M., Gao, B., and Thompson, H.J. (2007) Effects of dietary energy restriction on gene regulation in mammary epithelial cells. Cancer Res., 67, 12018–12025. 34 Alli, P.M., Pinn, M.L., Jaffee, E.M., McFadden, J.M., and Kuhajda, F.P. (2005) Fatty acid synthase inhibitors are chemopreventive for mammary cancer in neu-N transgenic mice. Oncogene, 24, 39–46.

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20 Overview of Acquired and Genetic Lipodystrophies Tisha Joy and Robert A. Hegele 20.1 Introduction

Lipodystrophies are a clinically heterogeneous group of disorders characterized by adipose tissue loss either in localized or generalized regions of the body [1]. These disorders are often accompanied by metabolic abnormalities such as insulin resistance, glucose intolerance, lipid proﬁle disturbances (hypertriglyceridemia and low high-density lipoprotein (HDL) cholesterol), hypertension, polycystic ovary syndrome (PCOS), hepatic steatosis as well as an increased risk of premature coronary artery disease. Thus, these disorders represent an extreme form of the more common metabolic syndrome, and insights into the genetic and clinical characteristics of lipodystrophies may prove useful for the prevention and management of metabolic syndrome [2]. Lipodystrophies have been traditionally classiﬁed into two broad categories – acquired and genetic. Lipodystrophies may also be a component of rare inherited multisystem syndromes such as SHORT syndrome and the progeroid syndromes [1] (Table 20.1). The metabolic severity of each type of lipodystrophy is variable and often correlates with clinical severity, with changes in adipose tissue stores being just one of many possible instigators of altered metabolic ﬁndings among these patients [2, 3]. However, even within this somewhat arbitrary categorization of acquired and genetic lipodystrophies, subtle differences and similarities between lipodystrophies can be delineated based on clinical and laboratory features, including information from recent magnetic resonance imaging (MRI) characterization of these disorders (Table 20.2). Furthermore, genetic heterogeneity exists among molecularly characterized lipodystrophies, with their clinical phenotypes varying based on the causative factor. We will discuss the clinical and molecular features of each disorder, and important differences and similarities between the groups as determined using clinical, genetic, and laboratory features.

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Table 20.1 Lipodystrophy classification.

Congenital congenital generalized lipodystrophy (Berardinelli–Seip) CGL1 (due to mutations in AGPAT2) CGL2 (due to mutations in BSCL2) familial partial lipodystrophy FPLD1 (Kobberling) FPLD2 (Dunnigan, due to mutations in LMNA) FPLD3 (due to mutations in PPARG) Acquired acquired generalized lipodystrophy acquired partial lipodystrophy (Barraquer–Simons; some cases are associated with mutations in LMNB2) HIV-related lipodystrophy Lipodystrophy as part of another syndrome mandibuloacral dysplasia SHORT syndrome neonatal progeroid syndrome Hutchinson–Gilford progeria syndrome Werner syndrome

20.2 Congenital Lipodystrophies 20.2.1 Congenital Generalized Lipodystrophy (Berardinelli–Seip Syndrome) 20.2.1.1 Clinical Features Congenital generalized lipodystrophy (CGL) was ﬁrst described in the 1950s by Berardinelli from Brazil and Seip from Norway [4, 5]. This disorder is inherited in an autosomal recessive manner and is characterized by generalized near-absence of adipose tissue. Affected individuals are usually recognized soon after birth due to the generalized muscular phenotype. MRI quantiﬁcation of regional adipose distribution of individuals affected with CGL conﬁrms decreased stores of supraclavicular, gluteal, mid-thigh, and mid-calf fat, but also reveals variable stores of visceral fat, when compared to individuals with familial partial lipodystrophy (FPLD), acquired partial lipodystrophy (APL), and HIV-related lipodystrophy [6]. The childhood years in affected individuals are distinguished by the presence of voracious appetite, accelerated linear growth, advanced bone age, and typically, marked acanthosis nigricans [7, 8]. Acromegaloid features, including enlarged hands, feet, and jaw, are often present. Other associated features include umbilical hernia, hepatomegaly secondary to hepatic steatosis, and focal lytic lesions of the appendicular bones [7–9]. Cardiomyopathy and intellectual impairment may occur, but these seem to be related to the genotype [10].

Table 20.2

Features in some forms of lipodystrophy that have a genetic basis. Congenital generalized lipodystrophy AGPAT2 (CGL1)

Demographics age at onset gender preference ethnicity preference

limb fat loss

BSCL2 (CGL2)

LMNA Diabetic

Nondiabetic

Acquired partial lipodystrophy

PPARG

puberty to adulthood female N. European Hispanic

typically < 20 years

normal

normal to overweight

normal

0 0

0 0

0 to þ þ 0

þþþþ

þþþþ

þ þ þ þ (distal predominantly)

0 to þ þ þ variable loss of palm fat; no loss of retroorbital fat; normal or increased sole fat þ þ to þ þ þ (but, normal or increased fat in lower extremities)

shortly after birth

shortly after birth

puberty

puberty

female African-American

female Caucasian Chinese Lebanese

female N. European

female N. European

normal– underweight

normal– underweight

normal

þþþþ 0

þþþþ þþþþ

þþþþ

þþþþ

female Caucasian

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(Continued)

20.2 Congenital Lipodystrophies

Fat distribution body mass index (based on World Health Organization criteria) facial fat loss mechanical fat loss (retro-orbital/palm/ sole)

Familial partial lipodystrophy

Congenital generalized lipodystrophy AGPAT2 (CGL1)

trunk fat gluteal fat bone marrow fat hepatic steatosis Clinical features diabetes age at onset of diabetes hypertension acanthosis nigricans hirsutism sexual development/ function

early coronary artery disease Metabolic parameters

Familial partial lipodystrophy

BSCL2 (CGL2)

LMNA Diabetic

Nondiabetic

Acquired partial lipodystrophy

PPARG

#### #### #### þþþ

#### #### #### þþþ

"""" #### unknown 0 to þ þ

"""" #### unknown 0 to þ þ

"""" 0 to ## unknown þ þ to þ þ þ

## to ### 0 to "" 0 rare

very common adolescence

very common <10 years

absent NA

rare present can be present external genitalia pseudohypertrophy often present

typically present adolescence to adulthood present typically present common menstrual anomalies or PCOS often present

uncommon adulthood (<45 years)

rare present can be present no changes

unknown

unknown

present 35–45 years present present rare external genital pseudohypertrophy can be present PCOS rare 50%

typically rare

rare

present present rare Polycystic ovary syndrom (PCOS) rare

rare

can be present rare uncommon PCOS rare

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Table 20.2 (Continued)

increased fasting insulin increased triglyceride decreased HDL increased free fatty acids leptin adiponectin elevated C-reactive protein other features

þþþ þþþ 0 to þ þ a) unknown #### ####a) unknown

not associated with mental retardation

can be associated with mental retardation

cardiomyopathy can be present

higher risk of cardiomyopathy than CGL1 bone cysts may be present

bone cysts may be present a)

Findings are for CGL as a group, irrespective of genetic mutation.

þþ þþþ 0 to þ 0 to þ ## ### þ

þ þþ 0 to þ 0 to þ ## ### þþ

þþþþ þ þ to þ þ þ þ 0 to þ þ 0 to þ 0 to #### #### 0 to þ

0 to þ þ 0 to þ þ 0 0 # ## unknown Can be associated with: low C3 complement; autoimmune disorders; membranoproliferative glomerulonephritis

20.2 Congenital Lipodystrophies

þþþþ þþþþ 0 to þ þ a) unknown ### ####a) unknown

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Speciﬁc endocrine complications include fasting hyperglycemia, diabetes (often with signiﬁcant insulin resistance), hypertriglyceridemia (sometimes resulting in pancreatitis), and markedly low levels of adiponectin and leptin [11]. As well, among women, hirsutism, PCOS, and menstrual irregularities can occur, while reproductive function in men has been reported to be normal [12]. 20.2.1.2 Molecular Genetics There are two main molecular forms of CGL – type 1 caused by mutations in the AGPAT2 gene (Mendelian Inheritance in Man (MIM) 608594) and type 2 caused by mutations in the BSCL2 gene (MIM 606158). However, not all patients affected with CGL have mutations in one of these two genes, indicating that additional loci or pathways are responsible. AGPAT2, which is located on chromosome 9q34, encodes 1-acylglycerol-3-phosphate O-acyltransferase 2, also called LPA acyltransferase-b or 1-acyl-sn-glycerol-3phosphate acetyltransferase (EC 2.3.1.51) [13, 14], which is important for catalyzing reactions involved in the metabolism of lysophosphatidic acid, eventually leading to synthesis of triacylglycerol. It has also been linked to increased transcription and synthesis of cytokines including interleukin (IL)-6 and tumor necrosis factor (TNF)-a [15]. CGL1 caused by AGPAT2 often results from nonsense or aberrant splicing mutations with no obvious correlation demonstrated between mutation severity and phenotype severity. The BSCL2 gene in the CGL2 locus is located on chromosome 11q13 [16]. BSCL2 encodes the protein seipin – a 398-amino-acid integral membrane protein localized to the endoplasmic reticulum of eukaryotic cells, and expressed mainly in the brain and testes [16, 17]. The function of this protein remains to be determined. To date, more than 12 mutations in BSCL2 have been identiﬁed, and, similar to AGPAT2, mutations in the BSCL2 gene are typically of the nonsense or aberrant splicing variety with no obvious correlation demonstrated between mutation severity and phenotype severity. CGL2 appears to be a more severe phenotype than CGL1, with more extensive fat loss and biochemical changes, more severe cardiomyopathy and intellectual impairment, earlier diabetes onset, and possibly earlier mortality. 20.2.2 Familial Partial Lipodystrophy 20.2.2.1 Clinical Features FPLD, originally reported in the 1970s, is an autosomal dominant condition, subdivided into three varieties – FPLD1 (or Kobberling type; MIM 608600), FPLD2 (or Dunnigan type; MIM 151660) caused by LMNA mutations, and FPLD3 (MIM 604367) caused by PPARG mutations [18–20]. The most common type is FPLD2. FPLD consists of progressive and gradual subcutaneous adipose tissue loss from the extremities commencing in puberty (although the onset of FPLD3 may be later into adulthood). Thus, during infancy and childhood, affected individuals cannot be easily distinguished from unaffected individuals. The loss of adipose tissue from the

20.2 Congenital Lipodystrophies

extremities is accompanied by a variable degree of adipose tissue loss in the trunk and chest. FPLD1 subjects demonstrate normal or increased fat deposition in the face, neck, and trunk. FPLD2 subjects demonstrate decreased fat deposition in the trunk, and increased deposition in the neck and labia. FPLD3 subjects demonstrate decreased to absent facial fat. In addition, increased fat deposition within the muscles and within the liver can occur [21–24]. MRI quantiﬁcation of adipose stores among the different types of FPLD have revealed signiﬁcant disparities in abdominal adipose partitioning (measured at the level of L4) with FPLD2 subjects demonstrating more severe decreases in subcutaneous fat and increases in visceral fat compared to FPLD3 subjects, who demonstrate no decreases in subcutaneous and no increases in visceral fat. Further characterization by MRI has revealed that FPLD2 subjects have greater depletion of subcutaneous adipose tissue in the gluteal, thigh, and calf regions compared to FPLD3 subjects. Thus, MRI may be clinically useful in distinguishing FPLD3 from the more common FPLD2 since FPLD3 is associated with less signiﬁcant adipose tissue alterations [6]. However, FPLD3 subjects often demonstrate more severe clinical and endocrine abnormalities [24]. These endocrine manifestations for FPLD include hypertriglyceridemia, low HDL cholesterol levels, diabetes, acanthosis nigricans, and, among women, hirsutism, PCOS, and menstrual irregularities [25]. The risk of development of diabetes has been shown to be higher among women, particularly if multiparous and affected by excessive central adipose deposition, compared to men [26]. Leptin and adiponectin levels tend to be low, but higher than CGL and lower than APL [11]. Premature coronary artery disease is more common among diabetic individuals with LMNA mutations [27, 28]. 20.2.2.2 Molecular Genetics The molecular cause of FPLD1 remains unknown. Meanwhile, FPLD2 is caused by heterozygous mutations in the LMNA gene encoding nuclear lamin A/C (MIM 150330) [29]. Lamins, part of the intermediate ﬁlament family of proteins, aid in the structural integrity of the nuclear envelope, transcriptional regulation, nuclear pore functioning, and heterochromatin organization. Mutations in the genes encoding lamins can result in a variety of disorders termed laminopathies. Over 180 mutations have been associated with 13 such laminopathies, including FPLD2, Hutchinson–Gilford progeria syndrome (HGPS), Werner syndrome, Emery–Dreifuss muscular dystrophy, limb-girdle muscular dystrophy type 1B, dilated cardiomyopathy type 1A, and Charcot–Marie–Tooth, and various overlapping syndromes [30, 31]. The majority of causative LMNA mutations for FPLD2 are missense, with only one splicing mutation described thus far. These LMNA mutations are typically downstream of the nuclear localization sequence (NLS), which divides the lamin A protein into the structural rod domain on the N-terminal side and the DNA-binding domain on the C-terminal side, implying that a possible mechanism for FPLD2 may involve altered interactions of transcription factors or other DNA-binding molecules [30]. Recently, two FPLD2 mutations – LMNA D230N and R399C – have been found

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upstream of the lamin A NLS [32], indicating that alterations within the secondary and/or tertiary structure of lamin A may also be important for the development of FPLD2 [30]. Deletions or duplications of LMNA in FPLD2 patients have not yet been found (Hegele, unpublished observations). Despite these ﬁndings, the exact mechanisms by which LMNA mutations cause lipodystrophy remain to be elucidated. Interestingly, a recent freeze of National Center for Biotechnology Information AceView, which annotates genes by aligning cDNA and expressed sequence tags, has shown there is greater transcriptional variability in LMNA than previously thought, with perhaps more than 40 exons and more than 10 distinct mRNA transcripts present (http://www.ncbi.nlm.nih.gov/IEB/Research/Acembly). This information may thereby aid in identifying novel mutation-bearing regions for FPLD2 as well as other laminopathies. FPLD3 (MIM 604367) results from heterozygous mutations in the PPARG gene encoding peroxisome proliferator-activated receptor (PPAR)-c (MIM 601487) [33–41]. PPAR-c is a ligand-activated nuclear receptor that functions as a transcription factor. It is highly expressed in adipose tissue and important in mediating adipocyte differentiation, adipokine release, improvement in insulin sensitivity, and inhibition of expression of proinﬂammatory factors [42]. The proposed mechanisms for mutated PPAR-c in the pathogenesis of FPLD3 include (i) a dominant-negative mechanism, in which the mutant receptor competes with the wild-type for DNA binding, and (ii) a haploinsufﬁciency mechanism, in which the function of the PPAR-c receptor is adversely affected by a 50% reduction in gene expression. Careful cellular assays indicate that seven PPAR-c mutations (C114R, C131Y, C162W, FS315X, R357X, P467L, and V290M) act via a dominantnegative mechanism [34, 41], and six (–14A>G, F388L, E138fsDAATG, Y355X, R194W, and R425C) via haploinsufﬁciency [35–40]. For the dominant-negative PPARG mutations, receptor mutants were shown to lack DNA binding and transcriptional activity, but still translocated to the nucleus for interaction with PPAR-c coactivators while inhibiting coexpressed wild-type receptor, with attenuated expression of PPAR-c target genes, suggesting that the mutants restricted wild-type PPAR-c action via a non-DNA-binding, transcriptional interference mechanism involving the sequestration of functionally limiting coactivators [34]. Another proposed mechanism for certain dominantnegative mutant PPAR-c receptors was reduced promoter turnover rate with the mutant eventually out-competing the wild-type receptor for promoter binding sites [39]. However, the exact mechanisms by which mutant PPAR-c receptors lead ultimately to the expression of a lipodystrophy phenotype in FPLD3 remain to be fully elucidated. About 50% of FPLD patients do not carry a mutation in either LMNA or PPARG genes as determined by genomic DNA sequence analysis of known coding regions (Hegele, unpublished observations). The reasons for this may include (i) presence of mutation types not detected by DNA sequence analysis, such as copy number variations, (ii) genetic heterogeneity with new causative genes yet to be identiﬁed, and (iii) the presence of mutations in unrecognized functionally important sequences of LMNA or PPARG.

20.3 Acquired Lipodystrophies with a Possible Genetic Component

20.3 Acquired Lipodystrophies with a Possible Genetic Component 20.3.1 Acquired Generalized Lipodystrophy 20.3.1.1 Clinical Features Acquired generalized lipodystrophy (AGL) is typically recognized in the school-age and teenage years, with progressive loss of adipose tissue, typically affecting the face and extremities with varying changes in intra-abdominal fat. Although retro-orbital and intramarrow fat are preserved, affected patients may or may not have loss of adipose tissue in the palms or soles. Females tend to be more affected than males at a ratio of 3 : 1. During childhood, affected individuals may be noted to have a voracious appetite, acanthosis nigricans, and hepatic steatosis. Endocrine abnormalities include low levels of leptin and adiponectin, hyperinsulinemia, diabetes, hypertriglyceridemia, and low HDL levels. Unlike affected males, females demonstrated changes in reproductive parameters, including menstrual irregularities and PCOS. AGL has been subdivided into three groups – AGL associated with autoimmune disorders, AGL associated with panniculitis, and idiopathic AGL. Based on this classiﬁcation, Misra and Garg found the prevalence of diabetes and hypertriglyceridemia to be highest in both the autoimmune and idiopathic groups compared to the panniculitis group (approximately 88 versus 44% for diabetes and approximately 90 versus 59% for hypertriglyceridemia) [43]. 20.3.1.2 Molecular Genetics Using candidate gene sequencing of known lipodystrophy genes (AGPAT2, BSCL2, LMNA, PPARG, and LMNB2) and also candidate genes encoding nuclear envelope proteins (LBR, LMNB1, and emerin), we have to date found no putative causative or associated mutations (Hegele, unpublished observations). 20.3.2 Acquired Partial Lipodystrophy (Barraquer–Simons Syndrome) 20.3.2.1 Clinical Features APL (MIM 608709) was initially reported in the 1880s by Mitchell, followed by two reports in the early 1900s [44–46]. Like the other lipodystrophies described above, there is a female preponderance of ascertained cases at a ratio of approximately 4 : 1. Affected individuals develop adipose tissue loss affecting primarily the face, neck, arms, thorax, and upper abdomen in progressive cephalocaudal order, commencing in childhood or adolescence. Variable fat loss of the palms, but no loss of intramarrow or retro-orbital fat has been demonstrated [47]. The peripheral and central changes in adipose distribution have been assessed using MRI, which has also helped to distinguish APL from FPLD. In a study by Al-Attar et al. [6], APL-affected subjects were found to have increased subcutaneous calf adiposity compared to subjects with FPLD. In fact, APL-affected subjects

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demonstrated the highest adiposity within the mid-calf region compared to subjects affected with HIV-related lipodystrophy, CGL, or FPLD, thereby providing a key distinguishing feature [6]. Misra et al. have also demonstrated normal to increased subcutaneous adiposity in the glutei, thighs, and soles [47]. There is a strong association of APL with autoimmune disorders, low C3 levels, and membranoproliferative glomerulonephritis. Those with membranoproliferative glomerulonephritis have been reported to develop lipodystrophy at an earlier age compared to those without any renal disease. Although the prevalence of diabetes has been shown to be only around 10%, those who did develop diabetes were primarily females [47, 48]. 20.3.2.2 Molecular Genetics Thus far, only heterozygous mutations in the reannotated LMNB2 gene have been associated with APL. We studied nine unrelated patients with APL and in four of them, we found three new rare LMNB2 mutations: intron 1 –6G>T, exon 5 p.R215Q (in two patients), and exon 8 p.A407T. Compared with a multiethnic control sample of 1100 subjects, the relative risk of APL for carriers of these mutations was 110 (95% conﬁdence interval 36–271; P < 0.00001). These novel mutations were the ﬁrst reported in LMNB2 and the ﬁrst reported among patients affected with APL. The exact mechanism by which mutations in LMNB2 cause APL is unknown. Among our patients, there was no obvious genotype–phenotype correlation. However, our ﬁndings indicate how sequencing of a reannotated candidate gene can reveal new disease-associated mutations [48]. 20.3.3 HIV-Related Lipodystrophy

HIV-related lipodystrophy represents the most common form of lipodystrophy (acquired or genetic) recognized thus far. This form of lipodystrophy affects males and females equally, and has been related to antiretroviral therapy. Fat redistribution among HIV-infected individuals is quite common, affecting approximately 30–50% of individuals, although there is no standard deﬁnition [49]. Since not all individuals exposed to HIV and antiretroviral drugs develop lipodystrophy, a possible genetic susceptibility component can be postulated. Initially, patients with HIV-related lipodystrophy were mistakenly presumed to have Cushings syndrome due to the presence of a prominent dorsocervical fat pad. However, careful endocrinologic testing revealed that this adipose redistribution was unrelated to altered corticosteroid metabolism [50–52]. Later reports linked the presence of peripheral lipoatrophy (on the face and extremities) with central lipohypertrophy (speciﬁcally dorsocervical and truncal regions). The most recent prospective trials have shown that although peripheral lipoatrophy is indeed commonly found among HIV-infected individuals, fat distribution in the truncal region of HIV-infected individuals can range from lipoatrophy to lipohypertrophy. These studies have shown that peripheral lipoatrophy is not always linked with central lipohypertrophy [53–55].

20.4 Lipodystrophy Associated with other Syndromes

We have shown that HIV-related lipodystrophy demonstrates a similar body-wide adipose distribution to FPLD3 with increased supraclavicular and visceral as well as decreased thigh and mid-calf adiposity. Furthermore, subjects affected with FPLD3 or HIV-related lipodystrophy lacked changes in their subcutaneous abdominal and gluteal adipose stores, making these two lipodystrophies relatively similar phenotypically, except for depleted facial fat in HIV-related lipodystrophy [6]. Other endocrinologic manifestations of HIV-related lipodystrophy include hypertriglyceridemia, low HDL cholesterol, insulin resistance, impaired glucose tolerance/diabetes, and androgen deﬁciency [56, 57]. Hepatic steatosis has been observed among HIV-infected individuals [58]. PCOS has not been found to occur among HIVinfected females and acanthosis nigricans is also a relatively rare clinical ﬁnding, despite the signiﬁcant insulin resistance seen in this population [59]. In contrast to the other lipodystrophy forms discussed above, serum leptin levels tend to be preserved and even elevated in conjunction with low serum adiponectin [60, 61].

20.4 Lipodystrophy Associated with other Syndromes 20.4.1 Mandibuloacral Dysplasia 20.4.1.1 Clinical Features Mandibuloacral dysplasia (MAD; MIM 248370) is a rare autosomal recessive disorder characterized by mandibular and clavicular hypoplasia; postnatal growth retardation with delayed closure of cranial sutures; musculoskeletal anomalies including resorption of distal phalanges (acro-osteolysis), joint contractures, and short stature; and progeroid features of bird-like face, high-pitched voice, and ectodermal defects such as mottled hyperpigmentation, alopecia, and nail dysplasia. Fat distribution changes typically are noted in childhood and can be subdivided into two types – type A showing partial loss of subcutaneous adipose tissue in the extremities, and type B showing generalized loss of subcutaneous adipose tissue in the face, trunk, and extremities. Endocrine abnormalities include insulin resistance, hypertriglyceridemia, low plasma HDL cholesterol, and impaired glucose tolerance [62, 63]. 20.4.1.2 Molecular Genetics MAD has been associated with homozygous missense mutations in LMNA (MAD type A; MIM 248370) as well as compound heterozygous mutations in ZMPSTE24 (MAD type B; MIM 608612). LMNA mutations have been reported in several families and have involved homozygous mutations in R527H or A529V [64–67]. In nine affected individuals in ﬁve consanguineous families from Italy, the same homozygous missense R527H mutation was found and all shared the same haplotype, indicating a possible founder effect [64]. ZMPSTE24 encodes a zinc metalloproteinase involved in the post-translational processing of prelamin A to lamin A. Hence, mutations in ZMPSTE24 lead to accumulation of lamin A precursors. Thus far,

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mutations in ZMPSTE24 have been reported in two unrelated individuals, both of whom died prematurely from focal segmental glomerulosclerosis, which seems to be a unique feature of this genotype [68, 69]. 20.4.2 SHORT Syndrome 20.4.2.1 Clinical Features SHORT syndrome (MIM 269880) is a very rare disorder consisting of Short stature, Hyperextensibility of joints and/or inguinal hernia, Ocular depression, Rieger anomaly (defective development of cornea and iris), and Teething delay, affecting males and females equally. Inheritance can be either autosomal recessive or dominant. The majority of affected individuals demonstrate adipose tissue loss in the face, upper extremities, and trunk, with less effect on lower extremities. Other individuals have demonstrated adipose tissue loss of the trunk, gluteal region, and elbows only [70–72]. 20.4.2.2 Molecular Genetics The molecular basis for this disorder remains to be elucidated. 20.4.3 Neonatal Progeroid Syndrome 20.4.3.1 Clinical Features Wiedemann–Rautenstrauch syndrome (MIM 264090), also known as neonatal progeroid syndrome, shows autosomal recessive inheritance. Affected individuals are noted at birth to have progeroid features of enlarged skull, triangular-shaped and older-looking face, and minimal scalp hair. As well, a large anterior fontanel with prominent veins on the scalp is often noted. Fat loss is evident in a generalized pattern, with sparing of the gluteal and sacral regions. Premature death in the neonatal period occurs commonly and has been typically attributed to cerebral hemorrhage or respiratory causes [73]. 20.4.3.2 Molecular Genetics No mutations in known lipodystrophy genes or in candidate genes encoding nuclear envelope proteins have been found in affected patients by our group (Hegele, unpublished observations). 20.4.4 Hutchinson–Gilford Progeria Syndrome 20.4.4.1 Clinical Features HGPS (MIM 176670) was ﬁrst described in the late 1800s and was subsequently divided into two categories – classical HGPS and nonclassical HGPS. Classical HGPS follows an autosomal dominant inheritance pattern whereas nonclassical

20.5 Conclusions

HGPS has demonstrated autosomal recessive inheritance. The typical clinical features of HGPS include postnatal growth delay with normal skull growth, extreme short stature, premature loss of scalp hair with prominence of scalp vessels, decreased joint mobility, osteolysis, and typical facial features of an elderly person. Importantly, cognitive development is normal. The lipodystrophic features can begin as early as 6 months of age with initial adipose tissue loss involving the extremities, followed by the trunk, and ﬁnally the face with sparing of intra-abdominal fat. Cardiovascular and cerebrovascular atherosclerotic disease is very common among individuals with HGPS, and comprises the major cause of premature death. Individuals with nonclassical HGPS tend to follow a milder clinical course and can occasionally live into the third decade of life compared with the average age of death of around 13 years in classical HGPS [74]. 20.4.4.2 Molecular Genetics HGPS is one of the laminopathies, with mutations found in the LMNA gene. The most common mutation is the G608G in the LMNA gene of paternal origin, creating a cryptic splice site resulting in the formation of mutant prelamin A [75, 76]. Other LMNA mutations for HGPS include R471C, R527C, G608S, and c.2036C>T [77]. 20.4.5 Werner Syndrome 20.4.5.1 Clinical Features Werner syndrome (MIM 277700) is one of the progeroid syndromes. Affected individuals display clinical features characteristic of premature aging, including development of premature graying, diabetes, osteoporosis, cataracts, and predisposition to early death from cardiovascular disease or cancer. This syndrome is inherited in an autosomal recessive fashion, with premature death occurring in the late ﬁfth decade of life. Associated features of this syndrome include decreased subcutaneous fat deposition in the trunk, face, and extremities; insulin resistance; short stature; hypogonadism leading to reduced fertility; dermatologic manifestations of telengiectasia, hyperkeratosis, predisposition to trophic ulcers; and calciﬁcation of the blood vessels [78, 79]. 20.4.5.2 Molecular Genetics Werner syndrome occurs as a result of homozygous mutations in RECQL2/WRN (MIM 604611), encoding a DNA helicase important in maintaining genomic stability through functions in both single strand and double strand repair [80].

20.5 Conclusions

Lipodystrophies highlight the importance of adipose tissue distribution in human metabolism since alterations in adipose distribution are associated with

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the development of several metabolic and clinical abnormalities, including insulin resistance, glucose intolerance, hypertension, and hepatic steatosis. Not only has the critical role of adipocyte differentiation and adipokine release in human metabolism only been recently discovered, but also novel therapeutic options based on these pathways (e.g., PPAR-c agonists, leptin replacement) have been developed and utilized with positive improvements in the metabolic abnormalities of lipodystrophies. Importantly, molecular genetics has also added to our understanding of lipodystrophies. Thus far, genetic characterization of lipodystrophies includes mutations in two nuclear envelope structural components (LMNA and LMNB2), a nuclear hormone receptor (PPARG), a zinc metalloproteinase (ZMPSTE24), an integral endoplasmic reticulum membrane protein (BSCL2), and a lipid biosynthetic enzyme (AGPAT2). Signiﬁcant genetic heterogeneity exists in terms of both the number of genes involved and the variety of responsible mutations. Hence, there does not appear to be a single unifying mechanism for the development of lipodystrophy, but rather a complex and intricate interaction between multiple protein players and pathways, both intra- and extracellular. The exact mechanism by which mutations in these pathways result in the lipodystrophy phenotype and associated metabolic abnormalities remains to be clariﬁed. Nonetheless, further characterization of the pathways involved in the pathogenesis of lipodystrophy may be useful for the development of novel therapeutic options of more common clinical disorders, including the metabolic syndrome, HIV-related lipodystrophy, and diabetes. Acknowledgments

Supported by the Jacob J. Wolfe Distinguished Medical Research Chair, the Edith Schulich Vinet Canada Research Chair (Tier I) in Human Genetics, a Career Investigator award from the Heart and Stroke Foundation of Ontario, and operating grants from the Canadian Institutes for Health Research, the Heart and Stroke Foundation of Ontario (NA 5320), and the Ontario Research Fund, and by Genome Canada through the Ontario Genomics Institute.

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absence of adipose tissue matter in the upper half of the body. Am. J. Med. Sci., 90, 105–106. Barraquer, L. (1907) Histoire clinique dun cas datrophie du tissue celluloadipeux. Neurolog. Zentralblatt, 26, 1072. Simons, A. (1911) Lipodystrophia progressive. Z. Ges. Neurol. Psychiat., 5, 29–38. Misra, A. and Garg, A. (2004) Clinical features and metabolic and autoimmune derangements in acquired partial lipodystrophy: report of 35 cases and review of the literature. Medicine, 83, 18–34. Hegele, R.A., Cao, H., Liu, D.M., Costain, G.A., Charlton-Menys, V., Rodger, N.W., and Durrington, P.N. (2006) Sequencing of the reannotated LMNB2 gene reveals novel mutations in patients with acquired partial lipodystrophy. Am. J. Hum. Genet., 79, 383–389. Chen, D., Misra, A., and Garg, A. (2002) Lipodystrophy in human immunodeﬁciency virus-infected patients. J. Clin. Endocrinol. Metab., 87, 4845–4856. Lo, J.C., Mulligan, K., Tai, V.W., Algren, H., and Schambelan, M. (1998) Buffalo hump in men with HIV-1 infection. Lancet, 351, 867–870. Miller, K.K., Daly, P., Sentochnik, D., Doweiko, J., Samore, M., Basgoz, N.O., and Grinspoon, S.K. (1998) PseudoCushings syndrome in human immunodeﬁciency virus-infected patients. Clin. Infect. Dis., 27, 68–72. Roth, V., Kravcik, S., and Angel, J. (1998) Development of cervical fat pads following therapy with human immunodeﬁciency virus type 1 protease inhibitors. Clin. Infect. Dis., 27, 65–67. FRAM Investigators (2006) Fat distribution in women with HIV infection. J. Acquir. Immune Deﬁc. Syndr., 42, 562–571. Bacchetti, P., Gripshover, B., Grunfeld, C., Heymsﬁeld, S., McCreath, H., Osmond, D., Saag, M., Scherzer, R. et al. (2005) Study of fat redistribution and metabolic change in HIV infection (FRAM). Fat

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Helbling-Leclerc, A., DApice, M.R., Massart, C., Capon, F., Sbraccia, P. et al. (2002) Mandibuloacral dysplasia is caused by a mutation in LMNA-encoding lamin A/C. Am. J. Hum. Genet., 71, 426–431. Shen, J.J., Brown, C.A., Lupski, J.R., and Potocki, L. (2003) Mandibuloacral dysplasia caused by homozygosity for the R527H mutation in lamin A/C. J. Med. Genet., 40, 854–857. Simha, V., Agarwal, A.K., Oral, E.A., Fryns, J.P., and Garg, A. (2003) Genetic and phenotypic heterogeneity in patients with mandibuloacral dysplasia-associated lipodystrophy. J. Clin. Endocrinol. Metab., 88, 2821–2824. Garg, A., Cogulu, O., Ozkinay, F., Onay, H., and Agarwal, A.K. (2005) A novel homozygous Ala529Val LMNA mutation in Turkish patients with mandibuloacral dysplasia. J. Clin. Endocrinol. Metab., 90, 5259–5264. Agarwal, A.K., Fryns, J.P., Auchus, R.J., and Garg, A. (2003) Zinc metalloproteinase, ZMPSTE24, is mutated in mandibuloacral dysplasia. Hum. Mol. Genet., 12, 1995–2001. Agarwal, A.K., Zhou, X.J., Hall, R.K., Nicholls, K., Bankier, A., Van Esch, H., Fryns, J.P., and Garg, A. (2006) Focal segmental glomerulosclerosis in patients with mandibuloacral dysplasia owing to ZMPSTE24 deﬁciency. J. Invest. Med., 54, 208–213. Gorlin, R.J., Cervenka, J., Moller, K., Horrobin, M., and Witkop, C.J. Jr. (1975) Rieger anomaly and growth retardation (the S-H-O-R-T syndrome). Birth Defects Orig. Artic. Ser., 11, 46–48. Sorge, G., Ruggieri, M., Polizzi, A., Scuderi, A., and DiPietro, M. (1996) SHORT syndrome: a new case with probable autosomal dominant inheritance. Am. J. Med. Genet., 61, 178–181.

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N. (1983) Autosomal dominant partial lipodystrophy associated with Rieger anomaly, short stature, and insulinopenic diabetes. Am. J. Med. Genet., 15, 29–38. Pivnick, E.K., Angle, B., Kaufman, R.A., Hall, B.D., Pitukcheewanont, P., Hersh, J.H., Fowlkes, J.L., Sanders, L.P. et al. (2000) Neonatal progeroid (Wiedemann–Rautenstrauch) syndrome: report of ﬁve new cases and review. Am. J. Med. Genet., 90, 131–140. Hennekam, R.C.M. (2006) Hutchinson–Gilford progeria syndrome: review of the phenotype. Am. J. Med. Genet., 140A, 2603–2624. Eriksson, M., Brown, W.T., Gordon, L.B., Glynn, M.W., Singer, J., Scott, L., Erdos, M.R., Robbins, C.M. et al. (2003) Recurrent de novo point mutations in lamin A cause Hutchinson–Gilford progeria syndrome. Nature, 423, 293–298. De Sandre-Giovannoli, A., Bernard, R., Cau, P., Navarro, C., Amiel, J., Boccaccio, I., Lyonnet, S., Stewart, C.L. et al. (2003) Lamin A truncation in Hutchinson–Gilford progeria. Science, 300, 2055. Cao, H. and Hegele, R.A. (2003) LMNA is mutated in Hutchinson–Gilford progeria (MIM 176670) but not in Wiedemann–Rautenstrauch progeroid syndrome (MIM 264090). J. Hum. Genet., 48, 271–274. Lebel, M. (2001) Werner syndrome: genetic and molecular basis of a premature aging disorder. Cell. Mol. Life Sci., 58, 857–867. Hoepffner, H.-J., Dreyer, M., Reimers, U., Schmidt-Preuss, U., Koepp, H.-P., and Rudiger, H.W. (1989) A new family syndrome with impaired function of three related peptide growth factors. Hum. Genet., 83, 209–216. Navarro, C.L., Cao, P., and Levy, N. (2006) Molecular bases of progeroid syndromes. Hum. Mol. Genet., 15, R151–R161.

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21 Mouse Models of Lipodystrophy Jimmy Donkor and Karen Reue 21.1 Introduction

Lipodystrophy is the lack of functional adipose tissue. In humans, both congenital and acquired forms of lipodystrophy are known, and are associated with metabolic dysregulation characterized by insulin resistance, hypertriglyceridemia, and fatty liver [1, 2]. These phenotypes are also present in most lipodystrophic mouse models, making these models a valuable tool to elucidate the mechanisms of adipose tissue development and function, and to understand the abnormalities that occur in lipodystrophy [3–7]. The study of several genetically modiﬁed mouse models has revealed that lipodystrophy can result from defects in any of several genes that inﬂuence processes such as adipocyte differentiation, triacylglycerol biosynthesis, or energy metabolism. Here, we classify several lipodystrophic mouse models into groups based on the etiology of the lipodystrophy, and summarize the molecular, physiological, and metabolic characteristics of each model that may be pertinent to understanding and treating lipodystrophy in humans.

21.2 Physiological Mechanisms of Lipodystrophy in Mouse Models

Over the past several years, numerous mouse models of lipodystrophy have been developed and characterized. These include genetically engineered mouse models, mouse strains with spontaneous mutations in endogenous genes, and chemically induced lipoatrophy. In this chapter, we categorize lipodystrophic mouse models into groups based on the mechanism(s) by which adipose tissue development or function is impaired (Figure 21.1). The ﬁrst category is deﬁned as models in which lipodystrophy is caused by disruption of functions that are required during early

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Impaired Adipogenesis or Increased Cell Death

Impaired Triglyceride Biosynthesis

A/ZIP Tg aP2-SREBP-1c Tg PPARγ KO C/EBPα KO Zmpste24 KO aP2-DTA Tg RSK2 KO

GPAT1 KO

Lipin-1 deficient

FAT-ATTAC Tg

AGPAT6 KO DGAT1 KO

aP2-leptin Tg PPARδ Tg FOXC2 Tg

Enhanced Energy Expenditure Figure 21.1 Physiological basis for lipodystrophy in genetically modified mouse models. TG, transgenic; KO, knockout. Full names of each animal are given in the text.

stages of adipocyte differentiation. A second group of lipodystrophic models is characterized by deﬁciencies in enzymes required for triacylglycerol biosynthesis in adipocytes. The ﬁrst two groups of mouse models most closely resemble forms of congenital lipodystrophy in humans, typically affecting the initial development and accumulation of adipose tissue. A third group of models are animals in which lipodystrophy results from the enhanced utilization of stored triacylglycerol, resulting in decreased adipose tissue mass. Finally, we consider mouse models in which lipoatrophy occurs through the destruction of normal adipose tissue, through either genetic alterations or through diet or drug treatments. These models may be useful for elucidation mechanisms of acquired lipodystrophy in humans. These categorizations are proposed as a means of organizing a discussion of the several models, but it should be appreciated that in many cases a combination of these mechanisms may conspire to produce the resulting reduced fat mass and impaired metabolism. A general message from studies of mouse models of lipodystrophy is that normal adipose tissue function is required for glucose, lipid, and energy homeostasis. However, the speciﬁc differences that exist between models are also instructive, and these are highlighted in the descriptions of models that follow and are summarized in Table 21.1. Note that numerous additional mouse models with less profound decreases in adipose tissue, as well as models with increased adipose tissue, have also been of great value in elucidating the many physiological

Table 21.1

Metabolic characteristics of mouse lipodystrophy models. Level of WAT reduction (%)

BAT adiposity

Fatty liver

Dyslipidemia

Insulin resistance

Increased energy expenditure

Reduced leptin levels

Other molecular, physiological, and metabolic characteristics

A-ZIP/F1 Tg

>90

inactive

yes

yes

yes

no

yes

aP2-SREBP-1c Tg

>90

enlarged

yes

yes

yes

no

yes

Global PPAR-c KO (after embryonic rescue) Adipose-speciﬁc PPAR-c KO

>90

reduced

no

yes

yes

?

yes

development of hypertension and renal injury ﬁrst model to demonstrate leptin administration reverses metabolic dysregulation of lipodystrophy low blood pressure

>50

reduced and inactive

yes

yes

yes

?

yes

C/EBP-a KO (after liver rescue)

>90

normal

yes

yes

yes

?

yes

Zmpste24 KO

>90

?

?

?

?

?

?

GPAT1 KO

20–30

?

no

no

no

?

?

AGPAT6 KO

30–40

reduced

no

no

no

yes

?

DGAT1 KO

40

?

no

no

no

yes

?

(Continued)

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abnormal adipocytes, enlarged and reduced in number lipodystrophy affects some white adipose depots, but not others development of muscular dystrophy, cardiomyopathy, bone fractures, and premature aging triacylglycerol biosynthesis defect subdermal lipodystrophy, increased energy expenditure triacylglycerol biosynthesis defect, increased energy expenditure

21.2 Physiological Mechanisms of Lipodystrophy in Mouse Models

Mouse model

Mouse model

Level of WAT reduction (%)

BAT adiposity

Fatty liver

Dyslipidemia

Insulin resistance

Increased energy expenditure

Reduced leptin levels

Other molecular, physiological, and metabolic characteristics

Lipin-1-deﬁcient

>90

reduced and inactive

only in neonate

only in neonate

yes

yes

yes

Leptin Tg

>90

no

no

no

aP2-leptin Tg (6–9 weeks of age)

>90

reduced

no

no

no

yes

no

aP2-PPAR-d Tg

40

normal

no

no

no

yes

?

aP2-FOXC2 Tg

80

enlarged

no

no

no

yes

?

aP2-DTA Tg FAT-ATTAC Tg RSK2 KO Drug-induced lipoatrophy (HIV protease inhibitors) Diet-induced lipoatrophy (CLA)

>90 >90 80 25

reduced reduced normal ?

yes no yes ?

yes no yes yes

yes yes yes yes

? yes ? ?

? yes yes ?

85

reduced

yes

yes

yes

no

yes

spontaneous mutation; peripheral neuropathy, susceptibility to diet-induced atherosclerosis favorable metabolic proﬁle despite lack of WAT initial lipodystrophy followed by increased body weight, fat mass, and lipid accumulation at older ages (33–37 weeks) enhanced thermogenesis and fatty acid oxidation increased sensitivity to adrenaline and increased fatty acid oxidation Late-onset lipodystrophy reversible lipodystrophy late-onset lipodystrophy affects subcutaneous, but not visceral, adipose tissue depots lipoatrophy associated with inﬂammation of adipose tissue

Tg, transgenic; KO, knockout. Full names of each animal model are given in the text.

no

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Table 21.1 (Continued)

21.3 Lipodystrophic Models with Impaired Adipogenesis

roles of adipose tissue. These models have been reviewed in excellent articles elsewhere [3, 4, 8–12].

21.3 Lipodystrophic Models with Impaired Adipogenesis

White adipose tissue (WAT) is designed to provide efﬁcient long-term energy storage in mammals [13]. It carries out the synthesis, storage and hydrolysis of triacylglycerol – the most efﬁcient form in which to store energy. Importantly, WAT also secretes several proteins that have key roles in metabolism and energy homeostasis [14]. These include the enzyme lipoprotein lipase, which hydrolyzes exogenous triacylglycerol to release fatty acids for tissue uptake, inﬂammatory cytokines, components of the complement pathway involved in host defense, and peptide hormones with important functions in the regulation of food intake, glucose homeostasis, and energy expenditure. Brown adipose tissue (BAT) also stores triacylglycerol, but is functionally and morphologically distinct from WAT. In contrast to WAT, BAT has a specialized metabolic role in energy expenditure and heat generation, rather than energy storage [13]. In white adipocytes, the stored lipid is present as a single large droplet that occupies most of the cell volume, whereas brown adipocytes are ﬁlled with smaller lipid droplets that are often associated with abundant mitochondria. The hallmark difference between brown and white adipocytes is the expression of uncoupling protein (UCP)-1, which facilitates thermogenesis by promoting proton leakage across the inner mitochondrial membrane to uncouple proton transport from ATP generation. As BAT is more prevalent in small mammals such as mice than in humans, mouse models of lipodystrophy allow the study of effects on both white and BAT. WAT development involves the conversion of ﬁbroblast-like preadipocytes to mature, lipid-ﬁlled adipocytes through an ordered cascade of alterations in gene transcription and cell morphology [15, 16]. Transcriptional regulation of adipocyte differentiation is driven by the nuclear receptor peroxisome proliferator-activated receptor (PPAR)-c and members of the CAAT/enhancer-binding protein (C/EBP) family. PPAR-c expression is induced by C/EBP-b and -d during the very early stages of differentiation, and C/EBP-a has a role in maintaining PPAR-c expression in maturing adipocytes. PPAR-c and C/EBP-a activate several genes that are required for key adipocyte functions, such as triacylglycerol biosynthesis and storage, and secretion of adipocyte-derived factors such as leptin, adiponectin, and cytokines. PPAR-c and C/EBP proteins are also required, but not sufﬁcient, for the differentiation of precursor cells to brown adipocytes. Very recently it has been shown that differentiation of brown adipocytes is determined by the zinc ﬁnger protein PRDM16 [17]. The action of PRDM16 in brown adipocyte differentiation is not fully understood, but likely involves transcriptional activation of other genes that lead to expression of essential brown adipocyte proteins, such as UCP-1 and PPAR-c coactivator (peroxisome proliferator-activated receptor-c coactivatorPGC)-1a. Characteristics of mouse models with impaired adipocyte differentiation are described in the following subsections.

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21.3.1 A-ZIP/F1 Transgenic Mouse

The A-ZIP/F1 transgenic mouse was one of the ﬁrst genetically engineered mouse models that led to complete impairment of adipose tissue development [18]. The expression of a transgene encoding a dominant-negative transcription factor, termed A-ZIP, leads to the formation of stable heterodimers with endogenous B-ZIP transcription factors and prevents their binding to DNA [19]. In adipose tissue, AZIP interferes with the function of C/EBP-a and AP-1 during adipocyte differentiation, and preadipocytes are arrested in an undifferentiated state. A-ZIP/F1 transgenic mice lack WATand have reduced and inactive BAT [18]. As a consequence, they develop metabolic symptoms similar to those observed in human lipodystrophy, including fatty liver, elevated plasma glucose, insulin, fatty acid, and triacylglycerol levels, and extremely low leptin levels [18]. Transplantation of normal adipose tissue at ectopic locations, or restoration of leptin via transgene expression or injection, improves the hepatic steatosis, insulin resistance, and hyperphagia of A-ZIP/F1 mice, indicating that these symptoms are at least partially due to a lack of normal adipose tissue and adipokine function [20–22]. The A-ZIP/F1 transgenic mouse model has also revealed a relationship between lipodystrophy and hypertension and renal injury. The development of hypertension in A-ZIP/F1 transgenic mice is associated with a lack of perivascular adipose tissue and it is thought that impaired secretion of fat-derived relaxation factors in the vasculature may contribute to hypertension [23]. These results highlight an additional role for adipokines beyond the established roles in appetite regulation, glucose homeostasis, and energy expenditure. The A-ZIP/F1 transgenic mouse also develops renal injury that appears to be related to the absence of normal adipose tissue and the subsequent development of diabetes. The renal damage can be improved or prevented by replacement of leptin through administration of leptin or expression of a leptin transgene [24]. These ﬁndings highlight yet another role for leptin in metabolic homeostasis and also make the A-ZIP/F1 transgenic mouse a potentially useful model for the study of renal damage due to diabetic complications in humans. 21.3.2 aP2- SREBP-1c Transgenic Mouse

The aP2-SREBP-1c transgenic mouse model has provided insight into the metabolic disturbances that occur in congenital generalized lipodystrophy in humans. Sterol regulatory element-binding protein (SREBP)-1c is a transcription factor expressed in liver and adipose tissue, and has a key role in the regulation of genes involved in fatty acid synthesis and glucose utilization [25]. This protein is synthesized as a precursor that resides in the endoplasmic reticulum membrane until cleaved and released, allowing it to transit to the nucleus where it stimulates transcription of target genes. Expression of a transgene encoding the constitutively active nuclear form of SREBP1c in adipose tissue of mice unexpectedly led to the development of lipodystrophy characterized by nearly complete lack of WAT, but increased interscapular BAT

21.3 Lipodystrophic Models with Impaired Adipogenesis

mass [26]. The impaired WAT development in these mice is associated with reduced expression of the key differentiation factors PPAR-c and C/EBP-a, suggesting that the aberrant expression of activated SREBP-1c interferes with the normal adipogenic transcriptional cascade. However, a recent study indicates that embryonic ﬁbroblasts from aP2-SREBP-1c transgenic mice can differentiate into adipocytes in vitro, raising the possibility that the cause of lipodystrophy is impairment at a step beyond adipocyte differentiation [27]. The aP2-SREBP-1c transgenic mice develop insulin resistance with elevated glucose levels, fatty liver, and elevated plasma triacylglycerol levels, and dramatically reduced leptin levels [26]. In the ﬁrst study of its kind, leptin administration was shown to correct the diabetes and other metabolic abnormalities in these animals, without restoring adipose tissue mass, and revealed that leptin deﬁciency is partly responsible for the insulin resistance in lipodystrophy [28]. 21.3.3 Mouse Models with Altered PPAR-c Levels

As described above, PPAR-c is a key transcription factor required for adipocyte differentiation, and also has important roles in lipid and glucose metabolism. Studies of individuals with rare heterozygous mutations in the human PPARG gene have revealed that PPAR-c haploinsufﬁciency causes a partial lipodystrophy syndrome with loss of subcutaneous fat in the extremities, insulin resistance, and diabetes (reviewed in [29, 30]). Analogous to heterozygous human PPAR-c mutants, heterozygous PPAR-c knockout mice and hypomorphic PPAR-c mice with reduced expression have generalized loss of body fat and insulin resistance [31, 32]. Global inactivation of PPAR-c in the mouse causes embryonic lethality, indicating important roles for PPAR-c in placenta and cardiac development [33]. The embryonic lethality can be overcome in PPAR-c-deﬁcient embryos by allowing PPAR-c expression in trophoblasts [34]. The resulting adult mice have global PPAR-c deﬁciency, severe lipodystrophy, and insulin resistance, and have low blood pressure, which is thought to be associated with increased vascular relaxation. Mouse models in which PPAR-c expression levels have been spatially or temporally manipulated have contributed additional understanding of the role of PPAR-c in adiposity in insulin resistance. Adipose tissue-speciﬁc ablation of PPAR-c expression causes lipodystrophy characterized by abnormal fat cells that are large in size and reduced in number, and insulin resistance in fat and liver, but normal insulin sensitivity in muscle [35]. Despite the much lower levels of PPAR-c expression in muscle, its action in that tissue appears to be key in maintaining glucose homeostasis, as muscle-speciﬁc ablation of PPAR-c causes insulin resistance and failure to respond to thiazolidinedione drugs, which enhance insulin sensitivity by activating PPAR-c [36]. A recently reported model allows the reversible inactivation of PPAR-c and reversible lipodystrophy. Mice with an engineered PPAR-c allele that contains a Tet activator and a Tet activator-regulated PPAR-c transgene unexpectedly develop a dominantly inherited lipodystrophy [27]. The phenotype resembles that of the aP2SREBP-1c transgenic mouse described above in the occurrence of an enlarged buffalo hump of interscapular BAT and generalized lack of WAT, as well as insulin

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resistance and hyperlipidemia. A notable feature of this model is that the administration of doxycycline in the drinking water from midgestation onward can prevent lipodystrophy, or when administered to pubertal mice, can reverse the metabolic defects other than reduced WAT mass. This mouse strain may be valuable for elucidating the ongoing requirement for PPAR-c in mature adipocytes and as a model of acquired lipodystrophy. 21.3.4 C/EBPa-Deficient Mouse

As described above, C/EBP-a is a transcription factor that is required for adipocyte differentiation and is also expressed at high levels in the liver [37]. Total ablation of C/ EBP-a in all tissues leads to neonatal lethality due to severe hypoglycemia [38]. However, the lethality of C/EBP-a knockout can be rescued by C/EBP-a expression in the liver [39]. These mice do not develop subcutaneous, perirenal, or epididymal WAT, but do have BAT and mammary WAT. These mice also have postprandial hyperlipidemia, fatty livers, and elevated serum insulin despite normal glucose levels [39]. This mouse model suggests a requirement for C/EBP-a expression in WAT differentiation in some depots, but not in BAT or in mammary WAT differentiation. 21.3.5 Zmpste24-Deficient Mice

In humans, point mutations in the gene encoding lamin A/C – intermediate ﬁlament proteins that are components of the nuclear envelope – cause Dunnigan-type familial partial lipodystrophy (reviewed in [40]). Interestingly, distinct mutations in the same gene cause other disorders including muscular dystrophy, cardiomyopathy, and progeria. Mouse models deﬁcient in Zmpste24 – a protease that is required in the multistep processing of prelamin A to mature lamin A – have revealed that accumulation of an incompletely processed, lipid-modiﬁed (farnesylated) version of lamin A disrupts the nuclear architecture and results in many of the same phenotypes associated with lamin A mutations. Zmpste24 knockout mice exhibit lipodystrophy, muscular dystrophy, cardiomyopathy, and premature ageing associated with spontaneous bone fractures and reduced bone volume [41, 42]. A compound mouse model of Zmpste24 deﬁciency and heterozygous lamin A knockout revealed that some of the defects in Zmpste24 knockout mice can be overcome by reducing the amount of prelamin A that is synthesized and therefore accumulates due to impaired processing [43]. Further studies with the Zmpste24-deﬁcient mouse model demonstrated that farnesyltransferase inhibitors that prevent lipid modiﬁcation of prelamin A also reduce the incidence of misshapen nuclei, and increase body weight, bone integrity, and lifespan of Zmpste24-deﬁcient mice [44]. The insights gained in these studies have led to potential therapeutic strategies for disorders of lamin A mutations and also for lipodystrophy associated with protease inhibitor drugs used to treat HIV infection.

21.4 Lipodystrophic Models with Impaired Triacylglycerol Biosynthesis

HIV protease inhibitors cause a similar accumulation of farnesyl-prelamin A as seen in Zmpste24-deﬁcient mice. Thus, it appears that the inhibition of Zmpste24 activity by HIV protease inhibitors may promote the accumulation of unprocessed lamin A and contribute to the side-effects of these drugs, including lipodystrophy.

21.4 Lipodystrophic Models with Impaired Triacylglycerol Biosynthesis

Mature white and brown adipocytes synthesize triacylglycerol primarily through the glycerol phosphate pathway – a series of reactions in which glycerol-3-phosphate is acylated through the stepwise addition of three fatty acid moieties (reviewed in [45]). Distinct enzymes are required for each step in this process. Transfer of the ﬁrst fatty acid to the glycerol backbone is catalyzed by one of the glycerol phosphate acyltransferase (GPAT) enzymes, which reside in the endoplasmic reticulum and mitochondria. An additional fatty acid is subsequently transferred to monoacylglycerol through the action of acylglycerolphosphate acyltransferase (AGPAT) enzymes – a family consisting of several members. The resulting phosphatidate molecule can serve as a precursor for either phospholipids or for diacylglycerol (the immediate precursor of triacylglycerol). The conversion of phosphatidate to diacylglycerol occurs through the action of phosphatidate phosphatase (PAP)-1, also known as lipin-1. Finally, diacylglycerol is converted to triacylglycerol through the action of diacylglycerol acyltransferase (DGAT) enzymes. Despite the fact that triacylglycerol biosynthetic enzyme activities have been studied for decades, there are still many questions about the physiological role of speciﬁc enzymes. There appear to be at least four distinct GPATgenes, eight putative AGPAT genes, three lipin genes, and two DGAT genes. In humans, mutations in AGPAT2 cause generalized lipodystrophy, with absence of abdominal and subcutaneous adipose tissue, systemic insulin resistance, and hypertriglyceridemia (reviewed in [1]). Thus far, mutant mouse models for only a handful of triacylglycerol synthesis genes have been studied, but have begun to shed light on the physiological roles of various enzyme isoforms. Mice deﬁcient in each of the triacylglycerol biosynthetic enzymes studied thus far exhibit reduced adipose tissue, but only one model (lipin-1-deﬁcient mice) exhibits a severe lack of adipose tissue that leads to the metabolic consequences typical of lipodystrophy. These results suggest that compensatory mechanisms, such as action of other enzyme isoforms and/or increased energy expenditure, may safeguard against severely detrimental effects. Following are characteristics of mouse models with defects in triacylglycerol synthesis enzymes. 21.4.1 GPAT1-Deficient Mouse

Mice lacking mitochondrial GPAT1 – the enzyme that catalyzes addition of the ﬁrst acyl group to glycerol-3-phosphate during triacylglycerol biosynthesis – have modest

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20–30% reductions in body weight and percent body fat [46]. These mice also have 30–40% reductions in plasma and liver triacylglycerol levels. Unlike models with more severe reduction in adipose tissue, GPAT1-deﬁcient mice do not exhibit altered glucose or insulin levels and in fact have increased hepatic insulin sensitivity [47]. These results suggest that GPAT1 is not the sole GPAT activity in adipose tissue and that mutations in GPAT1 in humans may be unlikely to cause lipodystrophy. 21.4.2 AGPAT6-Deficient Mouse

AGPAT6 is one of eight putative AGPAT enzymes identiﬁed by predicted amino acid sequence similarities [48]. AGPAT6 is expressed at high levels in BAT and WAT, and in liver [49]. AGPAT6-deﬁcient mice have subdermal lipodystrophy, with nearly undetectable subcutaneous adipose tissue and 30–40% reductions in BAT and gonadal WAT fat pad mass, and smaller fat cell size and triacylglycerol content. These results suggest that the AGPAT6 isoform may have a unique role in subcutaneous, but not visceral, adipose tissue. These mice are also resistant to both dietinduced and genetically induced obesity, which could be accounted for, in part, by increased energy expenditure. Unlike models of more severe adipose tissue reduction, AGPAT6 mice have normal glucose and insulin levels. 21.4.3 DGAT1-Deficient Mouse

DGAT1 catalyzes the ﬁnal step of triacylglycerol synthesis. DGAT1-deﬁcient mice revealed an initially surprising phenotype – normal body weight and normal appearing white adipocytes – indicating that adipose tissue is capable of synthesizing triacylglycerol without DGAT1 [50]. However, these animals were found to have a 40% reduction in fat pad mass, and are resistant to diet-induced and genetically induced obesity, which can be accounted for in part by increased activity and metabolic rate [50, 51]. As with the GPAT1-deﬁcient mice, the modest reduction in fat tissue mass in DGAT1 animals is associated with increased insulin sensitivity [51]. In an interesting demonstration of the inﬂuence of adipose tissue-derived hormones on insulin sensitivity, adipose tissue from DGAT1-deﬁcient mice was transplanted into wild-type mice, and led to reduced adiposity and increased glucose disposal [52]. The mechanism may be related to increased secretion of adiponectin or other adipokines from the DGAT1-deﬁcient adipose transplants, and its effect on enhanced fatty acid oxidation and insulin sensitivity. 21.4.4 Lipin-1-Deficient Mouse

Spontaneous mouse mutations leading to lipodystrophy are very rare. To date, the fatty liver dystrophy (ﬂd) mouse is the only lipodystrophic mouse model linked to a spontaneous gene mutation. The ﬂd mouse phenotype is similar to human

21.4 Lipodystrophic Models with Impaired Triacylglycerol Biosynthesis

generalized lipodystrophy, characterized by a lack of body fat in most depots and insulin resistance [6]. These animals differ from other models of lipodystrophy in that they exhibit fatty liver and hypertriglyceridemia only during the neonatal period [53]. In addition, lipin-1-deﬁcient mice have a peripheral neuropathy due to impaired myelin formation, and are susceptible to diet-induced atherosclerosis. Using positional cloning, the ﬂd mouse mutation was identiﬁed as a rearrangement of a novel gene (Lpin1, encoding lipin-1) that led to a nonfunctional mRNA [54]. An independent mutation that occurred in a second mouse strain and resulted in a single amino acid substitution in the lipin-1 protein produced a similar lipodystrophic phenotype as the null mutation in the ﬂd mouse. Lipin-1 is expressed at very high levels in adipose tissue and skeletal muscle, and at lower levels in liver and other tissues [54]. Lipin-1 deﬁnes a family that includes two other closely related proteins, lipin-2 and lipin-3, which appear to have a role in triacylglycerol metabolism in tissues other than adipose tissue [55]. Recent studies have elucidated the molecular function of lipin-1 and shed light on the likely mechanisms of lipodystrophy in lipin-1-deﬁcient mice (reviewed in [56]). Adipocyte differentiation fails to occur in lipin-1-deﬁcient preadipocytes both in vivo and in vitro due to impaired adipogenic gene expression and lipogenesis [57]. The impaired lipogenesis is explained by the recent identiﬁcation of lipin-1 as a PAP-1 enzyme, which catalyzes the conversion of phosphatidate to diacylglycerol in the penultimate step of triacylglycerol synthesis [55, 58, 59]. Lipin-1 is the sole PAP-1 enzyme in adipocytes, explaining, in part, the lipodystrophy in lipin-1-deﬁcient mice [55]. However, additional mechanisms may also contribute, as adipocytes from lipin-1-deﬁcient animals express very low levels of PPAR-c and C/EBP-a, and therefore do not express target genes of these transcription factors that are characteristic of mature adipocytes [57]. The mechanism for this is not clear, but may be related to a second cellular function of lipin-1 – as a transcriptional coactivator. Thus far, transcriptional coactivator activity of lipin-1 has been demonstrated primarily in liver, where it interacts with PPAR-a and PGC-1a to modulate expression levels of lipid metabolism genes [60]. Similar interactions between lipin-1 and PPAR-c may occur in adipocytes. Finally, an additional mechanism that may contribute to lipodystrophy in lipin-1-deﬁcient mice is increased energy expenditure [61]. Thus, deﬁciency in several triacylglycerol synthesis proteins (AGPAT6, DGAT1, and lipin-1) is associated with increased energy expenditure, perhaps induced as a compensatory mechanism in response to the inability to store excess triacylglycerol in adipose tissue. Although no mutations in the human LPIN1 gene have been identiﬁed to underlie lipodystrophy, lipin-1 expression levels in adipose tissue of individuals with lipodystrophy occurring secondary to treatment with protease inhibitors are positively correlated with better conservation of limb fat mass and favorable metabolic parameters [62]. In addition, several studies have demonstrated a positive correlation between lipin-1 expression levels in human adipose tissue and insulin sensitivity in subjects ranging from healthy, young individuals to obese, insulin resistant individuals [63–67]. Differences in lipin-1 expression levels may be related to genetic polymorphisms in the LPIN1 gene, which also show associations with traits such as

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body mass index, insulin sensitivity, energy expenditure, hypertension, and risk for the metabolic syndrome [66, 68, 69].

21.5 Lipodystrophic Models with Enhanced Energy Expenditure

Another category of lipodystrophic mouse models are those with reduced adiposity due to increased energy expenditure. In contrast to most models discussed in previous sections, these animals typically have improved insulin sensitivity, suggesting a functional difference in the lipid-depleted adipose tissue in these animals from that of models in which adipose tissue does not develop normally. 21.5.1 Leptin Transgenic Mouse

Leptin is a hormone synthesized by the adipose tissue that modulates energy balance and metabolism. Leptin affects satiety by its action in the central nervous system and also has important peripheral actions, including effects on hepatic glucose production, energy expenditure, and insulin secretion (reviewed in [70]). Leptin-deﬁcient mice are obese and develop insulin resistance, which is improved by exogenous administration of leptin [71]. As described earlier, leptin replacement in several mouse models of lipodystrophy, as well as in human lipodystrophic patients [72], can ameliorate insulin resistance and fatty liver, indicating that reduced leptin levels is one of the key detrimental effects of adipose tissue loss. The characterization of a transgenic mouse model with constitutive leptin expression in liver has provided further support for this interpretation. The leptin transgenic mice have markedly reduced food intake, as expected from the effect of leptin on satiety, and therefore have a near-complete absence of adipose tissue [73]. Despite the adipose tissue deﬁciency, however, the transgenic mice exhibit enhanced glucose utilization, which is associated with increased insulin signaling in liver and skeletal muscle [20, 73]. Chronically elevated AMP-activated protein kinase activity in the muscle of leptin transgenic mice appears to contribute to the favorable metabolic state of these mice, presumably by increasing fatty acid oxidation to prevent lipid accumulation and insulin resistance in muscle and liver [74]. An independently developed leptin transgenic mouse model in which leptin is expressed in adipose tissue has also been characterized and provides insight into the metabolic effects of short- versus long-term effects of enhanced leptin expression in adipose tissue [75]. These animals exhibited distinct effects of hyperleptinemia at young (6–9 weeks) versus older (33–37 weeks) ages. At the young age, transgenic mice have very small or nonexistent white adipose depots and substantially reduced BATwith lipid deﬁcient adipocytes. These animals also have reduced food intake, and reduced plasma glucose, insulin, and triacylglycerol levels. Thus, the adipose tissue deﬁciency in this model is uncoupled from the metabolic dysregulation that occurs in most lipodystrophic models discussed in earlier sections, presumably due to the

21.5 Lipodystrophic Models with Enhanced Energy Expenditure

presence of high levels of leptin. A striking aspect of the phenotype of these mice is that beginning at 20 weeks of age, they begin to accumulate adipose tissue with increased lipid droplet content in both white and brown adipocytes, and plasma glucose and lipid levels increase. It is unclear whether the appearance of fat depots in older transgenic mice results from decreased responsiveness to the leptin pathway (i.e., leptin resistance) or to compensatory changes that supersede the effect of leptin. The fact that the older transgenic mice still exhibit a normal weight loss response to acute doses of exogenous leptin raises the possibility that molecular changes directly in the adipose tissue underlie the recovery of fat deposition. 21.5.2 PPAR-d Transgenic Mouse

Targeted activation of PPAR-d in the adipose tissue by expression of a ligandindependent active form of PPAR-d causes increased fatty acid oxidation in adipocytes, resulting in substantially reduced triacylglycerol accumulation in adipose tissue without lipid accumulation in liver [76]. One mechanism by which PPAR-d activation affects adiposity is through the enhanced expression of genes involved in fatty acid oxidation and thermogenesis. Furthermore, the treatment of genetically obese mice with a PPAR-d agonist leads to a reduction in stored lipids and an upregulation of thermogenic gene expression in both WAT and BAT. Studies of PPAR-d action in muscle and liver have shown that PPAR-d has important roles in hepatic carbohydrate metabolism and muscle fatty acid oxidation as well as its effect in adipose tissue, and represents a potential therapeutic target for the metabolic dysregulation that accompanies lipodystrophy and obesity [77, 78]. 21.5.3 FOXC2 Transgenic Mouse

FOXC2 belongs to the winged-helix/forkhead family of transcription factors and regulates adipocyte metabolism by increasing adipose tissue sensitivity to the badrenergic/cAMP/protein A signaling pathway. Adipose tissue-speciﬁc expression of FOXC2 in transgenic mice inﬂuences adipose tissue distribution, morphology, and oxidative capacity. The FOXC2 transgenic mice develop an enlarged BAT depot, but show an 80% reduction in WAT mass and improved insulin sensitivity [79]. Gene expression analysis revealed that WAT from the FOXC2 transgenic mice expresses brown adipocyte markers such as UCP-1 and PGC-1a, and has increased adrenergic receptor expression. Consistent with these changes in gene expression, isolated adipose tissue exhibits increased oxygen consumption. In addition, muscle and liver from the FOXC2 transgenic mice are protected from diet-induced insulin resistance associated with reduced fatty acyl-CoA accumulation and reduced hepatic glucose production, respectively [80]. Thus, the reduced adiposity is likely due to enhanced fatty acid oxidation, which leads to reduced triacylglycerol storage in adipose tissue as well as in skeletal muscle and a favorable metabolic proﬁle.

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21.6 Mouse Models with Acquired or Conditional Lipodystrophy

The ﬁnal category of lipodystrophic mouse models are unique in that animals are initially normal and lipodystrophy develops with age, or is induced upon administration of a chemical or dietary inducer. These models may be particularly valuable for the study of mechanisms that trigger adipose tissue destruction through apoptosis, which may have relevance to human acquired lipodystrophies. A few of the models discussed in previous sections (i.e., the Tet-activated PPAR-c transgenic and adiposespeciﬁc leptin transgenic mice) can also be considered as acquired or inducible lipodystrophic animals. 21.6.1 aP2-DTA Transgenic Mouse

The aP2-DTA transgenic mouse was the earliest mouse model of adipose tissue ablation developed. This strain expresses an attenuated diphtheria toxin A (DTA) chain speciﬁcally in adipose tissue [81]. Expression of high levels of the transgene causes lethality at birth, but mice with lower transgene expression levels survive. The aP2-DTA transgenic mice have normal adipose tissue until adulthood, when they begin to lose adipose tissue mass. They are resistant to obesity and infertility normally induced by treatment with monosodium glutamate. As with other lipodystrophic models, these transgenics develop insulin resistance, dyslipidemia, and fatty livers [81]. This strain thus represents a model of late onset lipodystrophy with some similarities to acquired lipodystrophy in humans. 21.6.2 FAT-ATTAC Transgenic Mouse

The fat apoptosis through targeted activation of the caspase 8 (FAT-ATTAC) transgenic mouse was the ﬁrst mouse model to be generated with inducible and reversible lipodystrophy [82]. The FAT-ATTAC transgenic mouse was created by the expression of an adipose tissue-speciﬁc caspase 8–FK506-binding protein (FKBP) fusion protein under the control of the adipose tissue-expressed aP2 promoter. The transgenic mice are normal until treated with a chemical dimerizer (a FK506 analog that binds FKBP), which causes homodimerization of the fusion protein, leading to the activation of caspase and resulting in adipose tissue apoptosis [83]. Treatment with the dimerizer causes generalized lipodystrophy, glucose intolerance, reduced leptin and adiponectin levels, and impaired systemic inﬂammatory response to endotoxin [82]. The lipodystrophy, reduced adipokine levels, and inﬂammatory response are reversed by discontinuation of the drug treatment for 6 weeks. Importantly, functional adipocytes can be recovered from the previously lipodystrophic mice after the cessation of drug treatment, making this an exciting model for the study of adipogenesis in vivo. Adipokine levels and glucose tolerance are also restored in the recovered mice. The use of similar technology could potentially be useful to study the metabolic role and

21.6 Mouse Models with Acquired or Conditional Lipodystrophy

development of speciﬁc adipose tissue depots (BAT, visceral WAT, etc.) once depotspeciﬁc gene regulatory sequences are identiﬁed. 21.6.3 RSK2 Deficient Mouse

Ribosomal S6 kinase 2 (RSK2) is a member of the serine-threonine kinase gene family and is most prominently expressed in WAT. RSK2-deﬁcient mice are normal at birth; however, as they age, they lose up to 80% of their WAT mass, although BAT is not affected [84]. They also develop fatty liver, insulin resistance, and mild diabetes, and have reduced leptin levels. Despite normal food intake, the RSK2 knockout mice are resistant to diet-induced obesity. Impaired glucose tolerance, and elevated fasting insulin and glucose levels are restored in these mice with low-level leptin administration [84]. The mechanism by which RSK2 deﬁciency causes lipodystrophy is not known, but RSK2 has been implicated in cell survival, and it has been suggested that its loss may lead to increased cell death in WAT. 21.6.4 Drug-Induced Lipoatrophy

In addition to congenital forms of lipodystrophy in humans, acquired lipoatrophy occurs during long-term treatment of HIV patients with antiretroviral therapy that includes protease and/or reverse transcriptase inhibitors (reviewed in [85]). This syndrome is characterized by a selective loss of peripheral adipose tissue stores, with fat accumulation in the abdomen and neck, accompanied by hypertriglyceridemia and insulin resistance. These effects have been replicated to some degree in mouse models, which provide useful tools to identify the cellular mechanisms involved in protease inhibitor-induced lipoatrophy. For example, treatment of mice with a lopinavir/ritonavir combination for 8 weeks caused a 25% reduction in peripheral inguinal, but not epididymal, adipose tissue mass [86]. There was an increase in SREBP-1c in inguinal WAT, suggesting a possible role of SREBP-1c in causing lipodystrophy in these mice, as has been observed in aP2-SREBP-1c transgenic mice [26]. One mechanism for the hypertriglyceridemia associated with protease inhibitor treatment was shown to be decreased lipoprotein lipase-mediated clearance of lipoprotein triacylglycerols and reduced fatty acid uptake in adipose tissue [87]. Additional studies performed in vivo and in vitro suggest that multiple mechanisms may contribute to the atrophy of adipose tissue and metabolic consequences in response to HIV protease inhibitors, including apoptosis, impaired adipogenesis, reduced glucose transport, and defective insulin signaling (reviewed in [88]). 21.6.5 Diet-Induced Lipoatrophy

Lipoatrophy and weight loss are also induced in mice by feeding high concentrations of trans-10, cis-12 conjugated linoleic acid (CLA), which occurs naturally in hydrogenated vegetable oils, meat, and dairy products. Numerous studies in mice have

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shown dramatic (85%) reductions in adipose tissue mass with resultant hepatomegaly, fatty liver, and insulin resistance (reviewed in [89]). Gene expression proﬁling revealed that CLA-induced lipoatrophy is associated with inﬂammatory and apoptotic gene expression, and macrophage inﬁltration of adipose tissue [90, 91]. These changes in adipose tissue lead to reduced levels of leptin and adiponectin, which may contribute to insulin resistance, hyperinsulinemia, and expression of lipogenic genes in liver. As with other forms of lipodystrophy, continuous leptin infusion of CLA-treated mice reverses the hyperinsulinemia and hepatomegaly [92]. Studies of CLA administration in humans to promote weight loss have proven less dramatic than in mouse, and may be related to differences in levels of the effective CLA isoforms in the diets used, age, and species differences in the physiology of lipogenesis and fatty acid oxidation (reviewed in [93]). 21.7 Conclusions

As shown in Table 21.1, the metabolic consequences of lipodystrophy appear to depend, at least in part, on the etiology of the adipose tissue deﬁciency. Severe generalized lipodystrophy usually results from the disruption of genes that are required for adipocyte tissue differentiation, such as PPAR-c, C/EBP-a, and lipin-1. The loss of adipose tissue mass in these mouse models is usually accompanied by severe metabolic consequences such as hyperglycemia, insulin resistance, dyslipidemia, and lipid accumulation in liver and skeletal muscle. Glucose and lipid homeostasis in these models are usually improved by the administration of leptin, illustrating the important role that this adipokine plays in systemic metabolic regulation. Lipodystrophic mouse models resulting from deﬁciencies in the triacylglycerol synthesis pathway typically suffer from less severe insulin resistance than models of impaired adipocyte differentiation (except in the case of lipin-1-deﬁcient mice, which have both impaired differentiation and triacylglycerol synthesis). Interestingly, impairment in triacylglycerol synthesis is often accompanied by a compensatory increase in energy expenditure, which may protect these animals from extensive ectopic lipid accumulation in liver. Models having reduced adiposity due to increased energy expenditure can reach the low levels of adipose tissue that are observed in lipodystrophies of other etiologies, but are nevertheless protected from insulin resistance and fatty liver. The likely mechanisms include efﬁcient oxidation of fatty acids to prevent their accumulation and interference with insulin action. Mouse models of lipodystrophy have greatly improved our understanding of adipose tissue development and function, and of the consequences of impaired adipose tissue function in conditions such as obesity and diabetes. Acknowledgments

This authors gratefully acknowledge support from NIH grants HL28481 and HL66621 (to K.R.), and Genomic Analysis Training grant T32-HG002536 (to J.D.).

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22 Caloric Restriction, Longevity, and Adiposity Leanne M. Redman and Eric Ravussin 22.1 Introduction

Antiaging research by modern scientists continue to echo the quest of the Spanish explorer Ponce de Leon, who searched for the Fountain of Youth on the shores of Florida in the early 1500s. In the last quarter of the twentieth century, healthcare efforts and medical interventions have resulted in a tremendous growth in the size of elderly populations. Concurrently, consumer-driven momentum is pushing scientiﬁc research endeavors towards identiﬁcation of the fundamental aspects of aging at the molecular level. Individuals are no longer satisﬁed with simply living longer, but want increased quality of life and prolonged health during their senior years. As a result, basic and clinical research is conducted to understand the physiological and molecular mechanisms of aging in order to postpone and possibly alleviate many of the illnesses typically associated with the aging process. Ironically, as researchers aim to unravel the mysteries of delaying the biological aging process, the current societal environment is marked by overabundant accessibility of food products coupled with a strong trend of reduced physical activity. As obesity rates have risen to over 30% among Americans [1], so has the prevalence of obesity-related chronic diseases such as diabetes mellitus, heart disease, strokes, and chronic obstructive pulmonary disease. This alarming increase in obesity is further coupled with a lower age of onset for the emergence of obesity-related comorbidities [1], and is pushing researchers into focusing on diet and physical activity as means of altering the scope of preventable morbidities. It is now well accepted that obesity may cause up to 300 000 deaths per year in the United States [2]. Furthermore, estimates have been provided for the number of years of life lost due to obesity [3]. Alarmingly, it now seems that babies born at the beginning of the twenty ﬁrst century will have shorter life expectancies than their parents [4].

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22.2 Physiological Changes with Aging

Aging is associated with increased risk of metabolic disorders including overweight, obesity, insulin resistance, type 2 diabetes, atherosclerosis, and cancer. Crosssectional and longitudinal studies in humans and experimental animal models suggest that over-consumption of energy dense foods and lack of physical activity are the leading causes of weight gain, obesity, and other related health issues [5]. Recently, researchers have learned that while increase adipose tissue per se is a health concern, the storage and distribution of fat within the body also has important implications for health risks. In particular, adipose tissues stored centrally in the visceral compartment of the abdomen and in nonadipose tissues, such as liver, heart, pancreas, and skeletal muscle, are considered to be metabolic abnormalities that precede the development of impaired glucose tolerance, hyperlipidemia, insulin resistance, and type 2 diabetes. As individuals age, body weight even if maintained is composed of an increase in fat mass and a decrease in fat-free mass [6, 7]. Adipose tissue stores within the abdominal visceral compartment increase [8], as does the accumulation of ectopic fat in the liver and skeletal muscle [9]. Associated with these changes in body composition and adiposity is the increased incidence and prevalence of diabetes and glucose intolerance in older persons [10, 11]. It seems in our current obesogenic environment that the link between aging and chronic disease may be inevitable. Interventions therefore that can attenuate the age-associated changes in body composition could delay or even prevent the onset of metabolic disturbances with aging and result in an extended lifespan.

22.3 Aging and Caloric Restriction

One approach of counteracting the effects of aging on the health and longevity of organisms has been through prolonged calorie restriction (CR). CR has been shown since the 1930s by McCay et al. to retard the aging process [12], extending the median and maximal lifespan in various models [13]. While the exact mechanisms through which CR is able to extend the lifespan have yet to be fully elucidated, CR reduces metabolic rate and oxidative damage, improves markers of age-related diseases including diabetes such as insulin resistance, and has been shown to alter neuroendocrine and sympathetic nervous system activity in animals [14] (Figure 22.1). Increased lifespan through prolonged CR has been demonstrated in yeast and worms [15, 16], ﬂies [17], ﬁsh, mice, and rats [18–20]. Results from studies on rhesus monkeys suggest that prolonged CR can also oppose many age-associated pathophysiological changes. These studies demonstrate alterations in a variety of body systems, and include learning and behavior changes [20], lower body temperatures [21], lower plasma insulin concentrations [22], and reductions in resting energy expenditure [23]. Since many changes associated with prolonged CR are important to the health and survival of humans, and excessive caloric intake is associated with morbidity and

22.4 Energy Restriction may Alter the Rate of Living

Figure 22.1 Impact of CR on factors related to aging. Nonmutually exclusive candidate mechanisms of CR include: improved mitochondrial function, decreased oxidative damage due to reduced ROS generation and increased ROS removal; altered neuroendocrine function including growth axis, thyroid axis, hypothalamic pituitary axis,

autonomic nervous system, and carbohydrate metabolism; decreased incidence of chronic diseases such as obesity, diabetes and cardiovascular disease; and delayed onset of aging-related markers (i.e., glucose, insulin, dehydroepiandrosterone sulfate, and body temperature).

development of chronic diseases, it has become an important research objective to assess the feasibility, safety, and effects of prolonged CR in well-controlled human trials.

22.4 Energy Restriction may Alter the Rate of Living

The aging process may be inﬂuenced by energy restriction through a reduction in the metabolic rate of living [24], leading ultimately to reduced oxidative damage. Of issue with the rate of living theory is that metabolic rate changes following dietary manipulation are due to several factors, including decreased tissue mass, reduced energy intake with decreased thermic effect of foods, and diminished size of the metabolizing mass [14]. An ongoing controversy among investigators in previous animal studies appears to be whether chronic CR leads to metabolic adaptation – a reduction in the metabolic rate that is out of proportion to the diminished mass of the organism [14]. In 1985, a Food and Agriculture Organization/World Health Organization/United Nations University report [25] deﬁned metabolic adaptation as A process by which a new or

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different steady state is reached in response to a change or difference in the intake of food or nutrients. Given this contextual deﬁnition, metabolic [26], social, or behavioral changes, or changes in expression or translation of genes may be viewed as adaptations. Results from rat and monkey studies on the changes in energy expenditure [27–31] suggest that most of the data collected should be re-evaluated using appropriate methods of normalizing the metabolic rate after adjusting for changes in body size and composition [32]. For example, Blanc et al. recently calculated a 13% reduction in resting energy expenditure after adjusting for fat-free mass in an 11-year study of energy-restricted monkeys [23]. Recently, however, using doubly labeled water to measure total energy expenditure, Selman et al. reported that caloric restricted rats expended 30–50% more energy than expected [33].

22.5 CR and Oxidative Stress

The free radical theory of aging or oxidative stress hypotheses are well-supported theories of aging. It is widely accepted that the metabolic rate of an organism is a major factor in the rate of aging and is inversely related to its lifespan [34]. Additionally, since 1–3% of consumed oxygen is associated with the production of reactive oxygen species (ROS), namely superoxide (O2. ), hydrogen peroxide (H2O2), and the hydroxyl ion (OH. ) [35], the production of these highly reactive molecules from normal aerobic metabolism is also in direct proportion to an organisms metabolic rate. The inﬂuence of CR on the aging process has been evaluated, in part, on the basis of longevity, disease patterns, and age-associated biological changes [36]. Additionally, many investigators have shown that modulation of the oxidative stress of an organism through prolonged CR is able to retard the aging process in various species, including mammals [37–40]. For example, Jung et al. recently reported that the potent antiaging, antioxidative action of CR through ROS reduction can effectively suppress the age-related downregulation of senescence marker protein-30 in rats [37]. Dandona et al. [41] found that protein carbonylation, a common measure of oxidative damage associated with age [42], was reduced after 4 weeks of CR in obese humans. Bevilacqua et al. have shown that the beneﬁcial effects observed with even short- and medium-term CR (40%) include decreased mitochondrial proton leaks, with consequential decreases in VO2, ROS production, and cellular damage [43]. As a result of increased oxygen consumption, aerobic exercise is associated with increased production of ROS in muscle tissues [44], thereby contributing to muscular fatigue and subsequent protein oxidation [45, 46]. Ironically, however, regular exercise is one of the major factors believed to be beneﬁcial in improving quality of life, retarding age-related declines in physiological functioning, providing protective adaptation against oxidative stress in aged organs [39], and possibly delaying the onset of age-related diseases [47, 48]. A major beneﬁt of regular nonexhaustive exercise is the exercise-induced mild oxidative stress. The repeated exercise results in adaptations in the skeletal muscle antioxidant capacity whereby through activation

22.8 What is Known from Humans?

of redox-sensitive signaling pathways [49], the expression of certain antioxidant enzymes (e.g., superoxide dismutase and glutathione peroxidase) are increased, protecting myocytes against the deleterious effects of oxidants and preventing extensive cellular damage [49]. As an essential component of lifestyle modiﬁcation, exercise has further been shown to possess antiatherogenic and antihypertensive effects [50, 51], and delay the development of type 2 diabetes in individuals at risk [52]. 22.6 CR and Cardiovascular Disease

Elevated levels of oxidized low-density lipoprotein (LDL), excessive ROS generation, hypertension, and diabetes are all potential causes for the development of endothelial dysfunction – a precipitating event in the progression of atherosclerosis. These factors are believed to initiate an inﬂammatory response in the injured endothelial tissue. Although studies on short-term energy restriction are inconclusive, long-term CR is associated with sustained reductions in factors related to endothelial dysfunction in humans, such as decreased blood pressure [53], reduced levels of total plasma cholesterol and triglycerides [54, 55], and reduced markers of inﬂammation such as C-reactive protein, interleukin-6, and plasminogen activator inhibitor type-1 [56–59]. A recent long-term CR study in humans supports the feasibility of using CR as a protective effect against atherosclerosis by showing a 40% reduction in carotid artery intima-media thickness in CR participants relative to a control group [60]. 22.7 CR and Insulin Resistance/Type 2 Diabetes Mellitus

Strong evidence in both monkeys [61–64] and humans [54, 65] shows that long-term energy restriction in lean and obese subjects improves insulin sensitivity – a mechanism by which CR may act to extend lifespan. Additionally, prolonged CR reduces fasting glucose and insulin concentration, two factors believed to contribute to the aging process due to protein glycation [66] and mitogenic action [67], respectively. This compelling evidence suggests that weight loss due to CR may be the most effective means of improving insulin sensitivity, thereby decreasing the risk for the development of diabetes mellitus. 22.8 What is Known from Humans? 22.8.1 Centenarians from Okinawa

Probably the most intriguing epidemiological evidence supporting the role of CR in lifespan extension in humans comes from the Okinawans [68]. Compared to most

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industrialized countries, Okinawa, Japan has 4–5 times the average number of centenarians with an estimated 50 in every 100 000 people [69]. Reports from the Japanese Ministry of Health, Labor, and Welfare show that both the average (50th percentile) and maximum (99th percentile) lifespan are increased in Okinawans. From age 65, the expected lifespan in Okinawa is 24.1 years for women and 18.5 years for men compared to 19.3 years for women and 16.2 years for men in the United States [70]. What is interesting about this population is that a low caloric intake was reported in school children on the island more than 40 years ago and later studies conﬁrmed a 20% CR in adults residing on Okinawa compared to mainland Japan [71]. Importantly, reports indicate that the diets which were typically rich in green leafy vegetables, soy, and some ﬁsh were similar with CR interventions providing adequate amounts of nutrients, essential vitamins, and minerals [70]. 22.8.2 Vallejo Study

To our knowledge there is only one other study that tested the effects of CR without malnutrition in nonobese humans [72]. This was a study of alternate-day feeding in 120 men whereby the 60 participants in the CR group received an average of 1500 kcal/day for 3 years whereas the 60 others were ad libitum. This amounted to approximately 35% CR compared to the control group. While the initial report was brief, post hoc analyses conducted several years later [73] indicated that death rate tended to be lower in the CR group and hospital admissions were reduced in these individuals by approximately 50% (123 days for CR versus 219 day for control). 22.8.3 Unexpected CR in Biosphere 2

The unexpected low availability of food during the 2-year Biosphere 2 experiment provided a unique opportunity to observe the effects of CR in a group of nonobese humans. Biosphere 2 is an enclosed 3.15-acre ecological laboratory that houses seven ecosystems or biomes resembling the Earth: rainforest, savannah, ocean, marsh, desert, and agriculture and human/animal habitats [74]. For 2 years, eight scientists were completely isolated within this mini-world where 100% of the air and water was recycled, and all the food grown inside. Due to unforeseen problems with agriculture early on, food supply was much lower than expected. Food intake for the eight individuals was projected at around 2500 kcal/day and estimates from food records maintained by one of the biospherians suggested diets were restricted by around 750 kcal/day in each person during the ﬁrst 6 months. The 17 5% resulting weight loss was associated with many physiological, hematological, biochemical, and metabolic alterations [74, 75], consistent with calorie-restricted rodents and primates, including reductions in insulin, core temperature, and metabolic rate [30]. Furthermore, cardiovascular disease risk factors such as a 25 and 22% decrease in systolic and diastolic blood pressures, respectively, and a 30% lowering of cholesterol support CR studies in animals [54].

22.8 What is Known from Humans?

22.8.4 Randomized Controlled Trials of Prolonged CR in Humans

As for randomized controlled trials, results from a 2-year study of CR in humans are only a few years away. The National Institute on Aging is sponsoring a trial called CALERIE (Comprehensive Assessment of the Long-term Effect of Reducing Intake of Energy) that will for the ﬁrst time scientiﬁcally test the effects of 25% CR in more than 150 nonobese (22 body mass index (BMI) < 28) healthy men and women aged 21–45 years. Three clinical sites are involved in the trial; Washington University (St. Louis, MO), Tufts University (Boston, MA), and Pennington Biomedical Research Center (Baton Rouge, LA). In preliminary studies (ﬁrst phase of CALERIE), besides feasibility and safety, each clinical site tested varying hypothesis and modalities of CR in studies ranging from 6 (Pennington) to 12 months (Washington University and Tufts). The ﬁndings of these studies are starting to be published [76–86]. The Phase 1 study conducted at Pennington Biomedical Research Center involved 46 men and women, randomized to one of four treatment groups for 6 months. For the CR group, the level of restriction imposed was a 25% reduction from the daily energy requirement for weight maintenance [76]. The other groups were: (i) CR plus exercise group where the calorie deﬁcit was also set at 25% reduction from weight maintenance energy requirement, but half (12.5%) was achieved by dietary restriction and half (12.5%) from an increase in energy expenditure by structured aerobic exercise; (ii) a low-calorie diet group where participants consumed 890 kcal/day to achieve a 15% weight loss and thereafter followed a weight maintenance diet; and (iii) a healthy diet control group that followed a weight-maintaining diet based on the American Heart Association Diet, Step 1. Six months of CR induced favorable outcomes in terms of physiological, hormonal, and biochemical parameters. Importantly, the 12 participants assigned to this treatment group completed the study and reported no development of eating disorder symptoms [82] or reductions in quality of life indices [87]. After 6 months of 25% CR, the group lost 10.4 0.9% of their body mass, attributed to both a loss in fat mass (CR: 24 3%) and fat-free mass (CR: 4 1%). Central adiposity was also improved with a 27% reduction in both visceral (women: 24 4%, men: 32 6%) and subcutaneous fat depots (women: 25 2%, men: 28 7%). Interestingly, the distribution of whole-body fat and speciﬁcally within the abdominal compartment was not altered by CR [78]. Abdominal fat cell size was reduced by around 20% and the deposition of lipid in the liver was lowered by around 37%, but no change was noted in the lipid content within skeletal muscle [77]. Importantly, the reduction in weight, visceral fat, and abdominal fat cell size were associated with a 40% improvement in insulin sensitivity and reduced acute insulin response to glucose as assessed by the frequently sampled intravenous glucose tolerance test [77]. We also observed favorable changes in the lipoprotein proﬁle [88]. Triacylglycerol was reduced by 21%, high-density lipoprotein (HDL) cholesterol increased by 9%, and Factor VIIc reduced by 10%. No changes were observed in ﬁbrinogen, homocysteine, or endothelial function. Based on combined changes in lipid and blood

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pressure values, the estimated 10-year cardiovascular disease risk declined by 29% in the CR group, but as expected remained unchanged in the control group. Based on combined favorable changes in lipid and blood pressure levels, CR favorably reduces risk for cardiovascular disease [88]. With regard to longevity, two out of three biomarkers of longevity [22] were improved in the CR group after 6 months [76]. Speciﬁcally, we observed signiﬁcant reductions in both fasting insulin and core body temperature. Interestingly, in parallel with the decrease in core temperature measured by telemetry, we observed a metabolic adaptation – lowering of metabolic rate larger than expected on the basis of weight loss – associated with reduced DNA damage, probably due to lower production of ROS [76]. These ﬁndings of course echo results previously reported in nonhuman primates and rodents on CR [30] and long-lived men in the Baltimore Longitudinal Study of Aging [22]. Importantly, CR was associated with an increase in the muscle expression of genes involved in mitochondrial biogenesis and mitochondrial fusion including peroxisome proliferator-activated receptor-c coactivator (PGC)-1a, mitochondrial transcription factor A, endothelial nitric oxide, sirtuin-1, and presenillin-associated rhomboid-like protein [79]. In parallel, mitochondrial content increased by 35 5% in the CR group with no change in the control group (2 2%). However, the activity of key mitochondrial enzymes of the tricarboxylic acid cycle (citrate synthase), b-oxidation (b-HAD), and electron transport chain (cyclooxygenase II) were all unchanged. This suggests that 6 months of CR in nonobese humans was sufﬁcient to improve biomarkers of aging and supports the theory that 24-h energy expenditure is reduced beyond that expected due to reduced metabolic size. Whether the observed metabolic adaptation translates into long-term overall reduced oxidative damage remains to be determined. The increased mitochondrial content in association with a decrease in whole body DNA damage is, however, an important indication that CR improves mitochondrial function in human skeletal muscle – a factor that may decrease cellular senescence. The Phase I study at Washington University was conducted in overweight (but nonobese) men and women aged 50–60 years [80, 83–85]. Participants were randomized to one of three treatment groups for 12 months: (i) CR group where the level of dietary restriction was 20% less than baseline energy requirements for weight maintenance, (ii) exercise group that had an identical prescribed energy deﬁcit of 20% only by daily aerobic exercise, and (iii) healthy diet control group. After 1 year of CR, body mass was reduced by around 11% and participants were now considered healthy weight for height based on BMI (24 0.6 kg/m2). With regard to adiposity, whole-body fat mass was reduced (around 25%), as were abdominal visceral (37%) and subcutaneous masses [80]. Associated with these favorable changes in body composition were improvements in lipid proﬁle and cardiovascular disease risk. Total cholesterol, triglycerides, and LDL cholesterol were decreased with the 12 month intervention, and HDL cholesterol increased, but not signiﬁcantly [85]. Insulin sensitivity assessed during an oral glucose tolerance test was improved as was fasting insulin concentrations [84]. Systolic and diastolic blood pressures were also

22.9 Could CR Increase Longevity in Humans?

improved by CR. According to the Framingham algorithm for estimating 10-year Coronary Heart Disease Risk, which considers age, total cholesterol, HDL cholesterol, blood pressure, smoking, and diabetes [89], the CR group had a signiﬁcant reduction in coronary heart disease risk at month 12 when compared to the healthy living control group [85]. In summary, this 12-month trial of moderate CR in older persons indicates that age-associated changes in body composition and cardiovascular disease risk can be improved. Whether or not CR will sustain these effects over the long term and increase lifespan is not known in humans. The Phase 1 study at Tufts University was designed to test the effects of CR with two different dietary macronutrient patterns [86]. Thirty-four overweight adults completed a 30% CR regimen for 12 months. Half of the participants were randomized to consume a diet with a high glycemic load (60% carbohydrate, 20% protein, 20% fat) and the other half to a diet with a low glycemic load (40% carbohydrate, 30% fat, 30% protein). In both diet groups, body mass and percent body fat declined similarly, reaching around 8% for weight and around 16% for fat after 12 months, respectively. There were also observed improvements in cardiovascular and metabolic factors. Total cholesterol, triglycerides, and LDL cholesterol were reduced by around 5, 15, and, 7%, respectively, and fasting insulin by around 20%. Taken together, the preliminary ﬁndings from the Phase I of CALERIE indicate promising beneﬁts of CR for 6–12 months on body composition whereby total fat mass, visceral fat, and ectopic fat stores are reduced. These body composition changes are associated with improvements in plasma lipids and reductions in cardiovascular and type 2 diabetes risk. Due to the pluripotent nature of energy restriction, the exact mechanisms by which CR extends lifespan are still being investigated and will likely remain a challenge. However, controlled human trials of prolonged energy restriction, such as the multicenter CALERIE study, are transforming this challenging investigation into a modern scientiﬁc reality.

22.9 Could CR Increase Longevity in Humans?

Taken together, the information obtained from humans so far is suggestive that CR is associated with clear health beneﬁts, but we lack sufﬁcient evidence to conclude that age-related morbidities are deﬁnitely reduced and/or lifespan is extended. It is not known from the studies of CR in humans if, through measurement of known biomarkers of aging, biological age is improved and if it results in an extension of chronological age (Figure 22.2). The wealth of CR literature in rodents, however, allows us to address some important questions relating to the practicality and feasibility of CR in humans. Relevant and practical questions that need to be asked are: i) How much CR do we need to improve age-related health and possibly longevity? ii) How long do we need to sustain CR in order to obtain these beneﬁts?

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• • •

Fasting Insulin Body Temperature

• •

DHEAS ?

?

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61 20

40

60 80 20 40 60 Chronological vs. Biological Age (y)

Figure 22.2 Can CR improve biological age and extend chronological age? This figure illustrates three validated biomarkers of longevity. It is hypothesized that CR will change the trajectory of these biomarkers, and therefore improve biological age and extend chronological age. For example, the left panel shows an individual aged 75 years. With

80

prolonged CR, it is hypothesized that insulin and core temperature will be reduced and the individual although 75 will have a biological age 14 years younger. Similarly, the individual on the right at 90 years with prolonged CR will be biologically similar to an individual aged 66 years. DHEAS, dehydroepiandrosterone sulfate.

22.9.1 How Much CR?

How much CR would be necessary to derive the beneﬁts observed in mammals? According to Merry [90], analysis of 24 published studies of CR in mice or rats undergoing varying degrees of CR indicated a strong negative relationship between survival and energy intake. Across a wide range of energy intakes a dose–response relationship is evident between the degree or severity of CR and survival such that lifespan is increasingly extended with increasing degrees of CR [90]. In general, these studies indicate that the more CR (up to 55%), the better the effect on maximal lifespan. However, while a CR of 30%, which is frequently used in the rodents and is the level of restriction imposed in the nonhuman primate colonies, is probably feasible in humans, 50% CR that has been successfully implemented in rodents is not. The preliminary phase of the randomized trial CALERIE, as discussed Section 22.8, imposed a 25% CR for 6 months with alterations in biomarkers of longevity consistent with the rodent and primate studies [76]. We will know more about the longer-term feasibility of this moderate level of CR when the 2-year investigation is complete. 22.9.2 How Long is CR Required?

How long does a CR intervention need to be sustained in order to reap antiaging beneﬁts? Induction of CR prior to sexual maturation has negative consequences on

22.10 CR Mimetics

growth and reproduction, especially in females. Analysis of several studies in rodents initiating CR in early adulthood indicates that the duration of CR is an important factor in lifespan extension. The interpretation from 36 published studies of 40–50% CR indicated a positive correlation between average and maximal lifespan and the duration of the CR intervention [90]. Hence, the longer the animal is maintained on CR, the greater the rate of survival. What can be taken away from the rodent data is that CR has greater beneﬁts when it is more extreme and sustained over a long period of time. Using the prediction equations derived from the rodent data above, we estimated that a 5-year life extension could be induced by 20% CR starting at age 25 and sustained for 52 years (i.e., the life expectancy of a male in the United States). However, if a 30% CR was initiated at age 55 for the next 22 years, the gain would only be 2 months. Enhancing life expectancy and delaying age-associated morbidities is certainly appealing; however, it seems that while living in the current obesogenic environment, both environmental and social pressures would make sustained CR very difﬁcult at best. Furthermore, it is difﬁcult to predict the quality of life in people engaging in CR. Certainly there are individuals belonging to the Calorie Restriction Society (www.calorierestriction.org) who self-impose CR with the CRON (Calorie Restriction with Optimal Nutrition) diet for health and longevity. A group of 18 CRONIES (only three women) from the United States and Canada have recently been studied after 3–15 years of CR [60]. Dietary analysis indicated an energy intake around 50% less than age-matched controls. In terms of body composition, the mean BMI of the males was 19.6 1.9 kg/m2 with an extremely low percent body fat of around 7%. Atherosclerosis risk factors including total cholesterol, LDL cholesterol, HDL cholesterol, and triglycerides fell within the 50th percentile of values for people in their age group. This report provides further evidence that longer-term CR is highly effective in lowering the risk of developing coronary heart disease and other age-related comorbidities [60]. It remains to be seen if the CRONIES live longer than their age- and sex-matched counterparts or, in the interim, if the surrogate biomarkers of longevity, including reduced insulin, dehydroepiandrosterone, and core temperature, are altered in favor of increased lifespan.

22.10 CR Mimetics

Given the extent and severity of CR necessary to obtain potential beneﬁts, it is no wonder that a large amount of research in the gerontology ﬁeld (and now pharmaceutical companies) is being geared toward identifying and testing compounds that mimic CR [91, 92]. Excitement is growing for resveratrol – a compound with antioxidant properties found in red wine, in the skin of grapes and berries, and in peanuts. Already resveratrol has been shown to increase lifespan in yeast, fruit ﬂies [93], and recently by around 60% in ﬁsh [94], but now data have been emerging that resveratrol increased the survival of mice even in conjunction with a high-calorie diet [95]. Furthermore, despite weight and fat gain, the resveratrol-treated animals

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had metabolic proﬁles that paralleled that of the control group and support lifespan extension [95]. In another recent study, resveratrol was found to improve the efﬁciency of skeletal muscle metabolism by increasing mitochondrial biogenesis and function [96]. Interestingly, besides being resistant to high-fat diet-induced obesity and insulin resistance, mice fed large doses of resveratrol could run twice as long as the control mice before exhaustion. We believe that while the rodent and primate data indicate lifespan extension is possible with CR, collective analysis of these data suggests that intensity and onset of CR required to induce these effects is probably not suitable for many individuals. Epidemiological studies certainly support the notion that a reduced energy intake that is nutritionally sound improves age-associated health. If compounds of CR mimetics prove viable in humans, individuals for the most part will opt to enjoy the effects of antiaging via a pill and not CR. 22.11 Conclusions

Since the ﬁrst report of prolonged lifespan in rodents more than 70 years ago, similar observations have been reported across a wide range of species, including yeast, worms, spiders, ﬂies, ﬁsh, mice, and rats. While the effects of CR in longer-lived species remain unknown, we now have the opportunity to test whether the mechanisms discovered in small animals are present in primates and humans. However, facing an obesogenic environment, it seems very unlikely that a public health message may be launched towards reducing the amount of ingested calories. For that reason, the mechanisms leading to the retarded senescence at the molecular and physiological levels are important to understand for the successful development of CR mimetics. Such natural compounds or botanical extracts would mimic the effect of CR without depriving people from their usual energy intake. This is a ﬁeld that will expand in the next few years, and will be headed by biotechnology and pharmaceutical companies, all eager to search for small molecules mimicking the effect of CR and representing the Fountain of Youth. Acknowledgments

This review partially supported by grant U01 AG20 478 from the National Institute on Aging, NIH. L.M.R. is supported by a Neil Hamilton-Fairley Training Fellowship awarded by the National Health and Medical Research Center of Australia (ID 349 553).

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23 Peroxisome Proliferator-Activated Receptor-c: A Key Regulator of Adipose Tissue Formation, Remodeling, and Metabolism Olga Astapova and Todd Leff 23.1 Introduction

The transcription factor peroxisome proliferator-activated receptor (PPAR)-c plays a vital role in many aspects of adipose tissue biology. It is required not only for the development of adipose tissue during embryogenesis, but also for the maintenance of normal adipocyte morphology and function, and of adipose tissue architecture in the adult animal. The central importance of PPAR-c in adipose tissue development and function is demonstrated by multiple independent lines of evidence. Reduction of PPAR-c activity by gene mutation, in both humans and mice, causes alterations in adipose tissue morphology and function, and severe metabolic abnormalities that share many features with type 2 diabetes. On the other hand, activation of PPAR-c with receptor-speciﬁc agonists (e.g., the thiazolidinedione (TZD) drugs) leads to distinct adipose tissue remodeling and improvements in the metabolic abnormalities associated with type 2 diabetes. Thus, the amount of PPAR-c protein present in adipose tissue, and the level of its transcriptional activity, must be maintained within a fairly speciﬁc range for normal metabolic balance and proper adipose tissue function. Deviations from this ideal level have profound effects not only on speciﬁc cellular processes within adipocytes, but also on global (whole-body) metabolic parameters that affect the health of the organism. From this very large body of research, it is clear that a complete understanding of adipose tissue function will require an in-depth knowledge of the molecular and cellular function of PPAR-c. This chapter focuses on the speciﬁc role of PPAR-c in adipose tissue function and will not discuss the closely related family members, PPAR-a and PPAR-d, that may also play important roles in adipose tissue function. The reader is referred to one of the many excellent reviews on the biological activities of these transcription factors.

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23.2 Molecular Biology of PPAR-c 23.2.1 PPAR-c Structure and DNA Binding

Nuclear receptors are ligand-activated DNA-binding proteins that regulate transcription of target genes via recruitment of histone-modifying enzymes and altering the transcriptional activity status of the chromatin. PPAR-c is a member of type II nuclear receptors, deﬁned by dependence on heterodimerization with other members of the nuclear receptor family. In the case of PPAR-c, it dimerizes with the retinoic X receptor (RXR)-a, and the PPAR-c/RXR-a heterodimer is its functional unit for DNA binding and transcriptional activation. The type II class of nuclear receptors also includes the vitamin D and thyroid hormone receptors, among others. All nuclear receptors share structural similarities (Figure 23.1), including a DNA-binding domain comprised of two zinc ﬁnger motifs, a liganddependent transactivation domain (AF-2) in the C-terminus of the protein, and

Figure 23.1 PPAR-c gene structure and RNA expression: four promoters in the human PPARc gene give rise to four unique mRNA species, PPAR-c1–4. Of these RNAs 1, 3 and 4 are translated into the identical protein, PPAR-c1, containing exons 1–6. PPAR-c2 mRNA is

translated into the PPAR-c2 protein, identical to PPAR-c1 except for an additional 28-amino-acid sequence at the N-terminus which is encoded by exon B. Domain structure of the PPAR-c peptide is shown in the bottom panel. Adapted from [67].

23.2 Molecular Biology of PPAR-c

ligand-independent transactivation domain (AF-1) at the N-terminus. The ligandbinding domain of PPAR-c contains coregulator interaction interfaces and also forms extensive interactions with the ligand-binding domain of RXR-a [1]. The function of the AF-1 domain is not well understood, but it contains interaction sites for some coregulators [2]. PPAR-c is expressed as several isoforms as the result of differential splicing and promoter use. The two most abundant isoforms, PPAR-c1 and PPAR-c2, are derived from differentially spliced RNAs initiated from distinct promoters and generate two PPAR-c proteins that are identical with the exception of a 28-amino-acid N-terminal sequence present in PPAR-c2, but not the shorter PPAR-c1. While the activities of these two isoforms appear to be quite similar with respect to DNA binding and activation of gene transcription in vitro, they display distinct tissue distributions in vivo. PPAR-c1 is expressed in many tissues, including adipose tissue, liver, kidney, macrophages, muscle, and the intestine, while PPAR-c2 expression is more strongly restricted to adipose tissue. The functional signiﬁcance of this tissue speciﬁcity remains poorly understood, but it is likely an important determinant of the many nonadipose tissue functions of PPAR-c [3]. PPAR-c/RXR-a heterodimers bind DNA at deﬁned sequences referred to as PPAR response elements (PPRE)s, which are located in the regulatory regions of PPAR-c target genes. The consensus PPRE is a direct repeat of the core sequence AGGTCA with one spacing nucleotide – this conﬁguration of a bipartite response element is referred to as a direct repeat (DR)-1. Unlike the steroid hormone receptors that are localized to the cytoplasm until ligand binding, class II nuclear receptors are believed to be bound to DNA at their cognate response elements, even in the inactive unliganded form, and this appears to be the case for PPAR-c. The structure of intact PPAR-c/RXR-a heterodimer bound to the consensus PPRE was recently resolved by X-ray crystallography [1], providing insights into the speciﬁc interactions between the receptor and the DNA molecule, as well as between PPAR-c and RXR-a. The C-terminal extension of the DNA-binding domain of PPAR-c makes extensive contact with the DNA minor groove just outside of the 50 end of the PPRE [1], suggesting a role for these ﬂanking sequences in DNA binding of PPAR-c. These ﬂanking sequences may also be involved in another aspect of PPAR biology that is poorly understood. The three PPAR family members (PPAR-c, -a, and -d) all recognize canonical PPRE sequences, but regulate overlapping but distinct sets of PPRE-containing target genes [4]. The ability of these closely related receptors to differentially regulate certain target genes may be due to their interaction with these ﬂanking sequences. Some well-characterized PPAR-c target genes have a conserved AAACT sequence directly upstream of the DR-1 in their regulatory regions, suggesting a role for these ﬂanking sequences in DNA sequence recognition and possibly in determining which PPRE-containing genes are activated by PPAR-c and to what degree [5]. However, more recent genome-wide studies of PPAR/RXR DNA binding indicate that the 50 -ﬂanking nucleotides of DR-1 sequences are not signiﬁcantly conserved and do not contribute appreciably to the prediction of PPAR-c-binding sites or help distinguish binding sites of different PPAR subtypes [6]. It is possible that the C-terminal extension of PPAR-c interacts only with

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the minor groove of the DNA molecule in a sequence-independent fashion [1] and functions mainly to orient the heterodimer in its correct polarity on the DR-1 sequence. Several recent genome-wide studies have characterized the global set of PPAR-cbinding sites using chromatin immunoprecipitation and computer-based (in silico) methods [7–10]. These studies challenge the traditional view of PPREs as localized to the conserved proximal promoter regions of genes (i.e., within 5 kb upstream of the transcription start site) – a dogma that has perhaps hindered the discovery of bona ﬁde PPREs, as investigators limited their region of investigation to the proximal promoters. In fact, only a minority (fewer than 10%) of all binding sites identiﬁed in these recent studies are located in the proximal promoter region. Rather, most PPARc/RXR-a binding sites are found within the gene-coding regions, in both introns and exons. In addition, a high percentage of PPREs are located within the extended promoter regions (up to 100 kb upstream of the transcription start site) or in the distal (up to 100 kb) 30 -untranslated regions. Many sites are also located in the intragenic regions or gene deserts (more than 100 kb from the nearest gene coding region). It is still unclear whether there exists a relationship between the transcriptional activity of the PPAR-c/RXR-a complex and the proximity of its binding site relative to the transcription start site of the target gene. Limited evidence suggests that PPAR-c DNA binding and transcription induction are biased toward the PPREs located within 10 kb upstream of the transcription start site [9, 10], although it seems likely that the more distal binding sites also play an important role in the regulation by PPAR-c of many target genes. 23.2.2 Transcriptional Regulation by PPAR-c

The traditional view of how transcription factors function to activate expression of target genes is founded on the assumption that they form stable, long-lasting protein/DNA complexes that direct multiple rounds of gene transcription by RNA polymerase. Recently, a more dynamic model has been proposed in which gene regulation is accomplished through a rapid transcriptional activation cycle initiated by binding of an agonist (in the case of the nuclear receptors) and terminated by the ejection of the transcription factor from the promoter [11]. For PPAR-c-induced genes, this cycle would be initiated by the binding of a ligand to the receptor that is already present on the PPREs of target genes (Figure 23.2). Ligand binding alters the three-dimensional structure or conformation of the DNA-bound receptor that leads to a change in the set of auxiliary proteins or coregulators bound to the receptor (described in more detail below). These coregulators convert the local chromatin structure to a more open, transcriptionally active, conﬁguration, and promote the binding of RNA polymerase II and the initiation of transcription. Once transcription has been initiated, the liganded DNA-bound PPAR-c is subject to proteolytic degradation and removal from the promoter, restoring the gene to its basal, unstimulated state [12, 13].

23.2 Molecular Biology of PPAR-c

Figure 23.2 Transcriptional activation by PPAR-c: PPAR-c binds PPREs as a heterodimer with RXR. In the unliganded state (top), PPAR-c interacts with corepressor molecules capable of modulating chromatin structure via HDAC activity, leading to a closed, or inactive, chromatin configurations and reduced gene transcription. Ligand binding induces a conformational change in PPAR-c that favors release of the corepressors and recruitment of coactivators (bottom), leading to acetylation of

histones and the conversion of chromatin to an open configuration that is more accessible to the transcriptional machinery. The presence of coactivators such as TRAP220 also leads to the recruitment of the mediator complex, which acts as a bridge between the transcription factors and RNA polymerase (POL II), and is required for the assembly of a functional transcription complex and the initiation of transcription.

The conformation change in PPAR-c protein structure induced by ligand binding inﬂuences the interactions between the receptor and the set of coregulator proteins that mediate its transcriptional activity (for a detailed review of nuclear receptor coregulator interactions, see [14]). In the unliganded form, the DNA-bound receptor is found in a complex with corepressor proteins such as receptor-interacting protein-140 (RIP140) [15], nuclear receptor corepressor-1 (NCOR1) [16], and silencing mediator of retinoic acid and thyroid hormone receptor (SMRT) [17] that block the transcriptional activity of the receptor and maintain the local chromatin structure in an inactive conﬁguration via recruitment of histone deacetylases (HDACs) [18, 19].

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Ligand-bound PPAR-c assumes a conformation that reduces its afﬁnity for corepressors and promotes the recruitment of coactivator proteins such as CBP/p300 [20], SRC1/p160 [21], and thyroid hormone receptor-associated protein complex 220 kDa component (TRAP220) [22]. This results in a remodeling of the chromatin structure and docking of the RNA polymerase II at the promoters of PPAR-c target genes to initiate gene transcription. Recruitment occurs via interaction of the AF-2 domain of PPAR-c with LXXLL motifs of the coactivator molecules; histone acetyltransferases and histone methyltransferases are then directed to the chromatin [23]. In addition, a large multiprotein complex termed Mediator associates with nuclear receptors via the LXXLL motif in TRAP220 and is associated with docking of RNA polymerase II to the promoter, possibly via direct interaction between the C-terminal domain of RNA polymerase II and members of the mediator complex [24]. While many of these coactivators mediate transcriptional activity of nuclear receptors in general, PPAR-c coactivator-1a(PGC-1a) is more speciﬁc for the PPARs, and mediates the upregulation of functionally related genes that participate in mitochondrial biogenesis and oxidative phosphorylation in muscle, brown fat, and other tissues [25, 26]. In considering the biological function and signiﬁcance of the PPAR-c regulatory system, it is important to recognize that while PPAR-c activation induces a highly orchestrated transcriptional program involving hundreds or even thousands of PPARc target genes, there is a great deal of variation in the degree and timing of the response among the individual target genes. Many factors contribute to this variation in how individual genes respond to what is essentially the same stimulus. These include the interaction of PPAR-c with PPREs that vary in their DNA sequences, the presence of distinct sets of PPAR-c coactivators and corepressors in different tissues and different physiological conditions, as well as the interaction of PPAR-c with variable sets of transcription factors depending on the promoter environment. For instance, the traditional view of ligand binding as the sole determining factor of coregulator composition is challenged by recent reports that some PPAR-c target genes are transcribed in a ligand-independent manner [2], and some coactivators and corepressors bind PPAR-c regardless of the presence of ligand [25]. Corepressor RIP140 has been shown to interact with ligand-bound nuclear receptors and actively repress gene transcription via HDAC recruitment as well as HDAC-independent mechanisms [19, 27]. In addition, target gene identity may contribute to recruitment of speciﬁc coactivators via the A/B domain of PPAR-c [2]. The regulation of PPAR-c activity is also affected by its interaction with other DNA-bound transcription factors that may be present on a subset of target genes. One well characterized example of this is the interaction of PPAR-c with the adipogenic transcription factor CCAAT/enhancer-binding protein (C/EBP)-a. DNA binding of C/EBP-a is spatially and temporally associated with PPAR-c binding, and the two proteins cooperate in inducing the adipogenic gene transcription program [7, 8, 28]. Therefore, to fully appreciate the biological function of PPAR-c and to take full advantage of its therapeutic potential, it will be necessary to decipher the details of each of these highly complex processes at the molecular level.

23.3 PPAR-c is a Master Regulator of Adipose Tissue Development

23.3 PPAR-c is a Master Regulator of Adipose Tissue Development

One of the primary functions of PPAR-c is to serve as a master regulator of the cellular differentiation process that converts ﬁbroblast-like preadipocytes into terminally differentiated lipid laden adipocytes. In vivo, this means that PPAR-c is absolutely required for the formation of adipose tissue. As will be discussed in more detail below, animals that are missing PPAR-c do not have any adipose tissue and die immediately after birth. Our knowledge of how exactly PPAR-c functions to regulate adipogenesis comes mainly from studies in tissue culture model systems in which ﬁbroblastic cell lines are stimulated with a hormonal cocktail to differentiate into cells that resemble in many ways native adipocytes found in fat tissue. This aspect of PPAR-c biology is summarized in the following sections. The reader is referred to the many excellent reviews of the gene regulatory networks that control adipogenesis for a more detailed treatment of the subject (e.g., see [29–31]). 23.3.1 Role of PPAR-c in Adipogenesis – Cell Culture Studies

In vitro studies of adipogenesis commonly involve culture of immortalized 3T3-L1 mouse preadipocytes, which can be induced to differentiate into adipocytes by treatment with a cocktail of dexamethasone, insulin, and methylisobutylxanthine. Immediately following treatment of preadipocytes with the adipogenic cocktail, the cells undergo a burst of cell division referred to as mitotic clonal expansion [32, 33]. Although poorly understood, it appears that one of the proadipogenic functions of PPAR-c is to suppress the expression of genes involved in this cell cycle progression. These include actin organization genes, RNA-splicing enzymes, translation initiation and elongation factors, and cyclins. The decline in expression of these genes is consistent with decreased cell division and intracellular reorganization during early adipogenesis. Following the clonal expansion phase, two transcription factors in the C/EBP family, C/EBP-b and -d, are activated and function to stimulate the expression of PPAR-c and C/EBP-a, and probably other adipogenic genes. Downstream genes characteristic of terminally differentiated adipocytes are induced by PPAR-c/RXR-a heterodimers that bind to PPREs associated with thousands of target genes, frequently at locations that are in close proximity to C/EBPa response elements [8]. The coordinated action of PPAR-c and C/EBP drives the expression of the large set of genes that characterize the fully differentiated adipocyte [7–9]. Later in the differentiation pathway, PPAR-c-dependent induction of metabolic genes takes place, with enrichment of genes involved in glucose and lipid handling, such as lipoprotein lipase, perilipin, phosphoglycerate kinase, and hexokinase, reﬂecting the need to maintain adipocyte functions of lipogenesis, lipid storage, and metabolism. Interestingly, during adipogenesis PPAR-c occupancy is signiﬁcantly higher in the vicinity of upregulated genes compared to the set of downregulated genes [7, 8], suggesting that downregulated genes may not be direct transcriptional targets of PPAR-c, but may be regulated by other transcription factors

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induced by PPAR-c. There are no comparable studies to date of PPAR-c-binding sites during maintenance of fully differentiated adipose tissue. 23.3.2 PPAR-c is Required for Adipose Tissue Development In Vivo

As already mentioned, the whole-body PPAR-c gene knockout is lethal. The small number of animals that were born with this mutation were completely devoid of adipose tissue and died immediately after birth [34]. Although this clearly demonstrated a crucial role for PPAR-c in adipose tissue development, it did not provide a useful model to study the mechanisms of PPAR-c function. Of greater utility in this regard were several different mouse models of adipose-speciﬁc ablation of PPAR-c. Despite distinct differences in severity of the phenotype and in the physiological response to adipose-speciﬁc PPAR-c ablation, all of these animal lines share the characteristics of lipodystrophy and hyperlipidemia. This again highlights the fundamental importance of PPAR-c in maintaining the ability of adipose tissue to function as a lipid storage depot. Knockout of PPAR-c2 by homologous recombination, generating a mainly adipose-speciﬁc PPAR-c loss, resulted in severe lipodystrophy, growth retardation, and death of young pups [35, 36]. Surviving animals overcame the growth retardation but the fat pads remained small due to decreased number of adipocytes, demonstrating the necessity of the PPAR-c2 isoform for adipose tissue differentiation. Ablation of adipose PPAR-c (both isoforms) immediately following adipocyte differentiation using cre recombinase [37, 38], or in adult mice using an inducible cre recombinase [39], also produced a lipodystrophic phenotype with compensatory hypertrophy of surviving adipocytes after the majority of existing adipocytes underwent apoptosis. Together these studies provide unequivocal evidence of a central role for PPAR-c in both differentiation of adipocytes as well as for post-differentiation maintenance of adipose tissue. The lipodystrophic remodeling of adipose tissue caused by PPAR-c ablation was accompanied by increased plasma free fatty acids due at least in part to resistance of the PPAR-c-deﬁcient adipose tissue to insulin-mediated inhibition of lipolysis. Adipokine secretion was decreased in all animals; glycemia and systemic insulin sensitivity were consistently normal on standard diet. However, the mutation rendered mice in some studies [37] more susceptible to high fat diet-induced insulin resistance, while similar mice in other studies [38] were protected from diet-induced insulin resistance. These differences may be accounted for by differential contribution of the liver to systemic insulin sensitivity. Liver PPAR-c expression was upregulated in the latter study as a compensatory response to adipose tissue insulin resistance, leading to increased hepatic glucose disposal. In the former study, on the other hand, the lipid overload caused hepatic insulin resistance, although the skeletal muscle was able to compensate for this under normal feeding conditions. Similarly, the metabolic derangements brought on by lipodystrophy in the Koutnikova et al. study [35] were compensated by increased glucose uptake into the muscle. Regardless of whole-body phenotypic differences, all studies point to the crucial role of

23.4 Metabolic Functions of PPAR-c

PPAR-c in adipose tissue lipid storage and together form an excellent illustration of the complexity of the multiorgan homeostatic systems that regulate metabolic balance. When gene expression patterns were examined in these PPAR-c adipose-speciﬁc knockout models, decreased expression of lipogenic and adipogenic markers (aP2, C/EBP-a, lipoprotein lipase) was uniformly observed. Notably, both upregulated and downregulated expression of known PPAR-c target genes was observed after PPAR-c loss, consistent with the recent report that PPAR-c can both induce and suppress expression of speciﬁc target genes [25]. Moreover, these changes in expression were tissue-speciﬁc, highlighting the inﬂuence of cellular environment on PPAR-c activity.

23.4 Metabolic Functions of PPAR-c

Agonist stimulation of PPAR-c activity in vivo leads to metabolic changes in adipose tissue that result in reduced plasma free fatty acids and triglycerides, improved glucose tolerance and insulin sensitivity, and suppression of inﬂammatory state, among other effects in a wide variety of tissues. The exact mechanism behind insulin sensitization by PPAR-c is unknown, but the reduction in blood lipid levels is believed to reverse lipotoxicity – the aberrant accumulation of lipid in nonadipose tissues such as liver and muscle. Within adipose tissue, PPAR-c stimulation promotes tissue remodeling, leading to greater number of smaller adipocytes. These smaller adipocytes have an increased lipid storage capacity and an altered pattern of adipokine secretion characterized by increased production of adiponectin, which is antidiabetic, and reduced expression of proinﬂammatory cytokines associated with insulin resistance. It is known that different adipose tissue depots have distinct endocrine and metabolic properties. Visceral fat pads, represented most notably by omental, mesenteric, and retroperitoneal depots, secrete more inﬂammatory adipokines, including cytokines interleukin (IL)-6, IL-8, tumor necrosis factor (TNF)-a, and plasminogen activator inhibitor (PAI)-1. However, it is not clear whether these factors originate from adipocytes or other cells resident within the adipose depot, such as macrophages or stromal cells [40]. Many other adipokines such as adiponectin and leptin are preferentially released from subcutaneous adipocytes. In addition, visceral adipocytes are more sensitive to adrenergic stimulation of lipolysis and less sensitive to insulin suppression of lipolysis than subcutaneous adipocytes, while basal lipolysis is higher in the latter. Although PPAR-c is expressed in both visceral and subcutaneous fat depots, PPAR-c activity induced by systemic agonist administration is depot-speciﬁc. PPAR-c activation in subcutaneous adipose tissue leads to recruitment and differentiation of new adipocytes resulting in a smaller cell size in this depot. This remodeling of subcutaneous adipose tissue is accompanied by enhanced anabolic capacity, namely increased fatty acid uptake and esteriﬁcation in this tissue. Induced

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expression of known PPAR-c target genes lipoprotein lipase, adipocyte fatty acidbinding protein (aP2), and diacylglycerol acetyltransferase is in line with the observed changes in lipid handling in subcutaneous fat. This is in distinct contrast to the kinds of changes observed in visceral fat upon PPAR-c activation. Visceral adipocytes also become smaller in response to PPAR-c activation, but rather than cell recruitment, the decrease in cell size is mediated by increased lipolysis, fatty acid oxidation, and thermogenic energy expenditure in this tissue – changes that are reﬂected in the observed changes in gene expression, namely the induction of pyruvate dehydrogenase kinase, carnitine palmitoyl transferase, and PGC-1 [41]. Similarly, glucose uptake and utilization are increased in subcutaneous but not in visceral fat in this animal model [42]. Adding to the complexity of depot speciﬁc function is recent ex vivo evidence that the subcutaneous adipose tissue depot can be divided into functionally distinct subcompartments – superﬁcial and deep. Deep subcutaneous depots fall functionally between the visceral and the superﬁcial subcutaneous depots in terms of TZD-induced differentiation, gene expression, and adipokine secretion differences [43]. Overall, fatty acid uptake and re-esteriﬁcation compensate for the increased lipolytic release of fatty acids into the blood stream, leading to the decrease in lipemia observed with PPAR-c agonist treatment. Agonist-induced glucose disposal into subcutaneous fat is also consistent with the expected systemic insulin sensitization. The fact that PPAR-c activation has such distinct metabolic and gene expression effects in different adipose beds provides further evidence for the importance of tissue-speciﬁc factors in modulation of PPAR-c activity.

23.5 White versus Brown Fat-Specific Functions of PPAR-c

Rodents contain two kinds of fat tissue – white adipose tissue (WAT), which is the fat storage tissue discussed above, and brown adipose tissue (BAT), which is a highly oxidative tissue used primarily for heat generation. PPAR-c is expressed in brown fat and is essential for normal function of brown adipocytes, as evidenced by atrophy of brown fat pads in the PPAR-c knockout mice described above and decreased proliferative potential of brown preadipocytes deprived of PPAR-c in vitro [44]. Interestingly, C/EBP-a, the known transcriptional partner of PPAR-c in directing white adipocyte differentiation, is not required for brown adipocyte differentiation. C/EBP-b, on the other hand, induces PGC-1a expression in brown but not white adipocytes and shifts the differentiation path of preadipocytes towards the brown fat phenotype [45]. Possibly, the C/EBP transcription factors determine the speciﬁcity of adipocyte type when partnering with PPAR-c on speciﬁc gene promoters during differentiation. Until recently it was believed that human adults, contrary to their murine counterparts, lacked brown fat; however, studies have recently pointed to a possible existence of BAT depots in speciﬁc anatomic locations of the adult human body [46–48]. These

23.5 White versus Brown Fat-Specific Functions of PPAR-c

ﬁndings, although preliminary, reveal a novel area of potential PPAR-c activity in the human, and in light of the metabolic differences between white and brown adipocytes, careful study will be required to understand the role of PPAR-c and the effects of TZDs in brown fat. Given the diverse effects of PPAR-c activation on the metabolism of different WAT compartments, one may expect such tissue speciﬁcity to be extended onto brown fat as well. Indeed, recently published work indicates that TZD induction of PPAR-c in vivo does not result in the same changes in BAT as it does in either of the white fat depots [42]. Thus, TZD treatment of adult mice induced lowdensity lipoprotein lipid uptake and recruitment of new adipocytes in WAT as well as BAT. On the other hand, TZD-induced glucose uptake into brown adipocytes was reduced per cell, such that whole-tissue glucose disposal was unchanged in brown fat. Metabolically, PPAR-c induction in BAT stimulates glycogenolysis and the conversion of intracellular glucose into glycerol and triglycerides, suggesting a switch from glycogen to triglycerides as the primary energy storage mode. With regard to gene expression in these two types of fat tissue, PPAR-c upregulates the expression of the BAT-speciﬁc gene uncoupling protein (UCP)-1 [41, 44, 49–51] – a mitochondrial inner membrane protein necessary for heat generation in nonshivering thermogenesis. This is likely accomplished through PPAR-c binding to a well-characterized PPRE in the enhancer region of the UCP-1 gene. In line with this, TZD treatment of brown adipocytes increases their thermogenic capacity upon sympathetic stimulation [49], and the interaction between TZDs and norepinephrine in activating the thermogenic process has been described as synergistic [52], possibly via increased fatty acid storage as described above, as well as enhanced UCP-1 expression. However, the physiologic relevance of these ﬁndings is still not clear, as the relative contribution of other PPAR species to the PPRE-dependent UCP-1 transcription is unknown nor has the necessity of the PPRE for UCP-1 gene expression been demonstrated. Speciﬁcally, PPAR-a, the predominant PPAR species in differentiated brown adipocytes, also induces UCP-1 expression in BAT. In contrast, PPAR-c induces some UCP-1 expression in white adipocytes and thus the tissue speciﬁcity of these results should be assessed carefully. Finally, exogenous stimulation of PPAR-c activity does not always mimic its activity state in the in vivo environment. Important questions persist regarding the activity of PPAR-c in BAT, such as whether it regulates the same set of genes in these depots, and whether PPAR-c plays a role in guiding the path of adipocyte differentiation between the brown and the white phenotypes. PR domain-containing protein 16 (PRDM16), a zinc-ﬁnger DNA-binding transcription factor, has been identiﬁed as the master regulatory protein that turns on the brown switch in differentiating adipocytes. Some studies suggest that PPAR-c activation in white adipocytes promotes a brown phenotype [40], evidenced by UCP-1 expression, but no interaction between PRDM16 and PPAR-c during adipogenesis has been reported. Gene expression proﬁling studies in BAT will provide some answers as to the role of PPAR-c in brown fat gene regulation.

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23.6 PPAR-c Function in Adipose Tissue Maintenance and Remodeling

Unlike the highly detailed understanding of PPAR-cs molecular and cellular functions, the speciﬁc mechanisms that govern PPAR-cs effects on adipose tissue morphology and architecture are not very well understood. As described in the preceding sections, much of our knowledge of PPAR-c biology has been generated using an excellent well-developed set of PPAR-c-speciﬁc molecular and pharmacological tools. These tools have also been valuable in exploring the role of PPAR-c in the creation and maintenance of the adipose tissue architecture, and its remodeling that occurs in response to a variety of metabolic and hormonal stresses. In spite of our imperfect understanding of speciﬁc mechanisms, it is quite clear that PPAR-c plays a major role in this important aspect of adipose tissue function. For example, we know that generally, reduction of PPAR-c activity by naturally occurring or experimentally engineered mutations in the PPAR-c gene leads to reduced adipose mass and, in some cases, the complete loss of functional adipose tissue. In addition, we know that activation of PPAR-c with pharmacological agents leads to a major change in adipose tissue architecture, characterized by increased numbers of smaller adipocytes. These changes in adipose tissue architecture correlate with speciﬁc effects on whole-body metabolic parameters such as glucose homeostasis and insulin sensitivity, and illustrate again the close linkage between adipose tissue function and whole-body metabolic balance. The speciﬁc effects of the experimentally reduced adipose PPAR-c expression in mice are described above and the effects of the human PPAR-c mutations on adipose tissue function are presented in detail elsewhere in this volume. The following is a discussion of how activation of PPAR-c by the antidiabetic TZDs leads to speciﬁc changes in adipose tissue morphology and architecture, and how these changes go on to alter whole-body metabolic balance. TZDs are synthetic PPAR-c ligands that are currently used in clinical practice as insulin-sensitizing agents. These drugs were discovered two decades ago as part of a broad search for chemicals that lower blood glucose in animal studies, and their recognition as PPAR-c agonists generated the extensive body of information on this nuclear receptor and its role in adipose tissue physiology that we now have. The glucose-lowering effects of TZDs, most commonly rosiglitazone and pioglitazone, in type 2 diabetic patients are well-documented in numerous clinical studies such as the PROactive, Diabetes Control and Complications Trial, and United Kingdom Prospective Diabetes Study trials as robust reductions in glycosylated hemoglobin and fasting blood glucose [53–55]. However, the side-effects associated with long-term TZD therapy – plasma volume expansion, weight gain, and decreased bone density due to PPAR-c activity in the bone marrow – deter many physicians from prescribing these agents as ﬁrst-line therapy and relegate TZDs to be used most commonly as add-on or second-line therapy in diabetics when glycemia is poorly controlled with ﬁrst-line treatments such as metformin, sulfonylureas, and insulin. In addition, the effect of TZD treatment on the development of diabetes-related cardiovascular complications is complex and requires further investigation.

23.6 PPAR-c Function in Adipose Tissue Maintenance and Remodeling

TZDs reduce hyperglycemia through improved systemic insulin sensitivity, in large part via PPAR-c activation in adipose tissue. Activation of PPAR-c in adipose tissue of diabetic patients leads to proliferation and remodeling of some fat depots, and an alteration in metabolic properties and adipokine secretory proﬁles of adipocytes [56–58]. The proliferative effects of PPAR-c are selective to subcutaneous adipose compartments, while the visceral depots, such as the omental and mesenteric abdominal fat pads, are resistant to TZD-induced differentiation. Anatomic organization of adipose tissue is related to its metabolic and secretory activity, and to cardiovascular risk and metabolic syndrome, with intra-abdominal fat pads contributing more to the development of these diseases than the subcutaneous depots. Decreasing the ratio of abdominal to subcutaneous adipose tissue is, in part, how TZDs improve the metabolic proﬁle [59]. An additional therapeutically beneﬁcial effect of TZDs on adipose tissue may be the PPAR-c-dependent proliferation that has been observed in some adipose beds. This proliferation involves recruitment or differentiation of new adipocytes, reducing the average adipocyte size [58, 60]. Since smaller adipocytes are more insulin-sensitive, this outcome of TZD treatment contributes to healthier adipose tissue by reducing lipolysis and circulating free fatty acids. Reduced delivery of fatty acids to nonadipose tissue will improve systemic insulin resistance by reducing ectopic lipid accumulation in the liver and skeletal muscle. Inappropriate lipid accumulation in nonadipose tissue, a phenomenon termed lipotoxicity [61], is thought to be one of the major causes of insulin resistance in muscle and liver [62]. The expansion of subcutaneous fat pads contributes to the weight gain observed in most patients on TZD monotherapy. However, contrary to the well-known association between body weight and the metabolic syndrome, this increase in weight is associated with improved insulin sensitivity and reduced blood glucose, and thus can be used as a measure of TZD responsiveness. Finally, TZD administration, independently of its effect on adipose tissue distribution, results in increased secretion of the antidiabetic adipokine adiponectin, and decreased secretion of the prodiabetic adipokine resistin and the cytokines IL-6, PAI-1, and TNF-a, leading to improved insulin sensitivity [40]. The effects of PPAR-c agonists on cardiovascular complications associated with diabetes are less clear than their glucose-lowering activity. Since these secondary complications are a major cause of death in these patients, and especially in light of the recently published results of the Action to Control Cardiovascular Risk in Diabetes trial demonstrating no improvement in survival of diabetic patients after intensive glucose-lowering therapy [63], it is important for clinicians to understand the cardiovascular outcomes of TZD treatment. In the PROactive study, pioglitazone offered modest protection of diabetic patients from serious cardiovascular events such as myocardial infarction and stroke [64]. However, this study, as well as a metaanalysis of seven randomized control TZD studies [65], found a signiﬁcant increase in congestive heart failure in patients treated with TZDs (although the risk of death from heart failure was not affected). Peripheral edema is a well-documented side effect of TZD treatment and probably contributes to the observed increase in congestive heart failure incidence. As a result of these studies, TZDs are now contraindicated in type 2 diabetic patients with congestive heart failure and are

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used cautiously in patients with macrovascular complications of diabetes. In other patients the glucose- and free fatty acid-lowering beneﬁts of TZD treatment are weighed against the risks of side-effects mentioned above, and in type 2 diabetic patients, TZDs are commonly prescribed in combination with ﬁrst-line drugs. In particular, coadministration of TZDs with metformin may eliminate TZD-induced weight gain [66].

23.7 Conclusions

It is now clearly understood that PPAR-c plays a crucial role in nearly all aspects of adipose tissue function, controlling not only the development of the tissue from progenitor cells and the architecture of the tissue in adult animals, but also many aspects of its metabolic function. Alterations in PPAR-c activity, either generated by genetic mutation or by pharmacological manipulation, have profound effects on adipose tissue structure and function, and on the metabolic health of the animal. Although the last decade has seen enormous growth in our understanding of PPAR-c function at all levels of biological organization, there is clearly much more to be learned about this fascinating protein and the role it plays in adipose tissue function in health and disease.

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24 Early-Life Programming of Adipogenesis and Adiposity Roselle L. Cripps and Susan E. Ozanne 24.1 Introduction

The current global epidemic of obesity is showing no signs of abating. This poses many problems as obesity is a key risk factor for many other diseases such as the metabolic syndrome [1], hypertension [2], and some forms of cancer, including breast [3] and prostate cancer [4]. Obesity, therefore, has a striking effect on both morbidity and mortality as well as being a signiﬁcant drain on healthcare resources. In the United States, where the prevalence of adult obesity has doubled in recent decades, reaching 32.2% in the 2003–2004 National Health and Nutrition Examination Survey [5], there has been signiﬁcant economic impact on the healthcare system with costs estimated at $78.5 billion in 1998 [6]. This is mirrored in other countries, including the United Kingdom, where predictions for 2010 indicate that one in three adults will be obese [7]. It is not just adult obesity that is increasing worldwide, but also childhood obesity. This too has striking effects on later disease risk and tracks with both adult obesity and increased adult mortality [8], and also has signiﬁcant negative effects on emotional development [9, 10]. The rapid shift towards an overweight/obese society indicates that a change in genes cannot be the cause, but highlights environmental or epigenetic causes, or perhaps interactions between environment and genes. There is increasing evidence that an increased susceptibility to developing overweight or obesity may begin in utero and early life (reviewed in [11]). As the development of obesity involves both adipocyte hypertrophy and hyperplasia, there is increasing interest into the effect of early-life experiences on the adipogenesis that drives adipocyte hyperplasia leading to increased fat mass. Adipocytes are no longer viewed as simple lipid stores, but act as heterogeneous endocrine tissue secreting a range of adipocytokines, such as leptin, that are vital in maintaining energy balance (reviewed in [12]).

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24.2 Theories for the Developmental Origins of Obesity

The link between maternal nutrition and nutrient supply to the fetus and later disease risk is now well established, with low-birth-weight individuals having increased susceptibility to hypertension [13], cardiovascular disease [14], the metabolic syndrome [15], and type 2 diabetes [16]. The thrifty phenotype hypothesis was proposed by Hales and Barker in 1992 to explain this link [17]. The hypothesis states that the fetus makes adaptations in the face of suboptimal conditions in utero to improve both immediate survival and chance of reaching reproductive age in postnatal conditions similar to those seen in utero [17]. These include brain sparing at the expense of other tissues, such as the endocrine pancreas, and potentially alterations in metabolism leading to an increased propensity to store nutrients as fat. If the postnatal conditions are suboptimal, then the adaptations are advantageous; however, if the postnatal environment has adequate or abundant nutrition, then the programming in utero clashes with this postnatal environment, and so obesity and metabolic diseases including the metabolic syndrome are proposed to be the result (Figure 24.1) [17]. In this case the fetus predictive adaptive responses become maladaptive [18].

24.3 Evidence for the Developmental Origins of Obesity

There is increasing evidence supporting the developmental origins of obesity. This has come both from epidemiological studies in humans and more mechanistic data from animal models.

Figure 24.1 Schematic showing the thrifty phenotype hypothesis when extended to adipocyte generation and function. Adapted from [17].

24.3 Evidence for the Developmental Origins of Obesity

24.3.1 Data from Humans

There is a U-shaped curve relating birth weight to later obesity risk [19] with both lowand high-birth-weight individuals at increased risk of developing adult obesity. The earliest work linking early-life nutrition and later obesity risk was carried out looking at individuals who where in utero during the Dutch Hunger Winter of 1944–1945. During this short deﬁned period of World War II, daily rations were reduced to 400–800 kcal per person in a region of the Western Netherlands. Individuals that experienced famine in early gestation were at increased risk of developing obesity, whereas those who experienced famine in late gestation and in early neonatal life were at decreased risk of developing obesity [20]. This shows that experiences in both gestation and early life are important in determining later obesity risk. This work has been built on studying individuals who were part of the Hertfordshire cohort and experienced more normal nutritional experiences while in utero. In this group, men of a low birth weight have increased adiposity compared to normal-birth-weight men [21]. To remove confounding variables such as maternal factors and genetic variability, studies in monozygotic twins can be used. A study looking at male adult twins showed that although the heavier twin at birth was also heavier in adulthood, he had a lower waist-to-hip ratio, less subcutaneous fat, and more lean mass that the twin that was lighter at birth [22]. The results from studies of individuals affected by the Dutch Hunger Winter highlight the different consequences of the same nutritional experiences in gestation and early life. Those that are born small and then undergo rapid catch-up growth postnatally are at increased risk of developing later obesity [23]. Indeed, the rate of growth during the ﬁrst week of life alone has been shown to affect later obesity risk with rapid growth in this period increasing later obesity susceptibility in adulthood [24, 25]. Randomly controlled trials of neonatal feeding regimes for premature infants in controlled hospital environments have evaluated breast milk, formula milk, or enriched formula milk. These provide differing amounts of energy, fat, and protein to the infant, and growth rates are affected accordingly with breast-fed infants growing slower than formula-fed infants. When these infants are followed into adolescence, the breast-fed infants who grew slower during this early period had reduced cardiovascular disease risk factors [26] and reduced risk of developing insulin resistance [27]. Recent meta-analyses have shown that breast-fed infants are protected against the later development of obesity when compared to formula-fed infants [28]. Maternal obesity prior to pregnancy has been linked to being overweight in childhood [29]. Studies of siblings who were discordant for maternal diabetes from the Pima Indian population have shown that maternal diabetes when experienced in utero predisposes offspring to develop a obese, diabetic phenotype [30]. 24.3.2 Data from Animal Models

A variety of animal models have been employed to try and elucidate the mechanisms linking early-life experiences to increased later disease risk, including increased

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obesity susceptibility. These models have included maternal nutrient restriction, both a global reduction in calories and a reduction in macronutrients such as protein, maternal high-fat feeding, experimental models of maternal diabetes, experimental models of placental insufﬁciency, exposure to exogenous glucocorticoids both via the mother or directly to the fetus, and altering litter size during lactation (reviewed in [31]). This wide range of models all have a similar phenotype, suggesting a common pathway of these insults to act on later disease risk. Maternal calorie restriction in rats during gestation (up to 70% reduction) leads to a reduction in birth weight, and offspring go on to develop an obese, sedentary, and hyperphagic phenotype in adulthood [32, 33]. Similarly maternal protein restriction during just gestation in mice reduces birth weight; if the pups subsequently undergo catch-up growth during lactation they have increased susceptibility to diet-induced obesity [34]. Maternal high-fat feeding is detrimental to the offspring even if restricted to pregnancy or to lactation alone, leading to hypertension, hyperinsulinemia, and increased adiposity in adulthood [35]. These animals have altered body composition with increased fat percentage and higher amounts of abdominal fat [36]. The development of obesity involves a massive expansion of adipose depots. This is not as simple as hypertrophy of the adipocytes as they ﬁll with lipid, but also involves hyperplasia through adipogenesis forming new mature adipocytes.

24.4 Adipogenesis

Adipogenesis is the process by which numbers of adipocytes are increased and involves two separate processes. First, preadipocytes – the undifferentiated ﬁbroblast-like cells – must be recruited and proliferate. These cells then must differentiate into adipocytes, becoming cell cycle arrested permanently and taking on the characteristics of a mature adipocyte, such as a spherical, lipid-ﬁlled shape [37]. This process occurs throughout an organisms life time in response to an increased need for energy storage when in positive energy balance [38] as well as in response to normal cell turnover. Adipogenesis is a tightly controlled sequence of events with strict temporal regulation involving multiple signals that are both stimulatory and inhibitory. The end result is a switching on of adipocyte-speciﬁc or -associated genes in cells that are able to store increased amounts of lipid. It has been studied in detail using in vitro experimental systems such as 3T3-L1 cells. 24.4.1 Adipogenesis In Vitro

In cell culture experimental models, before preadipocytes can differentiate into adipocytes they must ﬁrst undergo growth arrest. This occurs on reaching conﬂuence by cell–cell contact inhibition and the cells are then growth arrested in the G1/S

24.4 Adipogenesis

phase [39]. At this point there is a downregulation in expression of inhibitory proteins such as preadipocyte markers such as preadipocyte factor-1 and an increase in expression of early differentiation markers (e.g., collagen type VI). The majority of information regarding the mechanism of adipogenesis has come from the study of experimental in vitro models. These generally require hormonal induction and the cocktails of proadipogenic signals used vary from system to system. For example, in the commonly used 3T3-L1 system, hormonal induction is brought about using methylisobutylxanthine, dexamethasone and supraphysiological concentrations of insulin [40]. This is used to ensure synchronous entry, maximizing the number of preadipocytes undergoing differentiation and so accelerating the differentiation process. Once cells have undergone hormonal induction, expression of c-fos, c-jun, and cmyc is increased [41]. These are believed to act to increase mitogenesis and their expression is only increased transiently. The transcription factors CCAAT/enhancerbinding protein (C/EBP)-b and -d also begin to be expressed during this time [42], and are thought to initiate mitosis [43]. The cells replicate their DNA and the cell number doubles [44]. During this period the DNA becomes accessible to transcription factors that regulate the expression of adipocyte differentiation. The ﬁnal steps of adipogenesis occur once the cells have become growth arrested again in the GD state [45] – this commits the cells to terminal differentiation. During this time expression of C/EBP-a [46] and peroxisome proliferator-activated receptor (PPAR)-c [47] increases, and this plays a role in initiating the arrest and hence terminal differentiation of the preadipocyte into mature functional adipocytes. Within 3 days the cells express markers indicative of late adipocyte differentiation [40]. Terminal differentiation occurs with an increase in the expression of proteins involved in glucose and lipid metabolism, and an increase in capacity for lipogenesis. The expression of cell surface receptors also takes on a more mature adipocyte pattern with upregulation of insulin receptor and glucose transporter 4 making the cells responsive to insulin [48]. There is also a change in the pattern of adrenoreceptor subtypes with a loss of b1 and an increase in both b2 and b3 subtypes, and an overall increase in number [49, 50]. The cells begin to have the physical characteristics of mature adipocytes with a spherical shape and the accumulation of lipid within the cell. The expression of adipocyte-secreted products such as leptin, adiponectin, and resistin begins, indicating the establishment of the cell as a mature adipocyte. 24.4.2 Control of Adipogenesis

The study of the control of adipogenesis has primarily used in vitro where cells in culture are exposed to various factors to see if the rate of proliferation or differentiation is altered. This has led to conﬂicting results, mainly due to the differences in cell lines, cell culture conditions, and hormonal induction protocols used. There are some common requirements, however, including insulin-like growth factor (IGF)-1, cAMP, and glucocorticoids [40].

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There is evidence from rodent models of obesity that paracrine regulation may be important in inducing adipocyte hyperplasia. This makes up part of the critical fat mass hypothesis [51]. When an animal is in positive energy balance, excess nutrition is stored as lipid primarily in adipocytes. However, if this state of positive energy balance is maintained the adipocytes soon will have reached their maximum lipidstoring capacity. At this point it is suggested that paracrine signals from these lipid-full adipocytes induce fresh adipogenesis [51]. This theory has been tested using in vitro systems, exposing preadipocytes to media exposed to adipose tissue and this induces differentiation [52]. When adipocytes are exposed to media from obese individuals this has a larger effect on inducing differentiation than material from nonobese individuals [53]. The attempts to identify the paracrine signals that when secreted by enlarging adipocytes induce adipogenesis has highlighted IGF-1 [54]. IGF-1 causes an increase in both the proliferation [55] and differentiation [56] stages of adipogenesis. It is thought to act through a pathway common to insulin, which induces the expression of transcription factors required for differentiation such as C/EBP-a and PPAR-c [57]. Endocrine regulation of adipogenesis involves insulin, which acts through the same pathway as IGF-1 via the IGF-1 receptor [58] prior to late differentiation and expression of the insulin receptor [56]. Glucocorticoids can also enhance adipogenesis in the presence of insulin [59] by causing transcriptional changes, most importantly in C/EBP-d, which leads to C/EBP-a expression and so differentiation. In vivo experiments have shown that growth hormone acts as a lipolytic agent in an anti-insulin like manner and leads to less lipid ﬁlling of adipocytes [60]. Growth hormone actions are less clear when studied in vitro, where initially it appears to have an insulin-like response [61]. Other factors can act to inhibit adipogenesis including epidermal growth factor [62] and tumor necrosis factor-a [63]. Dietary fat has also been shown to affect adipogenesis [64], and the source and composition of the fatty acids is important in determining the effect on both proliferation of preadipocytes and their differentiation into mature adipocytes. If rats are fed a diet rich in v-3 polyunsaturated fatty acids they have lower adipose tissue growth [65] and reduced lipid accumulation [66]. During adipogenesis, fatty acids can act as ligands for the PPAR transcription factors [67] and increase the expression of their target genes [68]. It is therefore important that both the amount and type of dietary fat consumed is optimal to prevent obesity. 24.4.3 Developmental Alterations to Adipogenesis

In humans, the process of differentiation leading to the formation of mature adipocytes begins in utero [69] and continues into early life. By birth, most of the differentiation processes have occurred, including lipid accumulation [70]. This process may be susceptible to changes in circulating nutrients or other metabolic signals in the fetus or neonate (e.g., fatty acids, IGF-1, and insulin). It has been shown that the nutritional environment experienced by sheep fetuses can affect the amount of adipose tissue that is deposited [71]. In rats, the offspring of dams fed a highly

24.5 Potential Mechanisms?

palatable diet in both gestation and lactation, which included free access to processed high-fat and/or high-sugar foods, including cake, potato, crisps cheese, and biscuits, have increased fat mass [72]. This increase in fat mass was accompanied by hypertrophy, in fact a doubling in adipocyte size, and adipocyte density, but no change in cell number [72]. In sheep, a reduction in maternal nutrition between early and mid gestation leads to an increase in the gene expression of IGF-1 and -2 receptors in conjunction with an increase in adipose tissue deposition [73]. If sheep are over-fed during late pregnancy gene expression of the proadipogenic transcription factor PPAR-c is increased, as is the gene expression of leptin and adiponectin [74]. As well as changes in nutrient availability that could alter rates of adipogenesis, many of the factors implicated in the control of adipogenesis are altered in models of developmental programming. IGF-1 levels are reduced in the rat fetus when the dam is fed an isocaloric low-protein diet and levels of IGF-binding proteins are increased [75]. In the long-term, IGF-binding protein-1 plasma concentration in old age in humans is correlated with birth weight [76]. Maternal insulin concentrations are also affected by maternal diet. In rats, feeding the dam an isocaloric low-protein diet increases maternal insulin in early gestation, but decreases it in late gestation [77]. While this insulin cannot cross the placenta it may signal to the fetus or affect the fetus nutrient availability. Proposed candidates for molecules involved in developmental programming of adipogenesis are shown in Figure 24.2.

24.5 Potential Mechanisms? 24.5.1 Glucocorticoids

Fetal exposure to increased glucocorticoid concentration leads to a similar phenotypic outcome to maternal nutritional models of programming and so may provide a mechanistic link. If rats are fed an isocaloric low-protein diet during pregnancy, there is a reduction in the activity of placental 11b-hydroxysteroid dehydrogenase (HSD)-2 that would increase the fetal exposure to maternal glucocorticoids as seen by increased activity of the glucocorticoid induced glutamine synthetase enzyme [78]. Rat dams fed a reduced calorie diet (50%) of control dams show reduced gene expression of placental 11b-HSD-2 at term [79]. Increased exposure to glucocorticoids during the time when adipogenesis is being initiated in the fetus may therefore affect the effect of the adipose tissue that is deposited. 24.5.2 Leptin

Leptin has been put forward as a candidate molecule for the programming of energy balance. This is primarily due to its effects on the developing hypothalamus, including the energy balance circuitry. In mice lacking a functional leptin gene,

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Figure 24.2 Schematic showing adipogenesis and candidate molecules for developmental effects on adipogenesis; these are both proadipogenic and antiadipogenic as indicated. SREBP1c, sterol regulatory element-binding protein-1c. Adapted from [90].

there is a reduction in the number of neurons projecting from the arcuate nucleus to the paraventricular nucleus in the hypothalamus and this reduction can be restored by giving leptin in early life [80]. Leptin concentrations peak in rodents during early life [81] before the brain is responsive to leptin effects on food intake and energy expenditure [82, 83]. As well as its neurotrophic effect, leptin can also affect adipogenesis. It has been shown in culture to act directly to inhibit the maturation of mature adipocytes [84]. If alterations in maternal and infant diet affect the timing and size of the leptin surge this could have long-lasting effects on the amount and function of the adipose tissue. It has been shown that the offspring of mice fed a globally calorie-reduced diet have a premature leptin surge [85]. If this is replicated exogenously in the offspring of control fed mice, they go on to develop obesity [85]. In rats, offspring of mothers fed 30% of ad libitum fed control mothers go on to develop a hyperphagic, obese, and sedentary phenotype in later life [32, 33]. This can be reversed by leptin treatment during lactation [86]. Leptins effect on adipose tissue

24.6 Future Perspectives

development may explain some of these consequences of alterations in leptin concentrations during this early period. 24.5.3 Epigenetic Alterations

The long-term, in some cases permanent, effects of brief experiences early in life suggest a role for epigenetic changes as an underlying mechanism. Permanent changes in the methylation status of genes have been observed in animal models. For example, maternal behavior has been shown to affect later glucocorticoid receptor expression in the brain of the offspring [87]. If an isocaloric low-protein diet is fed to rat dams then there is a reduction in methylation of the PPAR-a gene and an increase in mRNA expression [88]. This is also true of the glucocorticoid receptor [88]. These changes are prevented by folic acid supplementation [88]. These changes may be the consequence of a reduction in DNA methyltransferase-1 expression [89].

24.6 Future Perspectives

This area of study is still very much in its infancy, but understanding the mechanisms by which early-life experiences including nutrition program later adiposity would provide insight and potential healthcare options. Programmed changes in adipogenesis could lead to changes in fat mass and body composition, and changes in adipocyte function could alter feeding responses and satiety due to the important role of adipocytokines in regulating energy balance. 24.6.1 Optimizing Early Life Nutrition?

Data from animal studies, randomly controlled infant feeding trials, and epidemiological studies in humans will provide information on the optimal nutrition required during pregnancy and early life. If this can be translated into healthcare initiatives improving maternal and infant nutrition, then a large reduction in the prevalence of obesity, type 2 diabetes, and metabolic syndrome may result. 24.6.2 Interventions?

As more and more is uncovered regarding the mechanism linking early-life experiences and later disease risk, it may become possible to intervene either during pregnancy or postnatally to reverse the development of the detrimental phenotype. Currently there is much more mechanistic detail needed before this can be attempted, but potential interventions that alter adipogenesis provide key candidate molecules for investigation.

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H., Masuzaki, H., Nakao, K., Kawamura, M., Takemura, M., Kakui, K., Ogawa, Y., and Fujii, S. (2005) Role of premature leptin surge in obesity resulting from intrauterine undernutrition. Cell Metab., 1, 371–378. Vickers, M.H., Gluckman, P.D., Coveny, A.H., Hofman, P.L., Cutﬁeld, W.S., Gertler, A., Breier, B.H., and Harris, M. (2005) Neonatal leptin treatment reverses developmental programming. Endocrinology, 146, 4211–4216. Weaver, I.C., La Plante, P., Weaver, S., Parent, A., Sharma, S., Diorio, J., Chapman, K.E., Seckl, J.R., Szyf, M., and Meaney, M.J. (2001) Early environmental regulation of hippocampal glucocorticoid receptor gene expression: characterization of intracellular mediators and potential genomic target sites. Mol. Cell. Endocrinol., 185, 205–218. Lillycrop, K.A., Phillips, E.S., Jackson, A.A., Hanson, M.A., and Burdge, G.C. (2005) Dietary protein restriction of pregnant rats induces and folic acid supplementation prevents epigenetic modiﬁcation of hepatic gene expression in the offspring. J. Nutr., 135, 1382–1386. Lillycrop, K.A., Slater-Jefferies, J.L., Hanson, M.A., Godfrey, K.M., Jackson, A.A., and Burdge, G.C. (2007) Induction of altered epigenetic regulation of the hepatic glucocorticoid receptor in the offspring of rats fed a protein-restricted diet during pregnancy suggests that reduced DNA methyltransferase-1 expression is involved in impaired DNA methylation and changes in histone modiﬁcations. Br. J. Nutr., 97, 1064–1073. Feve, B. (2005) Adipogenesis: cellular and molecular aspects. Best Pract. Res., 19, 483–499.

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25 Evolutionary Aspects of Obesity and Adipose Tissue Function Jonathan C. K. Wells

25.1 Introduction

Substantial increases in the prevalence of adult obesity in the United States over recent decades strongly contradict any notion that human obesity is primarily a genetic condition. Rather, obesity is best considered a normal response to an abnormal environment rather than vice versa [1], although this then requires that we identify which particular components of the environment are abnormal from an evolutionary perspective. With obesity rapidly increasing in modernizing as well as industrialized populations, any simple concept of modern afﬂuence as the environmental abnormality is clearly inadequate. A more sophisticated evolutionary approach to fatness is therefore critical for understanding why an obesity epidemic should have emerged so rapidly in so many populations inhabiting diverse geographic regions and socioecological environments. Such a perspective is only recently emerging [2–5]. Obesity researchers have consistently considered the human propensity to store fat during positive energy balance as an adaptation to cycles of feast and famine in our past [6–8], although such cycles are rarely described in more detail. Reference is frequently made to our thrifty genotype, although again it remains unclear as to whether this refers to between-population variability in energy metabolism, or a trait of humans per se. The concepts of uncertainty in energy supply and genetic adaptation in response are clearly relevant to an evolutionary understanding of human adipose tissue biology and the prevalence of obesity in the contemporary world. Nevertheless, they represent an insufﬁcient framework with which to approach this issue. Adipocytes are specialized vertebrate cells that store triglyceride for export to other tissues as required, buffering their requirement for glucose from perturbations in energy supply [9]. Regional variation in insulin metabolism aids the allocation of energy between tissues. Across the vertebrate range, individual species vary widely in their size, anatomy, morphology, behavior, and equally in their adiposity. Indeed, Pond has stated that no other organ is so variable between species as adipose tissue and that lipid storage signiﬁcantly expands the range of niches in which different

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animal species can live [9]. If all stored energy represents metabolic thrift, then it is clear that vertebrate species vary extensively in their proﬁles of thriftiness, reﬂecting their diverse ecological environments and life history strategies. To understand human adipose tissue, and its variability, we must therefore understand the particular metabolic stresses that characterized hominin and human evolution. In this chapter, I argue that the human proﬁle of adiposity is as fundamental to our species as are our other trademark features such as bipedalism, large brains, and long lifespans. Indeed, I argue that human adipose tissue biology is intricately connected with these other traits, that they emerged as a suite of complementary adaptations selected to facilitate a colonizing reproductive strategy [10], and that a slow-growing, large-brained, colonizing hominin would be biologically impossible in the absence of complementary adaptations in adipose tissue biology.

25.2 Thrifty Genotype and Phenotype Hypotheses

Neels thrifty genotype hypothesis [11] is often considered the ﬁrst evolutionary perspective on human metabolism, and our contemporary propensity to obesity and diabetes, and represents an early reference to cycles of energy uncertainty. Thus it must be remembered that during the ﬁrst 99% or more of mans life on earth, while he existed as a hunter and gatherer, it was often feast or famine. Periods of gorging alternated with periods of greatly reduced food intake ([11], p. 355). In such an environment, Neel argued, natural selection would favor both the capacity to store fat when plasma glucose levels were high and the capacity to draw on that fat when plasma glucose levels were low. Central to his hypothesis was the notion of a quick insulin trigger; in other words, enhanced insulin production, favoring storage of circulating glucose in adipose tissue. Subsequent work has failed to support this concept of enhanced insulin production, but has replaced it with the notion of insulin resistance in muscle tissue [12], which would likewise function to divert circulating glucose to adipose tissue. The thrifty genotype hypothesis has been highly inﬂuential in both anthropological and biomedical research on human energy metabolism and the metabolic syndrome [13]. However, it has been used in different ways and it is important to clarify how it contributes to understanding the evolutionary biology of adipose tissue. Neels primary argument was that a thrifty genotype would predispose to diabetes and obesity, once uncertainty in energy supply was attenuated following the emergence of agriculture. Recent analyses of the ethnographic and archaeological records have failed to support the hypothesis that energy uncertainty was high prior to the emergence of agriculture and then decreased. No statistical difference in the frequency or extent of food shortages between preindustrial or recent foragers versus agriculturalists was apparent in a sample of 94 populations [14], while the archaeological record reveals signiﬁcant deteriorations in human health following the emergence of agriculture in diverse regions of the world [15–17]. Thus, there is little

25.2 Thrifty Genotype and Phenotype Hypotheses

evidence to support the hypothesis that preagricultural populations regularly gorged and fasted on a scale greater than that characteristic of farming populations. Contemporary foraging populations exploit a substantially wider range of resources compared to agricultural populations and can respond to seasonal uncertainty in resource availability by mobility. It could be argued therefore that agriculture, rather than foraging, might have exposed populations to regular famines and hence selected for thrifty genotypes. Certainly there is plentiful evidence of regular famines throughout the history of human agriculture [18], whilst humans have also developed a diverse range of subsistence practices across the global range of ecological conditions and have therefore exposed themselves to localized selective pressures. Such local niche construction [19] appears to have favoured a variety of polymorphisms of genes that regulate energy metabolism [20]. Where Neels hypothesis may be more relevant, therefore, is in accounting for contemporary population variability in vulnerability to diabetes and obesity. Contemporary humans, whether foragers, horticulturalists, pastoralists, or farmers, and whether nomadic or sedentary, occupy a wide range of socioecological environments. The longer a population has inhabited any given niche, the greater the opportunity for natural selection to have favored genetic adaptation. It is certainly plausible that populations that undertook difﬁcult migrations, as has been hypothesized during the colonization of Polynesia and the North American continent, for example, may have undergone particularly strong selection for thrifty genes [21]. Nor is such selection limited to the distant past, with the slave trade and mass immigration from Ireland to the United States, for example, both proposed to have exposed migrating populations to signiﬁcant selective pressures on metabolic thrift [18]. The evidence from polymorphism studies is that there is a wide range of thrifty genes, all of which contribute to variability in susceptibility to obesity. Contemporary genetic variability may therefore contribute to subtle variation in the susceptibility to obesity, diabetes, and the metabolic syndrome. Populations now exposed to the Western diet, high levels of overweight, and sedentarism can show markedly contrasting prevalences of diabetes. Native American, Polynesian, and Australian aboriginal populations appear to have particular genetic proﬁles that maximize their vulnerability to diabetes, whereas Europeans appear to have a degree of protection [13], but these examples probably represent different points on a continuum of susceptibility. Nevertheless, recent experience in the United States suggests caution when considering the signiﬁcance of thrifty genes. The extraordinary speed with which obesity is increasing in this population strongly suggests that, given sufﬁcient exposure to the obesogenic environment, most individuals can become obese. This is particularly the case given that environmental exposure is now known to commence in utero. Increasingly, research suggests that the genetics of thinness [22] may be more relevant than the genetics of obesity. A small proportion of people appear relatively protected against excess weight gain, whereas the majority are more susceptible to obesogenic factors. The fact that obesity appears to be highly heritable need not imply a particularly strong role for speciﬁc genetic factors.

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The contribution of ontogenetic experience to obesity susceptibility is addressed using an approach broadly known as the thrifty phenotype hypothesis [23]. This approach focuses on the concept that exposure to environmental factors during fetal life and infancy impacts on developmental trajectory during windows of early-life plasticity [23]. Evidence increasingly supports the hypothesis that maternal phenotype inﬂuences offspring ontogeny through a variety of pathways, including hormonal programming, behavior, and epigenetics [24]. Detailed research on the Pima Indians, a population with high susceptibility to diabetes and obesity, has highlighted the signiﬁcance of maternal diabetic status during pregnancy on the risk of obesity and diabetes in the offspring [24]. Thus, even in populations considered to have a genetic susceptibility to obesity, nongenetic factors appear critical in the etiology of the condition. While the thrifty genotype is therefore a useful concept when considering population variability in susceptibility to diabetes and obesity, it is of limited value for understanding the broader global trend towards an epidemic of obesity. To address this issue it is necessary to turn away from contemporary variability in genotype and environmental conditions, and to focus on the longer-term evolutionary history of the hominin lineage. Anatomically modern humans are currently estimated to have evolved between 150–200 000 years ago [25], however recent genetic studies indicate that while the majority of genetic variation between nonAfrican populations emerged since this speciation event, African populations retain genetic variability that emerged over a much longer time period, through admixture between ancient African hominin populations. Thus, both contemporary human variability and those genetic factors common to all living humans reﬂect selective pressures acting throughout hominin evolution, with many of these selective pressures deriving from environments very different to those occupied by modern. Addressing this issue is notoriously difﬁcult, due to the absence of soft tissue in the fossil record. Nevertheless, although fat represents a physical trait, it can also be considered as a strategy. Our understanding of the characteristics and functions of adipose tissue in contemporary humans allows us to attempt a reconstruction of the evolutionary history of hominin body fat, using an indirect approach proposed by Tinbergen [26] for studying behavior.

25.3 Ethological Approach

Tinbergens approach focused on four separate perspectives, considering the proximate causes, ontogeny, ﬁtness value, and evolutionary history of a given trait [26]. This approach allows propositions of the function of a trait to be well grounded in what is known of physiology and ontogeny. The proximate causes of fat deposition have been considered in detail elsewhere, and include genetic factors, diet and physical activity levels, feeding behavior, endocrine and inﬂammatory factors, and

25.3 Ethological Approach

psychological and broader environmental factors [5]. This chapter addresses the remaining three issues. 25.3.1 Ontogeny

The ontogeny of fat deposition in nonobese populations provides important clues as to the relative value of energy stores at different points in the life course and between the sexes. Human neonates stand out from other species in having high body fat content at birth and increasing it further during infancy [3]. During childhood growth, relative body fat content declines and growth is disproportionate in lean mass. However, during adolescence the sexes increasingly diverge. Girls gain limited lean mass, but substantial body fat, whereas the weight gain of boys is largely attributable to increases in lean mass [5]. By adulthood, all human populations display signiﬁcant sexual dimorphism in body composition, with males being taller, and having greater muscle mass and less fat (Figure 25.1). Females not only have more body fat, but a markedly different distribution with fat being located primarily on the thighs, buttocks, and breasts, in contrast to the abdominal deposition of fat in males [27]. The life-course pattern of fat deposition in both sexes is illustrated in Figure 25.2, and indicates greater value of adipose tissue stores during infancy and in reproducing females.

24

Lean mass index Fat mass index Line of identity

20 16 Male value (kg/m2) 12 8 4 0

0

4

8

12

16

20

24

Female value (kg/m2) Figure 25.1 Sex differences in lean mass index (lean mass divided by height squared) and fat mass index (fat mass divided by height squared) in 35 non-Western populations plotted on the line of identity. In all populations, relative fatness is greater in women than in men,

especially at the higher end of the spectrum of fatness. Equally, relative lean mass is greater in men than in women, although the magnitude of the sex difference is very low in some populations.

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Female Adulthood Relative fatness Male

birth

Relative lean mass Figure 25.2 Ontogenetic development of body composition in males and females. The lines describe changes in relative fat mass relative to height (y-axis) and lean mass relative to height (x-axis) between birth and adulthood. Body fat stores are high at birth and increased

substantially during infancy. They then decline during childhood, but increase markedly during adolescence in females, whereas males gained primarily lean mass during the same period. Reproduced with the authors permission from [65].

25.3.2 Fitness Value

The ﬁtness value of body fat stores refers to their effects as opposed to their causes [26]. Selection has acted not only on energy stores per se, but also the capacity to acquire them relatively rapidly. These energy stores are then used for a variety of functions, many of which are particularly important in the biology of our species. As recognized by many authors, fat represents a store of energy for buffering ﬂuctuations in energy supply. Fat stores are not appropriate for rapid bursts of physical activity, as lipid mobilization is slow in relation to the immediate rate of energy utilization [9]. Each kilogram of fat contains 37 MJ, equivalent to around 6 days energy supply for a typical adult individual. Adult males and females of average body fat content could theoretically survive starvation for around 60 or 90 days, respectively, as demonstrated in the literature [4]. However, total famine is likely to be a relatively new stress in hominin evolution and fat stores can clearly buffer moderate energy constraint for substantially longer than famine. Data from contemporary populations inhabiting highly seasonal environments illustrate signiﬁcant annual ﬂuctuations in body weight, of which the majority has been demonstrated to comprise gains and losses in fat mass [28]. The role of fat in addressing such perturbations is in no sense unique to humans. A recent elegant study of orangutans, which also inhabit an environment characterized by seasonal variation in the availability of fruit, demonstrated marked variations in energy balance during the year. When energy intake was less than energy utilization, ketones (produced by lipolysis) were detected in urine samples, particularly in pregnant or lactating females [29]. Thus, the use of fat stores to accommodate seasonal energy uncertainty is a general mammalian and primate trait, merely conserved in humans. However,

25.3 Ethological Approach

humans certainly show the capacity to gain such fat stores rapidly. Data from a West African population show that a traditional fattening period, characterized by up to a 3fold increase in energy intake, resulted in a 25% increase in body weight over 2 months, of which approximately 70% was fat [30]. Nutritional status exerts powerful effects on female reproductive function, such as inﬂuencing the regulation of menarche and conception. In the 1970s, the critical threshold hypothesis of Frisch predicted that reproductive viability could only occur above a certain body fat content [31]. This idea has remained highly controversial. On the one hand, Ellison [32] has argued that energy ﬂux plays a more important role than energy stores per se and that there is a graded continuum of female fertility. On the other hand, humans are capital breeders [33] and readily fund the expensive period of lactation from energy stores, although energy intake can also increase. It is clear that malnourished women have a decreased risk of conception and an increased risk of miscarriage early in pregnancy or stillbirth. In mice, the hormone leptin plays an important role in the regulation of reproductive function and administration of the hormone to leptin-deﬁcient mice restores function [34]. A similar mechanism is predicted to be found in humans [34]. However, while insufﬁcient fat is one constraint on reproductive function, excess fat stores may likewise impose constraints. Obesity is associated with a number of disorders that inﬂuence both the probability of conception and the course of pregnancy [35]. The beneﬁts of body fat to female reproductive ﬁtness therefore appear to apply primarily to those at the lower end of the scale of nutritional status. Energy stores appear particularly important in mediating the impact of the expensive human brain on offspring growth rate. Brain tissue is metabolically costly and due to its negligible plasticity imposes obligatory demands on the energy budget. This is particularly the case during early life, when the brain is a relatively large proportion of body weight and accounts for over 80% of basal metabolism [3]. High levels of body fat during infancy have been attributed to their value in buffering the inﬂexible and expensive infant brain from ﬂuctuations in energy intake [3]. Consistent with this hypothesis, malnourished infants have been found to switch rapidly from carbohydrate to fat metabolism during fasting, with over 90% of energy expenditure derived from fat metabolism during malnutrition [36]. These energy stores must, however, be obtained from the mother during the periods of pregnancy and lactation. High levels of body fat in reproducing females have therefore been attributed to their capacity to ensure offspring energy demands can be met throughout the period of parental care. Consistent with this hypothesis, breast-milk transfer appears relatively robust to ﬂuctuations in maternal energy supply [37]. In addition to buffering the brain, fat stores also fund the immune system – a function again of particular importance during early life. The immune response is now known to be metabolically costly and leptin has been shown to mediate the allocation of energy to this function [5]. Finally, body fatness is subject to sexual selection [4]. Body shape in females may act as a signal of fertility, expressing fecundity and the presence or absence of energy stores. However, signaling systems need not be honest and sexual selection may operate through runaway bandwagon mechanisms, whereby attractive traits

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become favored despite having no clear functional correlate. Both mechanisms appear plausible in relation to human fatness [5], the signiﬁcance here being that a particular phenotype may promote reproductive success without fat stores themselves contributing to any of the physical functions described above. 25.3.3 Evolutionary History

Body fat stores in any animal are the product of detaching energy intake from an exact association with energy utilization in basal metabolism, physical activity, and growth. The ecological pressures that favor fat stores differ widely between species [9]. It is possible to identify several distinct ecological pressures that have favored energy stores in humans and to estimate broadly when during hominin evolution they might have acted. Contemporary great apes (gorillas, chimpanzees, and bonobos) are thought to have shared a common ancestor with hominins around 6–8 million years ago, with the hominin line splitting from chimpanzees and bonobos around 5–7 million years ago. Ancestral gorillas, chimpanzees, bonobos, and hominins appear to have entered different habitats, characterized by varying duration of the dry season [38]. Whereas gorillas remained in a more forested niche, with only a short dry season, chimpanzees occupied an intermediate environment, and hominins pushed eastwards into a new emerging savanna niche with a longer and more variable dry season [38]. This environment, imposing greater seasonality of energy supply, may have begun selection for energy stores to buffer seasonal ﬂuctuations. It is probable that seasonality acted more strongly in some periods of hominin evolution than in others. Nevertheless, its pressure is likely to have been felt from the australopithecine radiation onwards. Although the australopithecines differed from nonhominin apes in developing bipedalism, relative brain size remained very similar. A marked transition towards greater brain size occurred with the emergence of the genus Homo, with Homo ergaster initiating a trend to encephalization continued by H. sapiens [39]. Changes in relative brain size generate profound impact on an organisms energy budget. For example, 20% of adult human basal metabolism is directed to the brain, in contrast to 9% in the chimpanzee [3]. However, the high energy costs of the human brain impact most strongly at the start of life and hence also on the maternal energy budgets due to the mothers role in supplying the energy for offspring growth. Figure 25.3 demonstrates the changes in body proportions that occur during human growth [40]. The signiﬁcance of the increase in brain size is that, despite considerable historical attention to the supposed importance of hunting and tool use by males in hominin evolution, females are likely to have been subject to the strongest selective pressures for energy stores [10]. Sexual dimorphism in body size and composition is characterized both by reduced lean mass in females and increased fat mass, relative to males [27] (Figure 25.1). This proﬁle reduces basal energy needs whilst increasing stores of energy for buffering energy ﬂuctuations and funding the requirements of reproductive events.

25.3 Ethological Approach

Figure 25.3 Changes in body proportions during growth from birth to adulthood. Of particular significance for human energetics is the decrease in the relative size of the brain. Reproduced from [40].

Along with such maternal characteristics, the ontogenetic developmental schedule of offspring also appears to have responded to selective pressures. Hominin evolution appears to have been characterized by increases in body size and increases in life expectancy, which in turn act to prolong the growth period. Childhood and juvenile growth in humans are strongly canalized, and relatively insensitive to nutritional inﬂuence. During fetal life and infancy, however, increased nutritional supply promotes growth with long-lasting effects. Given that early-life nutrition derives from the mother, who continues to provision each offspring even after weaning, this sensitivity of early growth nutrition allows alignment of offspring growth trajectory with maternal phenotype [24, 41]. The human pattern of growth appears to have steadily extended during hominin evolution – a broader trend already evident in primates. Thus, the lengthy growth period in H. sapiens likely reﬂects the extreme energy costs imposed by the large brain, reducing the load on the maternal energy budget [24, 42]. For the majority of hominin evolution, environmental stochasticity deriving from seasonality and longer-term climate trends is likely to have been the key selective pressure on body fat, with increases in brain size further increasing vulnerability of hominins to those pressures. Within the last 2 million years, however, the genus Homo has itself generated new selective pressures, which must have induced further selection for energy stores.

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First, H. ergaster spread rapidly between Africa and Asia, and colonized an enormous geographical range, with little evidence of reliance on technology [43]. Such dispersal may not have involved the movement of long distances by individuals, but large-scale colonization must inevitably have involved continual exposure of the Homo gene pool to novel ecological niches. The foraging patterns of H. ergaster must likewise have impacted on the environment, thus shaping the selective pressures acting back on the species. This evolutionary process of niche construction [19] is likely to have maintained selection for energy stores for buffering reproduction against uncertainties and ﬂuctuations in energy supply. Those characteristics shared by all humans today must have been shaped during this evolutionary period, prior to the evolution of H. sapiens. Second, following the emergence of H. sapiens and a second rapid colonization of the majority of the Earths land masses, localized selective pressures have given rise to population-speciﬁc genetic variability. This variability now manifests as the range of thrifty genes that has been linked with differential susceptibility to obesity and diabetes, discussed above [20]. Such selective pressures may have included broad climate, local ecology and dietary niche, seasonality, and disease load. Within the last 12 000 years or so, agriculture rapidly emerged as a new and powerful selective pressure, again acting locally in combination with climatic and ecological factors.

25.4 Significance of Agriculture

Agriculture emerged independently in several regions of the world [44]. Recent models suggest that the shift in subsistence strategy from foraging to farming, that occurred independently in several regions of the world within a narrow time frame, was favored by a sudden global reduction in ecological stochasticity [45]. Far from improving energy supply, agriculture is now understood to have worsened the human diet and to have induced downward secular trends in body size [16]. Regular famines, in combination with the poorer dietary range and the deterioration of stored foods, imposed severe energetic stress on farming populations. For example, the particular volatility of monsoon conditions may have contributed to the sensitive metabolic proﬁle of some Asian populations [46], although imperial economic policies over two centuries may have exacerbated these pressures (see Section 25.6). Furthermore, sedentary populations were exposed to a greater burden of infectious disease [47], in particular from diseases that jumped the species barrier from newly domesticated animal species [44]. Recent palaeodemographic research is increasing our understanding of the associations between human life history and the burden of disease. It is often assumed that few humans lived beyond 45 years until recent centuries, implying, for example, that adult obesity would not have been subject to major selective pressures. Such an approach ignores the fact that human life expectancy is most strongly inﬂuenced by mortality in early life. Recent modeling, based on long-term trends in child mortality between the Mesolithic and early mediaeval periods, suggests an

25.5 Significance of Colonizing

increase in the frequency of epidemics [48]. Shorter intervals between epidemics increase child rather than adult mortality and over generations convert infectious diseases into diseases of childhood. Given the energy costs of mounting an immune response, this evolutionary process may therefore have favored increased levels of body fat in younger age groups. Thus, the development of agriculture and the associated increased burden of disease may have favored both thrifty genes in different populations and also changes in the ontogenetic proﬁle of body fat. The development of agriculture involved far more than just changes in subsistence patterns and dietary intake. Trends in subsistence mode were accompanied by profound changes in social structure and generated major inequality in social status, which continues to be a primary determinant of health status in contemporary populations. Whilst agriculture may have been favored by increasing population size, it also would have acted to promote population growth. Many studies have demonstrated how changes in subsistence practices induce increases in fertility rate, although the relative contribution of various mechanisms remains unknown [49]. Despite poorer health, reﬂected in lower life expectancy, populations would therefore have grown rapidly in size as noted in the archaeological record [16]. As population size increased, the simpler kin relationships that characterize egalitarian societies would have given way to a new set of economic and political relationships, whereby access to resources increasingly depended on rights and obligations rather than subsistence productivity itself [16]. From around 5000 years ago, large-scale civilizations emerged apparently independently in several parts of the world, in which differential control of the means of production became extreme [16]. Social inequality then translates directly into inequality in access to resources during periods of hardship, inducing differential exposure to famine.

25.5 Significance of Colonizing

Contemporary humans occupy a huge geographical range in comparison with the modest territories inhabited by nonhuman apes. The most recent geographical expansion was aided by technology, but the earlier dispersal of H. ergaster was substantially achieved on the basis of physiological plasticity and behavioral variability [43]. Humans have been termed colonizing apes by a number of authors [50–52] who have noted both their wide habitat range and also their reproductive proﬁle. Relative to other apes, humans produce offspring at a faster rate and suffer reduced adult mortality. When juvenile mortality is reduced, this results in rapid population growth, as observed in many populations recovering from disease epidemics [52, 53]. Much of human biology ﬁts the proﬁle of a colonizing organism [10] and body fat may play a critical role in allowing hominin females to pursue a colonizing reproductive strategy even while encountering the challenges of novel habitats and environments.

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Consistent with the notion of humans as capital breeders [33], research increasingly highlights the role of both adipose tissue, and its hormonal product leptin, in the regulation of reproduction. Body fat stores and, more importantly, energy ﬂux regulate the probability of conception [32]. Baseline maternal nutritional status predicts birthweight of the offspring more strongly than does pregnancy weight gain [24]. Birth is predicted to occur when fetal energy requirements can no longer be met by placental nutrition, since breast-milk allows greater transfer of fat [32]. In postnatal life, maternal fat stores meet the high energy demands of lactation – of particular importance in women subject to constraints on energy intake. A recent longitudinal cohort study showed that age of menarche in the mother predicted both her own size and fatness, and also the growth rate of her offspring [54]. However, this maternal effect on offspring growth was limited to the infant period, which is both the period of growth under nutritional control and the period funded directly by maternal physiology. The implication is that the offspring senses maternal fat stores in utero and adapts its infant growth rate accordingly. At the end of infancy, growth rate becomes canalized and individuals tend to track along a given centile. The role of maternal fatness in guiding offspring growth trajectory and rate of maturation highlights the way in which human development is a function of maternal reproductive strategy. As a colonizing organism, the genus Homo could have encountered two types of situations on a regular basis. (i) Populations migrating to new environments could have found plentiful resources free of conspeciﬁc competition. Such circumstances would have favored a high rate of reproduction, until population density began to impact on the availability of resources. High levels of body fat could, along with plentiful energy supply, have contributed to the capacity of females to achieve such reproductive expansion. (ii) Such population booms would inevitably have predisposed to population busts, due to overexploitation of resources. Evidence for such local overexploitation is increasingly emerging from the archaeological record [55]. Body fat stores could then have contributed to the ability to withstand the onset of poorer conditions and the subsequent migrations of subpopulations in search of improved quality habitats. Selective pressures during hominin evolution may therefore have favored the evolution of traits that predisposed to rapid population growth, which in turn increases the risk of local population crashes. Body fat could contribute to such boom and bust cycles – fuelling the booms and accommodating the busts. The notion that humans have imposed on themselves the selective pressures for high levels of body fat resolves the problem of accounting for unusual features of human adipose tissue biology relative to other mammals despite occupying common geographical areas. The cycles of feast and famine so often alluded to in the literature may not be primarily determined by extraneous environmental factors, but rather have increasingly derived from aggressive human niche construction. Importantly, males as well as females would have been exposed to such boom and bust situations, which could help explain why both sexes are now susceptible to obesity.

25.6 Significance of Social Inequality

25.6 Significance of Social Inequality

Social inequality is by no means unique to humans and is observed in many animal species in the form of dominance ranking, maintained through a combination of physical force and attention-seeking behavior. Observations of chimpanzees, for example, highlight the role of alliances between individuals in addressing social challenges and violent raids against neighboring groups to acquire their territory [56]. Nevertheless, the emergence of civilization has potentiated a change in the scale of social hierarchy, towards large-scale coercive power. In socially stratiﬁed societies, social rank is a key factor affecting survival during difﬁcult periods [55]. Given that agriculture directly invoked regular famine, social inequality has been closely associated with differential selective pressures for several millennia. This has two implications for contemporary populations. (i) Social inequality may plausibly map onto genetic variability, due to cumulative ancestral exposure to different selective pressures. (ii) More importantly, over shorter timescales social inequality may impact by nongenetic mechanisms on forthcoming generations. As discussed above, phenotype of the offspring is highly sensitive to maternal phenotype during early life [24]. Although this sensitivity is broadly adaptive, low maternal social status inadvertently contributes to this mechanism and may induce a developmental trajectory in the offspring that reproduces adverse traits in the next generation. This scenario, arguably of increasing importance over the last few millennia following the evolution of profound social inequality, represents a metabolic ghetto [24], making it very difﬁcult to release individuals within their own lifespan from the consequences of the poor social circumstances experienced by their parents and recent ancestors. The metabolic ghetto may be particularly relevant to the experience of populations subjected to the forces of imperialism. Ecologists by convention assume organisms to select habitats in order to maximize ﬁtness – a concept known as the ideal free distribution [57]. At such a distribution, individuals in different habitats would have equal average ﬁtness; however, within any habitat, individual ﬁtness would decrease as a function of the abundance of conspeciﬁcs, favoring a dispersal to exploit new resources. If key resources are limited, those displaced from the optimum environment have poorer ﬁtness [58]. Humans are unique in the scale on which some individuals actively control the distribution of others. High status individuals not only monopolize the better resources, but also control the circumstances and territory of those of low status. The politicoeconomic process of imperialism enacted this manipulation on an extreme scale – the initial effect being systematically to malnourish entire indigenous populations. In recent centuries, European colonial powers destroyed indigenous industry and coercively introduced export agriculture at the expense of local food production. After 187 years of British rule, Nehru described Bengal as a miserable mass of poverty stricken, starving, and dying people ([59], p. 247). Imperialist policies, aided by the harsh monsoon climate, could be argued to have pushed the vast majority of the Indian population into the metabolic ghetto, reﬂected in their small contemporary average size and metabolic thrift. Such a view is supported by historical

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data on height, showing no long-term improvement in contrast to Europe [60], but merits further examination. In the decades since independence, global economics are increasingly imposing a second nutritional burden on these same populations, exposing them rapidly to the Western obesogenic niche (see Section 25.7) and the risk of diabetes. Thus, just as the emergence of agriculture generated large-scale social inequality, so the political and economic forces that emerged during the industrial revolution are continuing the manipulation of energy metabolism across entire populations.

25.7 New Obesogenic Environment

The present obesity epidemic can be attributed to the exposure of large-brained colonizing apes to a series of niches that ﬁrst enhanced selective pressures for energy stores and then within the last few generations negated them (the Western industrialized niche). Much of the contemporary effort to address the obesity epidemic comprises attempts to apportion greater responsibility to individuals for their food consumption and activity level While such efforts represent logical approaches to the manipulation of individuals bodyweight, the emergence of an obesogenic environment comprises a great deal more than simple secular trends in diet and activity level. The Western industrialized niche is closely associated with the capitalist economic model and the transformations in behavior that predispose to obesity have a fundamental connection with this model. It is no accident that the precursor for industrial capitalism that now dominates Western countries was the development of overseas sugar plantations exploiting slave labor [61]. This proto-capitalism not only initiated the displacement of individuals from their local subsistence niche, but also made possible the extraordinary domination of dietary intake by sugar. The capitalist model requires constant expansion of consumption in order to drive proﬁts. Sugar proved the ideal product, such that per capita intake of sugar has increased several thousand-fold over recent centuries in industrialized populations [61], displacing other dietary components. The Caribbean plantations that produced sugar acted as the model for both the evolution of the consumer and the manipulation of that consumer for the beneﬁt of capitalist enterprise. Central to the contemporary obesogenic environment are numerous constraints and pressures acting on individuals, such that secular trends in diet and activity level are not driven directly through individual volition, but rather indirectly through economic strategies intended to maximize corporate proﬁts. The fundamental role of vested interests substantially reduces the power of individuals to remove themselves from the obesogenic environment. Modern companies increasingly derive their proﬁts from manipulating behavior, rather than providing material goods. Such behavioral manipulations include increasing average waking hours and the construction of consumers of experience, which in practice is often remarkably passive. Other secular trends deriving from the same process include later age of

References

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j491

Index a acetyl-CoA carboxylase (ACC) 357 acquired lipodystrophies 393 – acquired generalized lipodystrophy 393 – – clinical features 393 – – molecular genetics 393 – acquired partial lipodystrophy (APL) 386, 393 – – clinical features 393 – – molecular genetics 394 – HIV-related lipodystrophy 394 – possible genetic component 393 acquired partial lipodystrophy (APL) 386, 393 acylation stimulation protein (ASP) 28, 290 acyl-CoA 109, 117–120 – activation 109 – homeostasis 120 – metabolism 116, 120 – – acyl-CoA-binding protein, role 116–118 – – FABPs/ACBP mediate 118 – – metabolic diseases 120 – partitioning, ACSLs/FATPs control 117 – pools, regulation/function 118 – trafﬁcking 119 acyl-CoA-binding protein (ACBP) 116, 117 – expression 117 – function 117 – role 117 acyl-CoA synthetases (ACSLs) 113 – ACSL-1, heart-speciﬁc overexpression 115 – activity 114 – isoforms 114 acylglycerolphosphate acyltransferase (AGPAT) enzymes 411 ADD-1 protein 7 adhesion markers, modiﬁcation 267 adhesion molecules 275

– intercellular adhesion molecule (ICAM) 275 – vascular cell adhesion molecule (VCAM) 275 adipocyte(s) 3, 55, 85, 130, 131, 157, 186, 270, 285, 348, 412, 449, 473 – Acrp30, see adiponectin – a2-adrenergic receptors, physiological stimulation 155 – b3-adrenoceptors 55 – biology 270–274 – – adipose tissue, paracrine cross-talk 272–274 – – chemoattraction 271 – deﬁnition 285 – functions 407 – insulin resistance 348 – leptin synthesis, transcriptional regulation 186 – lipid droplets 85 – lipogenesis, schematic presentation 130 – lipolysis, key players 83–88 – – droplet scaffold proteins 84–86 – – lipases 86–88 – – lipid droplets 84–86 – – schematic presentation 131 – pertussis toxin treatment 157 – precursors 59 – transcription factor target genes, identiﬁcation 9–13 adipocyte lipid-binding protein (ALBP) 151, 289 adipocytokines 129, 265 adipogenesis 4, 6, 30, 31, 459, 464, 466 – control 463 – developmental alterations 464 – differences 30–32

j Index

492

– early-life programming 459–467 – endocrine regulation 464 – human preadipocyte capacity 30 – in vitro studies 447, 462 – markers 31 – potential mechanisms 465 – – epigenetic alterations 467 – – glucocorticoids 465 – – leptin 465–467 – PPAR-g, role 447 – regional variation 31 – schematic presentation 466 – transcriptional control 4–9 – transcription factors, induction 6 adipogenic pathogens 307–312, 324 adipokines 248, 265, 269, 270, 274, 353, 378–380 – adiponectin, and cancer risk 353, 379, 380 – animal models 380 – cancer risk 380 – expression mechanisms 246 – IL-6/8 270 – leptin, and cancer 353, 378 – overview 378 – production 268 – role 248 adiponectin 202, 205–208, 210, 212, 251, 271, 353, 379 – circulating levels 251 – clearance 214 – expression 208 – – oxidative stress 212 – – PPARg, activators 213 – – regulation 212–214 – gene, ADIPOQ 202 – gene expression 10 – multimers 203, 204, 206 – – bioactivity 204 – – high-molecular-weight (HMW) 203 – – low-molecular-weight (LMW) 203 – – multimerization 210 – – signiﬁcance 204 – oxidative stress 253 – receptors 201, 215 – – AdipoR1/R2 201, 215 – role 205–207 – – brain 207 – – in liver 205 – – in skeletal muscle 206 – – in vasculature 206 – secretion, regulation 208–214 – signaling 216 – structure 202 – targets 208

adipose angiotensinogen expression 248 adipose depots 286, 288 – adipocyte function, heterogeneity 288–297 – – glucose uptake 290–297 – – insulin action 290–297 – – lipolysis 288 – – triglyceride deposition 289 – deﬁnitions 286 adipose-derived glycerol 139 adipose-derived stem cells (ASCs) 69 adipose differentiation-related protein (ADRP) 317 – expression 85 adipose glycerol channel 132–141, 143 – AQP7 132–134 – – in adipocytes, function/regulation 134 – – genetic mutation 137–139 – coordination 143 adipose organ 50, 55 – gross anatomy 50 – phenotype 55–60 adipose-speciﬁc genes 133 adipose tissue 49, 59, 231, 246, 249, 253, 266, 269, 286, 333, 424 – ACSL-1 expression 121 – acyl-CoA synthetase activity 115 – adipose tissue macrophages, importance 293–295 – anatomic location 246 – antilipolytic responses 157 – aromatase enzyme activity 376 – brown adipose tissue (BAT) 49, 149 – cell types 269 – changes in cachexia 333–335 – characterization 71–73 – – clonality 72 – – cytokine proﬁle 72 – – immunogenicity 71 – – immunophenotype 71 – – proteomic analysis 72 – – transcriptomic analysis 72 – composition 286 – contribution, systemic inﬂammation 266–268 – depots 268 – derived hormones, inﬂuence 412 – development 403 – dysfunction 246, 293 – embryology 69 – expansion pattern, defects 349 – fatty acid 114, 115 – – activation/channeling 115 – – partitioning 114 – function 403

Index – generated hormones, summary 351 – human, gene expression proﬁle 129 – immunocytes, importance 293–295 – in situ localization 69 – in situ microdialysis 162 – isolation methods 70 – lipid content 109 – lipodystrophic remodeling 448 – lipoprotein lipase activity, inhibition 163 – loss mechanism, cachexia 337 – mass, governing factors 336 – metabolic changes 449 – mRNA expression 253 – pathogens, interaction 323 – physiological function 115 – secreted factors 231 – – adipose tissue-speciﬁc secretory factor (ADSF)/resistin 231 – – preadipocyte factor (Pref)-1 231 – stem cells 69 – variable degree 391 – white adipose tissue (WAT) 49, 149 – – buffering function 149 – – physiological role 149 adipose tissue triglyceride lipase (ATGL) 87, 151, 152, 337 Adiposity, early-life programming 459–467 a2-adrenergic receptors 154, 155 – physiological role 155 b-adrenergic agonists 214 b-adrenergic antagonist 167 b2-adrenergic pathway 153 – b2-adrenergic receptor density, role 153 b-adrenergic receptor 153 – dependent lipolysis 153 – gene, polymorphism 154 – physiological contribution 153 ADSF/resistin 236, 237 – expression 237–239 – function 237–239 – identiﬁcation 236 – structure 236 adventitial-derived relaxing factor (ADRF) 256 aging process 424, 427 – free radical theory 426 – physiological changes 424 – protein glycation 427 agriculture, signiﬁcance 482 ALBP-HSL complex 151 AMP-activated protein kinase (AMPK) 201 – activity 113, 216 – dependent phosphorylation 217 androgens, role 377

angiogenic factors 294 – hepatic growth factor (HGF) 294 – leptin 294 – vascular endothelial growth factor (VEGF) 294 angiopoietin-like protein 4 (ANGPTL4) 100 angiotensin-converting enzyme (ACE) activity 247 angiotensinogen 247, 249, 255 – adipocyte-speciﬁc overexpression 247 – gene expression, menopausal women 249 – mRNA abundance 248 antibody-based methods 70 – ﬂow cytometry 70 – magnetic immunobeads 70 antidiabetic drugs 317 – thiazolidine-diones (TZDs) 23, 317 ApcMin/þ model 379 apes colonizing, signiﬁcance 483 apo-AV 103 – gene 103 – heparin interaction 103 – role, lipolysis 103 apoptosis 379 – induction 379 – susceptibility, regional variation 32 apoptotic fat cells 32 aquaporins (AQPs) 129, 132, 134, 138, 144 – AQP1, structural analysis 138 – AQP2 protein, phosphorylation site 135 – AQP7 129, 135, 136, 139, 140, 143, 151 – – adipose-derived glycerol 139 – – catecholamine-induced translocation 136 – – coordination 143 – – gluconeogenesis 139 – – insulin-mediated repression 136 – – intracellular glycerol contents 140 – – loss-of-function mutation 139 – – mRNA expression 143 – – nutrition-related changes 135 – – physiological function 139 – – physiological/pathophysiological involvement 143 – – physiological role, schematic presentation 140 – – schematic presentation 143 – AQP9, coordination 143 – discovery 132, 144 – – to physiopathology and clinic 129 – gene expression 134 – gene, single nucleotide polymorphisms (SNPs) 139 – protein, functional pore conformation 138 – role 132

j493

j Index

494

– structural/functional analysis 144 – suppression 144 – tissue distribution 134 atrial natriuretic peptide (ANP) 150, 160, 162 – lipid-mobilizing effect 162 – production 160 atherosclerosis 253 – apolipoprotein E null mouse model 120 autonomic nervous system 153

b Barraquer–Simons syndrome, see acquired partial lipodystrophy (APL) basal metabolic rate (BMR) 335 Berardinelli–Seip syndrome, see congenital generalized lipodystrophy bimolecular ﬂuorescence complementation experiments 92 body fatness, see body mass index (BMI) body mass index (BMI) 21, 59, 184, 249, 288, 369, 429 – cancer risk 370 – effect 370 bone marrow fat cells 23 Borna disease virus (BDV) 314 brainstem, schematic drawing 188 breast cancer 377, 379, 381 – protective factors 381 – risk factors 377, 378 breast tumorigenesis 377. see also breast cancer brown adipocytes 51, 55, 57 – markers, UCP-1/PGC-1a 415 – morphology 51–53 – protein expression 57 – proteins, UCP-1/PPAR-g coactivator 407 brown adipose tissue (BAT) 51–54, 132, 407, 450, 451 – gene expression proﬁling studies 451 – interscapular, buffalo hump 409 – physiology 54 – PPAR-g expression 450 – scanning electron microscopy 51 Bruce Speigelmans laboratory 4

c CAAT/enhancer-binding protein (C/EBP) family 4, 10, 407 – expression 464 – target genes 9 – transcription factors 7 cachexia 334 – adipogenic transcription factors 334 – mRNA 334

– protein, levels 334 calorie restriction (CR) 424 – aging, impact 425 – biological age 432 – cardiovascular disease 427, 430 – chronological age 432 – humans 427–431 – – intervention 432 – – lifespan extension 427, 432, 433 – – longevity 431 – – Okinawa 427 – – randomized controlled trials 429–431 – – survival and energy intake 432 – – Vallejo study 428 – insulin resistance/type 2 diabetes mellitus 427 – mimetics 433 – oxidative stress 426 – rate of living 425 calorie restriction with optimal nutrition (CRON) diet 433 cAMP poduction 152, 157, 163 – adrenergic control 152 – control by adenylyl cyclase inhibitors 157 – Gi-dependent inhibition 163 cAMP signaling pathways 158 cancers 369, 370, 374 – biological mechanisms 371–380 – – adipokines 378–380 – – candidate mechanisms 371 – – IGFs 372, 376 – – insulin, cancer hypothesis 372 – – nuclear factor-kB system 381 – – obesity-related inﬂammatory markers 380 – – oxidative stresses 381 – – sex steroids 376–378 – cachexia, fat-mobilizing substances 338–340 – C-peptide 372, 374, 375 – epidemiology 369–371 – – cancer mortality 370 – – cancer risk 369 – – excess body weight 369, 370 – mechanical mechanisms 381 – new research areas 382 – patients, energy expenditure 335 – risk 370 – serum insulin 374, 375 candidate systems 371 – adipokines 371 – insulin/insulin-like growth factors (IGFs) 371 – sex steroids 371

Index canine distemper virus (CDV) 313 cardiovascular disease 425 – cause 129 – risk factor 254, 369, 428 carnitine palmitoyl transferase (CPT)-1 354 catecholamines 150, 152 – induced lipolysis 152 – – b-adrenergic component 154 b-catenin, stimulation 372 CCAAT/enhancer-binding protein (C/EBP) 213, 231, 463 – transcription factors 463 cell – composition, depot differences 286 – proliferation, depot differences 296 – surface receptors, expression 463 central nervous system 187, 249 – leptin action 187–189 cGMP 160 – dependent protein kinase (PKG) 160 – PKA activation 160 – production 159, 160 – – natriuretic peptides control 159 – – stimulation 160 Chanarin–Dorfman syndrome 87, 152 channel-forming integral membrane proteins 132 – aquaporins (AQPs) 132 chemokines 274 – regulated upon activation of normal tcells expressed and secreted (RANTES) 274 chick embryo lethal orphan (CELO) 315 Chlamydia pneumoniae 320 cholesterol ester transfer protein (CETP) 28 chromatin immunoprecipitation 444 chronic acid reﬂux 382 chronic liver disease (CLD) 205 chronic obstructive pulmonary disease 423 chylomicron (CM) 98, 130 – intestine-derived 130 c-Jun N-terminal kinase (JNK) 216, 360 coactivator proteins 446 colony-stimulating factor-1 genes 274 – expression levels 274 colorectal cancer, risk 379 comparative gene identiﬁcation-58 (CGI-58) 87, 92, 152 complex insulin-cancer hypothesis, schematic representation 373 congenital generalized lipodystrophy 386 congenital lipodystrophies 386 – congenital generalized lipodystrophy 386 – – clinical features 386 – – molecular genetics 390

– familial partial lipodystrophy 390 – – clinical features 390 – – molecular genetics 391 conjugated linoleic acid (CLA) 417 – induced lipoatrophy 418 contemporary populations, implications 485 coregulator proteins 445 corepressor proteins 445 – nuclear receptor corepressor-1 (NCOR1) 445 – receptor-interacting protein-140 (RIP140) 445 – silencing mediator of retinoic acid and thyroid hormone receptor (SMRT) 445 C-reactive proteins (CRPs) 266, 288, 323 C-type natriuretic peptide (CNP) 159 Cushings syndrome 27, 394 cytokines 267, 337, 380, 449 – chemokines 267 – interleukins 267 – – IL-1 337 – – IL-6 164, 337 – tumor necrosis factor (TNF)-a 337

d dehydroepiandrosterone (DHEA) 377, 433 dehydroepiandrosterone sulfate (DHEAS) 377 delipidation process 59 delipidized adipocytes, morphology 61 Delta Serrate Lin12 (DSL) domain 232 de novo fatty acid synthesis 119 diabetes 373, 381 – diabetes mellitus 427 – type 2 – – hyperinsulinemia 373 – – patients 454 diacylglycerol acyltransferase (DGAT) enzymes 411 dietary fat 464 differentiation capacity 296 – depot differences 296, 297 dimeric transcription factors, AP-1 4 dimyristoylphosphatidylcholine (DMPC) liposomes 103 diphtheria toxin A (DTA) 416 DNA – binding transcription factor 451 – bound receptor 444, 445 – – structure/conformation 444 – chips 271 – damage 430 – methyltransferase-1 expression 467 dominant-negative mechanism 392

j495

j Index

496

double-label immunoﬂuorescence analysis 88 dragonﬂies, see Libellula pulchella Drosophila melanogaster 33 Dunnigan–Kobberling segmental lipodystrophy syndromes 24

e ectopic adiponectin expression 211, 212 ectopic fat deposition 348, 354, 424 – producing scenarios 348–355 – – adipose tissue, altered plasticity 349, 350 – – adipose tissue, impaired fat deposition capacity 350–353 – – peripheral tissue, substrate uptake vs. oxidative capacity 353–355 eicosapentaenoic acid (EPA) 340 electron transport chain (ETC) 360 endocrine cell, adipocyte 285 Endogenous Hormones and Breast Cancer Collaborative Group (EHBCCG) 377 endothelial nitric oxide synthetase, phosphorylation 251 enzyme(s) 117, 119, 151, 163, 411 – acetyl-CoA carboxylase 117 – acyl-CoA synthetases (ACSLs) 117 – allosteric inhibitors 119 – energy metabolism pathways 119 European Prospective Investigation into Cancer and Nutrition (EPIC) study 377 extracellular signal-regulated kinase (ERK) pathways 161, 235, 337, 372

f familial partial lipodystrophy (FPLD) 386, 390, 391 – FPLD2 391, 392 – – causative LMNA mutations, majority 391 – – development 392 – FPLD3 392 – – pathogenesis, dominant-negative/ haploinsufﬁciency mechanism 392 fasting-induced adipocyte factor (FIAF) 321 fat 473, 476 – apoptosis 416 – deposition, ontogeny 477 – depots, preadipocyte subpopulations, differences 32, 33 – distribution 23, 395 – – diversity 23, 24 – – types 395 – ethological approach 476–482 – evolutionary approach 473 – speciﬁc protein, CIDEA/CIDEC 85

fat cells 27, 29, 285. see also adipocytes – physiology 27–34 – progenitors 29 – triacylglycerols, hydrolysis regulation 151 – – adipose tissue triglyceride lipase 151 – – hormone-sensitive lipase 151 – – lipases, role 151 fat tissue 21–25 – cell dynamics 22 – function 21–23 – growth 24, 25 – – mechanisms 25 – – regional differences 24, 25 fatty acid(s) 109, 111, 113, 117, 118, 120, 121 – activation/channeling 113–116 – – fatty acid transport proteins 113 – – long-chain acyl-CoA synthetases 113 – contribution 120 – FABP-4-mediated transport 111 – homeostasis 120 – intracellular transport 109 – isotopes 111 – metabolism 118 – – Randle hypothesis 357 – partitioning 117, 121 – – ACSLs/FATPs control 117 – regulation/function 118 – trafﬁcking 119, 121 – – intracellular 109 – – proteins, expression 121 fatty acid-binding protein (FABP) 109–113, 131, 289 – FABP-4 110–112 – – function/regulation 110–112 – – promoter region 112 – FABP-5 112 – – function/regulation 112 – – overexpression 112 – – transgenic/knockout models 112 – isoforms 110, 113 – – expression 113 – – protein structures 110 – members 109 – transportation mechanism 110 fatty acid synthase (FAS) 10, 357 fatty acid translocase (FAT) 11, 131 fatty acid transport protein (FATP) gene 11, 113, 131 – FATP-1, knockdown cells 116 – FATP-4 expression 116 – isoforms 113 fatty liver dystrophy mouse, phenotype 412

Index 8–FK506-binding protein (FKBP) fusion protein 416 ﬂuorescence resonance energy transfer (FRET) 89 Framingham algorithm 431 free fatty acids (FFAs) 83, 111, 132 – transport 132

g gene – coding regions 444 – deserts 444 – expression proﬁles 116 genetically engineered mouse models 408 – A-ZIP/F1 transgenic mouse 408 genetic/hormonal mechanisms 254 genetic variability 475 genomic DNA sequence analysis 392 glucagon-like peptide (GLP)-1 166 glucocorticoids 233, 464 – receptors 233 glucose – metabolism, randle hypothesis 357 – tolerance 424 – transporter 1 (GLUT1) 322 – transporter 4 (GLUT4) 130, 334 – – expression 290 – – mRNA 334 – – vesicles, stimulation 114 glycerol 141, 142 – based gluconeogenesis 143 – oral administration 142 – plasma levels 142 glycerol kinase, enzymatic activity 140 glycerol phosphate acyltransferase (GPAT) enzymes 411 glycosylphosphatidylinositol-anchored highdensity lipoprotein-binding protein 1 (GPIHBP1) 97, 101, 102, 104 – expression 102 – role, triglyceride-rich lipoproteins, lipolysis 101, 102 G-protein complex 165 G-protein-coupled receptors (GPCRs) 158, 215 granulocyte-macrophage colony-stimulating factor (GM-CSF)-1 293 growth hormone 163, 164 – lipolytic sensitivity 164 – physiological contribution 163 – receptors 163 Gs/Gi-proteins, expression level 153 gut microbiota 320, 321 gut parasites 321, 322

h haploinsufﬁciency mechanism 392 helix-loop-helix transcription factor 7 heparan sulfate proteoglycans (HSPGs) 102 hepatic AQP9 143 – physiological/pathophysiological involvement, schematic presentation 143 hepatic glycerol channel 141, 143 – AQP9 141 – coordination 143 hepatic glycerol metabolism 142 – AQP9, role 142 hepatocyte growth factor (HGF) expression 294 heterotrimeric G-proteins 152 hexosamine biosynthesis 185 high-density lipoprotein (HDL) 99, 429 histone deacetylases (HDAC)-independent mechanisms 446 HIV-1 infection 193 – highly active antiretroviral therapy (HAART) 193 HIV-related lipodystrophy 386, 394, 395 – endocrinologic manifestations 395 homeostasis model assessment (HOMA) 372 Homo sapiens 482 hormone-sensitive lipase (HSL) 11, 84, 131, 151, 252 – activation 151 – activity 111 – vs. ATGL, biological signiﬁcance 88 – metabolic role 151 – triacylglycerol, hydrolysis 252 hormone-stimulated lipolysis 27, 86 – omental cells 27 hormone systems 167 – catecholamines 167 – natriuretic peptides 167 Hox genes 33, 34 – expression 34 – function 34 human 61, 62, 192 – adiponectin, domain structure 203 – adipose organ 61–63 – adipose tissue, cytokine families 268 – AQP7 gene 137, 138 – – genetic mutations 138 – fat cell – – adrenergic receptor distribution, heterogeneity 155 – – lipolysis, control pathways 150 – growth pattern 481 – leptin therapy 192, 193 – – congenital leptin deﬁciency 192, 193

j497

j Index

498

– – hypothalamic amenorrhea 193 – – lipodystrophic leptin deﬁciency 193 – obesity 183, 192 – – characterization 183 – – related hypertension, adipocyte RAS 249 – subcutaneous WAT 62 human primary adipose tissue-derived stem cells (hASCs) 317, 318 human telomere reverse transcriptase (hTERT) 30 Hutchinson–Gilford progeria syndrome (HGPS) 391, 396 – clinical features 397 d-b-hydroxybutyrate, antilipolytic effects 158 15(S)-hydroxyeicosatetraneoic acids (HETEs) 338 11 b-hydroxysteroid dehydrogenase (HSD)1 27, 248 hyperplasia 59 hyperplasty 349 hypertension 245 hypertriglyceridemia 141 hypertrophy 59 hypoglycemia 139 hypothalamus, schematic drawing 188 hypoxia-inducible factor (HIF)-1a 272

– binding protein (IGFBP) 33, 376, 465 – – potent inhibitor, PAPPA 33 – receptor 372 – synthesis 378 – system 376 – – ligands 376 – – receptors 376 insulin receptor substrate (IRS)-1 159, 358, 362 – expression 159 insulin response element (IRE) 137 integral endoplasmic reticulum membrane protein 398 intra-abdominal fat depots 286 intracellular cAMP 131 – glucose-6-phosphatase 136 – phosphoenolpyruvate carboxykinase (EPCK) 136 intracellular kinases 216 – c-Jun N-terminal kinase 216 – mitogen-activated protein kinase (MAPK) 216 intracellular lipids – de novo synthesis/hydrolysis 109 – transport proteins, role 109 intramyocellular lipids (IMCL) 355

i

j

IxB kinase (IKK)-b pathways 360, 381 immune system 479 immunoﬂuorescence microscopy 101 immunogold electron microscopy studies 135 inﬂammatory gene expression proﬁle 276 inﬂammatory markers, modiﬁcation 267 inherent cell dynamic mechanisms 28 – regional differences, contribution 28, 29 in situ microdialysis technique 153, 154 insulin 252 – antilipolytic agent controlling cAMP degradation 158 – antilipolytic effect 159 – dephosphorylation 159 – Ras/mitogen-activated protein kinase (MAPK)-dependent signaling effects 252 – role 158 – sensitivity 430 – sensitizing agent 379 – sensitizing drugs, thiazolidinediones 8 – signaling cascade, PDE-3B 159 – signaling pathways 251 insulin-like growth factor (IGF)-1 33, 372, 376, 378, 463 – actions 376

janus kinase (JAK)-dependent signaling 187 – STAT signal transduction pathway 189

k keratinocyte lipid-binding protein (KLBP) 289 knockout/transgenic animal models, striking phenotypes 121 Kruppel-like factors (KLF) family 6, 7 – zinc ﬁnger transcription factors 6

l lean mass index, sex differences 477 leptin 183–186, 192, 193, 249, 265, 378, 465 – cardiovascular effects 250 – effects 250, 466 – gene expression 185, 378, 465 – immunoreactive 59 – metabolic actions 250 – mRNA 186 – receptor 183 – – isoform, Ob-Ra 187 – synthesis 184–186 – therapy 192, 193 Libellula pulchella 321 ligand-activated DNA-binding proteins 442

Index ligand-dependent transcription factor 377 lipases, expression level 153 lipid – biosynthetic enzyme 398 – deﬁnition 109 – droplet-associating proteins, see PPAR-g target genes – homeostasis, white adipose tissue 348 – metabolic pathways, substrates 118 – metabolism, deregulation 167 – pathways 364 – species, accumulation 359 – vacuole 52 lipid mobilization 152, 156, 159, 162 – adrenergic control 152 – a2-adrenergic receptor-dependent modulation, demonstration 156 – induction by pharmacological doses, administration of ANP 162 – natriuretic peptides control 159 – physiological control, ANP, contribution 162 lipid-mobilizing factor (LMF) 166, 335 – cachexia-inducing tumors 166 lipin-1 411, 413 – molecular function 413 – protein 411, 413 lipodystrophic models 407, 411, 414 – animal models, leptin replacement therapy 193 – enhanced energy expenditure 414 – – FOXC2 transgenic mouse 415 – – leptin transgenic mouse 414 – – PPAR-d transgenic mouse 415 – impaired adipogenesis 407–411 – – aP2-SREBP-1c transgenic mouse 408 – – A-ZIP/F1 transgenic mouse 408 – impaired triacylglycerol biosynthesis 411– 414 – – AGPAT6-deﬁcient mouse 412 – – DGAT1-deﬁcient mouse 412 – – GPAT1-deﬁcient mouse 411 – – lipin-1-deﬁcient mouse 412 lipodystrophy 193, 350, 385, 387, 393, 395, 397, 403, 409, 413, 418 – acquired/genetic 385 – cause 409 – classiﬁcation 386 – elucidation mechanisms 404 – etiology 403 – features in forms 387–390 – genes 393 – Hutchinson–Gilford progeria syndrome 396

– – clinical features 396 – – molecular genetics 397 – mandibuloacral dysplasia 395 – – clinical features 395 – – molecular genetics 395 – metabolic consequences 418 – mouse models 403 – – altered PPAR-g levels 409 – – C/EBPa -deﬁcient mouse 410 – – metabolic characteristics 405, 406 – – physiological mechanisms 403–407 – – Zmpste24-deﬁcient mice 410 – neonatal progeroid syndrome 396 – – clinical features 396 – – molecular genetics 396 – overview 385 – short syndrome 396 – – clinical features 396 – – molecular genetics 396 – Werner syndrome 397 – – clinical features 397 – – molecular genetics 397 lipolysis 92, 149, 150, 152, 159, 167, 449 – activators 167 – adrenergic control 152 – adrenergic stimulation 449 – initiation, protein trafﬁcking 92 – insulin, role 149 – natriuretic peptides control 159 lipolytic pathways 163–166 – growth hormone 163 – IL-6 164 – lipolytic peptides 166 – TNF-a 165 lipolytic protein trafﬁcking 88–91 – ATGL, interactions 89, 90 – CGI-58, interactions 88, 89 – HSL, interactions 89 – lipid droplets 90 – – dispersion/disruption 90 – PLIN subcellular targeting 88 lipoprotein lipase (LPL) 97–99, 11, 97, 130, 289, 290, 336, 353, 354 – activity 290 – expression, regulation 97 – functional domains 99, 100 – overexpression 354 – tertiary folding pattern 99 lipotoxicity 350, 355, 360, 363, 453 – affecting factors 360–363 – – adipocytokines proinﬂammatory activity 361 – – adipose tissue macrophages 361 – contributing mechanisms 355–360

j499

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500

– – peripheral organs 355–360 – hepatic lipotoxicity 362, 363 – signaling effector pathways 361, 362 – in skeletal muscle 355, 356 – target, pancreatic b-cell 363 lipotoxicity-related metabolic networks 363 – identiﬁcation, computational methods 363, 364 lipoxygenase (LOX) 338 liver acyl-CoA synthetase 141 lipid-mobilizing factor (LMF), activity 339 low-density lipoprotein (LDL) 427, 430, 431 – cholesterol 430, 431

m macrophages 270 – colony-stimulating factor 323 – inﬁltration, aspect 271 magnetic resonance imaging (MRI) characterization 385 mammalian AQPs 133 – phylogenic analysis 133 mammalian body 130 – adipocytes/gycerol metabolism, characteristics 130–132 mammalian transcription factors, STAT family 5 mandibuloacral dysplasia (MAD) 395 maternal calorie restriction 462 maternal fatness, role 484 maternal high-fat feeding 462 maternal obesity, pregnancy 461 melanin-concentration hormone (MCH) 313 melanocortin-4 receptor (MC-4R) 191 – antagonism 191 a-melanocyte-stimulating hormone (MSH) 250 membrane proteins 114 mesenchymal stem cells (MSCs) 71 metabolic ghetto 485 metabolic syndrome 347, 385 microdialysis probe 155, 162 mitogen-activated protein kinases (MAPKs) 290, 291 – inhibition 161 – pathways 235, 337 – signaling 252 monocyte chemoattractant protein-1 (MCP-1) 253, 272, 274, 293 – endothelial cells 253 – expression levels 274 mouse embryonic ﬁbroblasts (MEFs) 234 mouse models 416

– acquired/conditional lipodystrophy 416–418 – – aP2-DTA transgenic mouse 416 – – diet-induced lipoatrophy 417 – – drug-induced lipoatrophy 417 – – FAT-ATTAC transgenic mouse 416 – – RSK2 deﬁcient mouse 417 mRNA expression 467 multisystem syndromes 385 – progeroid syndromes 385 – SHORT syndrome 385 murine FAS promoter 12 – STAT-5A-binding site 12 muscle lipotoxicity 356 – generation, molecular mechanisms 356–360

n N-acetyl cysteine (NAC) 212 NAD-dependent deacetylase, sirtuin-1 210 National Center for Biotechnology Information AceView 392 natriuretic peptides 150, 159, 161 – atrial natriuretic peptide (ANP) 150 – brain natriuretic peptide (BNP) 150 – in vitro/in vivo effects 161 – lipolytic effect 161 natriuretic peptide receptor (NPR)-A receptor antagonist 163 natural selection 474 Neels thrifty genotype hypothesis 474–476 neurogenesis 60 nicotinic acid 158, 167 – dyslipidemia, treatment 158 nonadipose tissues 424 nonalcoholic fatty liver disease (NAFLD) 205 nonesteriﬁed fatty acids (NEFA) 152, 158, 251, 336, 339 – plasma levels 251 – stimulators 152 – synthesis 336 noradrenergic parenchymal ﬁbers 55 norepinephrine 25, 152–154, 256, 451 nuclear factor-kB (NF-kB) pathway 381 nuclear hormone receptor (PPARG) 398 nuclear localization sequence (NLS) 391 nuclear receptors 442 – peroxisome proliferator-activated receptor (PPAR)-g 407 – type II class 442 nucleus of the solitary tract (NTS) 188 – leptin receptors 188 neuropeptide Y (NPY) receptor 157 – PYY receptor 157

Index

o obesity 3, 189, 190, 245, 266, 274, 314, 424 – adipose AQP7/hepatic AQP9 142 – – overexpression 142 – biological mechanisms 371–380 – – adipokines 378–380 – – candidate mechanisms 371 – – IGFs 372, 376 – – insulin-cancer hypothesis 372 – – insulin/C-peptide 372 – – nuclear factor-kB system 381 – – obesity-related inﬂammatory markers 380 – – oxidative stresses 381 – – sex steroids 376–378 – characterization 266 – complications 274, 275 – developmental origins 153, 460, 462 – – animal models 461, 462 – – evidence 460–462 – – humans 461 – – theories 460 – diet-induced 113 – epidemiology 369–371 – – cancer mortality 370 – – cancer risk 369 – – excess body weight 369, 370 – high-fat diet-induced obesity 434 – hyperleptinemia, complications 190, 191 – – leptin, pathologic effects 191 – – obesity-related hypertension 190, 191 – importance for 154 – infectious origin 307 – insulin resistance 143, 434 – – AQP7/AQP9, dysregulation 143 – leptin resistance 189, 190 – over-consumption of energy dense foods 424 – patients 249 – – arterial hypertension 249 – – candesartan role 249 obesity-related hypertension 132, 245–247, 248 – adipose tissue 245–254 – – adiponectin 250, 251 – – blood pressure control 245 – – concept 246 – – free fatty acids 252, 253 – – 11b-HSD-1 254 – – insulin 251, 252 – – leptin 249, 250 – – plasminogen activator inhibitor-1 252 – – RAS 247–249 – – regional distribution 254

– – resistin 253, 254 – animal models 254 – hemodynamic characteristics 245 – molecular mechanism 132, 246 – perivascular adipose tissue, potential role 155, 156 – visceral adipose tissue 255 obesogenic environment 486, 487 obesogenic niche 487 ob/ob obese diabetic mouse model 120 Ob-Rb mRNA, expression 187 omental depots 297 – FABP-4 297 – GLUT4 297 – leptin 297 omentin, C-type lectin 291 overweight, see obesity

p P2-UCP-1 transgenic mice 53 – white adipocytes 53 paracrine/autocrine mechanism 164 parathyroid hormone (PTH) 163, 166 – lipolytic pathways 166 PAT protein 85, 86 – biology 85 peptides, lipolytic potency 160 perilipin (PLIN) 84, 88 – phosphorylation 89 perivascular adipose tissue 256 peroxisome proliferator-activated receptor-c (PPAR-c) 4, 8, 9, 11, 12, 58, 112, 119, 134, 231, 392, 409, 430, 441, 442, 445, 447, 449–454, 467 – activation 119, 441 – agonists, effects 453 – coactivator 430 – dependent proliferation 453 – diverse effects 451 – DNA-binding domain 443 – expression 9, 409 – function in adipose tissue maintenance/ remodeling 452 – gene structure 443 – haploinsufﬁciency 409 – heterodimer formation 134 – importance 441 – ligands 112 – – binding domain 442 – master regulator of adipose tissue development 447–449 – – adipogenesis, PPAR-g, role 447 – – in vivo adipose tissue development 448 – metabolic functions 449

j501

j Index

502

– molecular biology 442–446 – – DNA binding 442–444 – – PPAR-g structure 442–444 – molecular/cellular function 441 – null mice 8 – phosphorylation 8 – proliferative effects 453 – receptors 381 – response elements 442 – role 9, 409, 441, 448, 454 – RXR-a complex 135 – – heterodimer, structure 442, 447 – – speciﬁc binding 135 – synthetic ligands 452 – – thiazolidinedione (TZD) drugs 452 – target genes 11, 12, 446, 449 – – downregulated expression 449 – transcriptional activation 445 – transcriptional partner 450 – transcriptional regulation 444 – transcription factors 464 – white vs. brown fat-speciﬁc functions 450 peroxisome proliferator response element (PPRE) 11, 134, 444 – heterodimer 134 phosphatidate phosphatase (PAP)-1 411. see also lipin-1 phosphoinositide-3-kinase (PI3K) activity 356 – pathways 372 physical activity level (PAL) 335, 340 plasma membrane, phosphorylation 114 plasminogen activator inhibitor (PAI)-1 252 – deﬁciency 252 – obesity 252 – pharmacologic inhibition 252 point mutations 410 polycomb group (PcG) proteins 34 polycystic ovary syndrome (PCOS) 385 polymorphisms 137, 154 – types 137 polyunsaturated fatty acids (PUFAs) 338 postmenopausal breast cancer 376 potential antilipotoxic therapeutic strategies 365 preadipocytes function 21, 29, 31–33 – cell lines 3 – differences 33, 34 – gene expression proﬁles 33 – replication 29, 30 Pref-1 232–236 – expression, regulation 233 – forms 233 – function 232–236

– in vivo effect 235, 236 – – adipogenesis 235, 236 – – glucose/insulin homeostasis 235, 236 – isoforms, domain structure 232 – mechanism 234, 235 – null mice 235 – structure 232, 233 – synthesis 232 pregnancy associated plasma protein A (PAPPA) 33 presenillin-associated rhomboid-like protein 430 proinﬂammatory cytokines 214, 291, 361 – CCL2 361 – interleukin (IL)-1 361 – interleukin-6 291, 361 – tumor necrosis factor (TNF) 291 promoter analysis 142 pro-opiomelanocortin (POMC) 187 – neuron-speciﬁc leptin receptor 191 protease inhibitors 413 protein/DNA complexes 444 protein kinase A (PKA) 84, 135, 151 – activation 83, 90, 91 – adipocyte lipolysis, regulation 84 – lipolytic protein trafﬁcking, model 91 protein kinase C (PKC) isoforms 358 protein kinase G (PKG) 151 protein phosphatase 2A (PP2A) 360 proteins 152, 407 – complementation experiments 89 – esterase/thioesterase/lipase subfamily 152 – roles 407 protein tyrosine phosphatases (PTPs) 159, 190 proteomic analysis 91 pulse-chase experiments 209 pyruvate dehydrogenase kinase (PDK)-4 13 – glycolysis, regulator 13

r rat interscapular, brown adipose tissue (BAT) 53 reactive oxygen species (ROS) 426 – generation 427 receptor protein 160 – guanylyl cyclase function 160 receptor-speciﬁc agonists 441 – thiazolidinedione (TZD) drugs 441 regenerative medicine 73 – differentiation 73, 74 – – ectodermal lineages 73, 74 – – endodermal lineages 73, 74

Index – – mesodermal lineages 73 – potential utility 73, 74 regional adipose deposition 254 – blood pressure regulation 254–256 regulated upon activation of normal T cells expressed and secreted (RANTES) expression 294 regulatory mechanisms, schematic presentation 137 renin-angiotensin system (RAS) 245 – obesity-related hypertension 255 resistin 238, 253 – acute infusion 238 – glucose homeostasis, regulation of 253 – physiological role 238 resting energy expenditure (REE) 335 retinoic acid X receptor (RXR)-a 134, 442 ribosomal S6 kinase 2 (RSK2) 417 RNA expression 443 RNA interference (RNAi) 139 Rous-associated virus (RAV)-7 314 – characterization 314 – obesity 314

s scrapie agents 322, 323 serum amyloid A (SAA) 268, 293 – isoforms 268 serum leptin 184, 185, 189 – determining factors 184–186 – – body fat distribution 184 – – caloric intake 185 – – gender 184 – – glucose 185 – – insulin 185 sex steroids 376–378 – androgens and breast cancer 377 – estrogen and breast cancer 376 – sex steroids and endometrial cancer 377 SHORT syndrome 396 signal transducers and activators of transcription (STAT) proteins 5, 6, 12, 13 single nucleotide polymorphisms (SNPs), effects 120 skeletal muscle antioxidant capacity, adaptations 426 social inequality, signiﬁcance 485, 486 soluble N-ethyl maleimide-sensitive fusion protein (SNARE) 211 stearoyl-CoA desaturase (SCD) genes 10 stem cell, deﬁnition 69 sterol regulatory element-binding protein (SREBP) 4, 7, 10, 117, 334, 362, 408, 409 – expression 409

– insulin-modulated transcription factor 7 – nuclear form 408 – target genes 10, 11 streptozotocin (STZ)-induced insulin deﬁcient mice 136 stromal vascular fraction (SVF) 269, 270 subcutaneous depots 50 subcutaneous vs. visceral adipocytes 27 suppressors of cytokine signaling (SOCS) 189, 362 sympathetic nervous system (SNS) 25, 248, 249

t 3T3-L1 cells 3, 5, 116, 214, 237, 463 – adipocytes, differentiation 133 – hairpin RNA knockdowns 116 – preadipocytes, lipid accumulation 317 – STAT expression 5 – treatment 237 thiazolidinedione (TZD) drugs 31, 201, 213, 441, 450–452, 454 – glucose-lowering effects 452 – induced differentiation 450 – pioglitazone 213 – therapy 451, 452, 454 thrifty phenotype hypothesis 460, 476 – schematic presentation 460 thrombospondin (TSP)-1 295 thyroid hormone receptor-associated protein complex 446 Tinbergens approach, perspectives 476 – causes 476 – evolutionary history 476, 480–482 – ﬁtness value 476, 478–480 – ontogeny 476, 477 tissue hypoxia 272 toll-like receptors (TLR-4) 273 total energy expenditure (TEE) 335 transcription – activators 5, 6 – factors 413, 415 – – winged-helix/forkhead family 415 transgenic mouse model, characterization 414 triacylglycerol – biosynthetic enzyme 411 – hydrolysis, intracellular fatty acids, role 118 – synthesis, ACSL-1 role 115 tumor – bearing animals, requirements 338 – development 376 tumor necrosis factor (TNF)- a 21, 150, 165, 251, 265, 273

j503

j Index

504

– converting enzyme 233 – induced lipolysis 165 – mechanisms of action 165 – receptors 165 tumor necrosis factor-a-converting enzyme (TACE) 233 – lentivirus-mediated overexpression 233

u ubiquitin-like protein 8 – small ubiquitin-like modiﬁer 8 uncoupling protein (UCP)-1 53, 236, 336, 407, 451 – gene expression 12, 62, 407 – syndrome 236

v

vascular noradrenergic ﬁbers 54 vasculogenesis 60 very-low-density lipoprotein (VLDL) 97, 98, 130 viruses 313–320 – adenoviruses 315–320 – – Ad-5 319 – – Ad-36 316–319 – – Ad-37 319 – – adipogenic potential 319, 320 – – SMAM-1 315, 316 – Borna disease virus 314, 315 – canine distemper virus (CDV) 313 – Rous-associated virus-7 314 visceral adipocytes 27

visceral adipose tissue 254, 287 – metabolic activity 254 – portal drainage, consequences 287 visceral fat cells 27 visceral fat depots 287 – deﬁnition 287, 288 visceral obesity 26 – metabolic impact 26 visfatin 291

w weight gain, see obesity Werner syndrome 397 Western industrialized niche 486 white adipocytes 51, 54 – morphology 51–53 white adipose tissue (WAT) 51, 52, 54, 56, 60, 132, 265, 333, 348, 410, 450 – adipocyte precursors 56 – physiology 54, 55 – retroperitoneal 60 – scanning electron microscopy 51 whole-body energy metabolism 121 Wiedemann–Rautenstrauch syndrome 396

x Xenopus oocytes 141

z

zinc a2-glycoprotein (ZAG) 166, 339, 340 zinc metalloproteinase 395, 398 Zmpste24-deﬁcient mouse model 410