Integration of Metabolism, Energetics, and Signal Transduction (Bioengineering, Mechanics, and Materials: Principles and Applications in Sport S.)

Integration of Metabolism, Energetics, and Signal Transduction Integration of Metabolism, Energetics, and Signal Tran...

Author: Robert K. Ockner

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Integration of Metabolism, Energetics, and Signal Transduction

Integration of Metabolism, Energetics, and Signal Transduction Unifying Foundations in Cell Growth and Death, Cancer, Atherosclerosis, and Alzheimer Disease Robert K. Ockner, M.D. University of California, San Francisco San Francisco, California

KLUWER ACADEMIC PUBLISHERS NEW YORK, BOSTON, DORDRECHT, LONDON, MOSCOW

eBook ISBN: Print ISBN:

0-306-48529-X 0-306-48471-4

©2004 Springer Science + Business Media, Inc. Print ©2004 Kluwer Academic/Plenum Publishers New York All rights reserved No part of this eBook may be reproduced or transmitted in any form or by any means, electronic, mechanical, recording, or otherwise, without written consent from the Publisher Created in the United States of America

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DEDICATION

This book is dedicated to my wife Elaine; my sons Jim, Matt, and Peter; my brother Steve; and to my parents, the late Lee and Sara Ockner.

v

PREFACE

Complex and unexplained phenomena tend to foster unorthodox perspectives. This publication is an example, as is a prior publication that emphasized the concept that intermediary metabolism might play a significant and determining role in hepatocyte proliferation and tumorigenesis1. Formulation of this hypothesis was based on an attempt to clarify several poorly understood phenomena; including the observations: 1) that xenobiotic peroxisome proliferators such as the fibrate hypolipidemic agents induce hepatocyte proliferation and carcinogenesis in rodents; 2) that benign and malignant liver tumors complicate the human syndrome of glycogen storage disease type I (glucose-6-phosphatase deficiency); and 3) that in this same syndrome, administration of glucose exerts an anti-tumor effect. Fatty acid and glucose metabolism are tightly linked in a wellestablished and profoundly inportant interplay. This connection, together with the fact that peroxisome proliferator-induced hepatocyte proliferation and carcinogenesis reflects inhibition of mitochondrial carnitine palmitoyltransferase-I and fatty acid oxidation, suggested the possibility that regulation of fatty acid metabolism could prove to be a pivotal determinant in the control of cell growth. In 1993, the year in which the paper cited above was published, insight into the importance of growth factors and signal transduction pathways in cell cycle regulation was increasing rapidly, but metabolic and energetic aspects of cell proliferation had attracted relatively little attention. Despite this, the concept seemed inescapable that the two seemingly distinct and unrelated determinants — signal transduction and metabolism — were integrally linked. Moreover, it is known that growth regulation in the earliest eukaryotes was governed largely or exclusively by nutritional, metabolic, and energetic factors hundreds of millions of years before the appearance of intercellular signaling in higher organisms. It seemed vii

viii

Preface

plausible, therefore, that these factors had retained a dominant regulatory role, at least under certain conditions, throughout the evolution of today’s more advanced and complex species. What has emerged through the development of this treatise is evidence strongly suggesting that metabolism and energetics, while at times referred to as “housekeeping” functions, are intimately and directly linked to those signal transduction pathways that are essential to survival of the cell, the organism, and the species. Initial work on the project (later to become Part II of the present volume) involved review of published contributions of many insightful and productive scientists. These addressed a broad range of relevant issues, including intermediary metabolism, signal transduction, and cell cycle regulation, and the effects of alternative substrates on mitochondrial energetics. Our preliminary 2 and unpublished experiments addressed the effects of endogenous and xenobiotic growth modulators on fatty acid metabolism and mitochondrial function, and provided critically important early insight and stimulus. Unexpectedly, it also became appreciated during this time that, despite suppression of glucose utilization in brain regions affected by Alzheimer disease, neurons in those regions remained viable. Viewed in the context of the current project, this otherwise seemingly unrelated observation suggested the possibility that utilization of alternative substrates might also prove to be a critical determinant in the energetics, function, and injury of the neuron. Pursuit of published research related to this hypothesis led to the recognition of what appeared to be important parallels in intermediary metabolism and signal transduction between neuronal activation (Part III of this volume) and cell proliferation (Part II). Initially, Parts II and III were developed as separate analytical reviews. With the passage of time, however, their sustained growth in size, scope, and relatedness made separate publication less feasible and less desireable. As a result, they are included in the present combined format, along with short introductory and concluding sections (Parts I and IV, respectively) that provide overall perspective. As work on the project progressed, it required ongoing review of increasing numbers of new publications in diverse fields; those findings most relevant to the integrated approach were selected for incorporation. Encouragingly, few published reports offered serious challenge to the present interpretations and hypotheses; of those, the more important are cited and discussed. Moreover, with continuing development of the project, its fundamental concepts became applicable to a growing number of surprisingly diverse additional areas, ranging from an early focus on liver regeneration to include programmed cell death 3 , cancer cachexia, atherosclerosis, ischemia-reperfusion injury, regulation of feeding behavior, aspects of synaptic function, and the pathogenesis of Alzheimer disease.

Preface

ix

While their broad relevance has provided additional support to many of the concepts, it has also required considerable stringency in the selection of publications for citation. As the regrettable but inevitable result, it was not possible to include reference to numerous other excellent publications that seemed less related to the present focus or less essential to its development. In any case, the project’s very nature dictates that it will remain a work in progress.

REFERENCES 1. R. Ockner, R. Kaikaus, and N. Bass, Fatty acid metabolism and the pathogenesis of hepatocellular carcinoma: review and hypothesis, Hepatology 18:669-676 (1993). 2. R. Ockner, N. Lysenko, N. Wu, and N. Bass, Hepatocyte growth inhibitors modulate mitochondrial and extramitochondrial fatty acid oxidation, Hepatology 24:253A (Abstract) (1996). 3. R. Ockner, Apoptosis and liver diseases: Recent concepts of mechanisms and significance, J Gastroenterol Hepatol 16:248-260 (2001).

ACKNOWLEDGEMENTS

I thank my bright, creative, and collegial former associates in fatty acid and fatty acid binding protein research at the University of California, San Francisco (UCSF), among them Nathan M. Bass, Ruth Brandes, David A. Burnett, Alfred Gangl, John L. Gollan, Albert L. Jones, Raja M. Kaikaus, Richard A. Weisiger, Nina Lysenko, Joan A. Manning, and, in collaborative projects, David H. Alpers, Jeffrey I. Gordon, and E. Raghupathy. I also wish to express appreciation for encouragement in the present project by other colleagues at UCSF: Rudi Schmid, D. Montgomery Bissell, Stephen DeArmond, Richard J. Havel, the late Ira Herskowitz, John Kane, Young S. Kim, Robert Mahley, Frank McCormick, Bruce Miller, R. Curtis Morris, and Marvin H. Sleisenger. Members of the UCSF Liver Center, Division of Gastroenterology, and Comprehensive Cancer Center have represented an ongoing and congenial source of information and stimulation. Sadie McFarlane provided expert assistance in preparation and formatting of the manuscript; she also accomplished the digital transformation of my original drawings to clear and informative illustrations. The project was supported in part at its inception by research grant R01 AM13328, and throughout its development by Liver Center Grant P30 DK26743, both from the National Institutes of Health. Figures 1 and 2 from Journal of Gastroenterology and Hepatology, 2001;16:248-260, are used with the permission of Blackwell Publishing. Finally, to the extent that my time and energies became increasingly devoted to this project, others in the Division of Gastroenterology and Liver Center have carried a correspondingly increased workload; their efforts are truly appreciated.

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CONTENTS

PART I: PROLOGUE

1

1. DISPARATE THEMES: ORIGINS AND INTEGRATION

3

PART II: FATTY ACIDS, MITOCHONDRIA, AND SIGNAL TRANSDUCTION: INTEGRATED CONTROL OF CELL PROLIFERATION, INJURY, AND DEATH 2. INTRODUCTION TO PART II 3. NUTRIENT AND ENERGY METABOLISM IN CELL PROLIFERATION 3.1. INTRODUCTION 3.2. INTERMEDIARY METABOLISM: GENERAL CONSIDERATIONS 3.3. ATP GENERATION, CARBOHYDRATE METABOLISM, AND CELL GROWTH 3.3.1. Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation 3.3.2. Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable 3.3.2.1. Broader Significance of wg/Wnt-Like Signaling 3.3.2.2. Lithium 3.3.2.3. Reactive Oxygen Species 3.4. REFERENCES 4. FATTY ACIDS AND GROWTH REGULATION 4.1. INTRODUCTION 4.2. OMEGA-6 FATTY ACIDS FATTY ACIDS 4.3. OMEGA-3 4.3.1. Metabolic Effects 4.3.2. Growth Effects

xiii

11 13 19 19 19 22 23 27 31 33 33 35 47 47 48 51 51 52

Contents

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4.4. FATTY ACIDS, MODULATION OF CELL GROWTH, AND THE AMP-ACTIVATED PROTEIN KINASE 4.4.1. AMP-Activated Protein Kinase 4.4.2. Stoichiometric Considerations 4.5. METABOLIC INTERACTION BETWEEN TUMOR AND HOST: CANCER CACHEXIA 4.6. REFERENCES 5. MITOCHONDRIAL FUNCTION IN CELL GROWTH AND DEATH 5.1. INTRODUCTION 5.2. PEROXISOME PROLIFERATOR-ACTIVATED RECEPTORS (PPAR) AND LIGANDS: MITOCHONDRIAL REDOX,MACROPHAGES, AND MITOSIS 5.2.1. 5.2.2. 5.2.3. 5.3. MITOCHONDRIAL FUNCTION IN APOPTOSIS 5.3.1. The Bcl-2 Protein Family 5.3.2. wg/Wnt-Like Signaling: Convergence of Mitochondria, p53, and Telomerase 5.3.3. Mitochondria and FasL-Independent Activation of Fas 5.3.4. Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer 5.4. REFERENCES 6. METABOLIC EFFECTS OF ANTIPROLIFERATIVE AGENTS 6.1. INTRODUCTION 6.2. BUTYRATE 6.3. TRANSFORMING GROWTH 6.4. SALICYLATES AND OTHER NONSTEROIDAL ANTI-INFLAMMATORY DRUGS (NSAIDS) 6.5. REFERENCES 7. FATTY ACIDS AND MITOCHONDRIA, CELL GROWTH AND INJURY: BROADER IMPLICATIONS 7.1. INTRODUCTION 7.2. ADVERSE EFFECTS OF FATTY ACIDS ON MITOCHONDRIAL FUNCTION AND CELL REDOX BALANCE 7.3. ORIGINS OF INTRAMITOCHONDRIAL OXIDATIVE STRESS 7.4. CONSEQUENCES OF INTRAMITOCHONDRIAL OXIDATIVE STRESS: ATHEROSCLEROSIS AND BEYOND 7.4.1. Atherosclerosis and Arterial Hypertension

53 57 58 61 64 77 77

78 78 84 85 85 88 90 93 95 98 121 121 121 125 127 132 143 143 143 145 148 149

Contents 7.4.1.1. Oxidized LDL 7.4.1.2. The Endothelial Lesion 7.4.1.3. Beneficial Effects of “Statin” Hypolipidemic Agents 7.4.2. Adverse Effects of Fatty Acid Oxidation-Induced ROS in Various Organs 7.4.2.1. Ischemia-Reperfusion Injury and Heat Shock/Stress Response 7.4.2.2. Paradoxical Benefits of Adversity 7.5. REFERENCES

xv 149 150 152 154 155 156 160

8. METABOLISM AND GENE EXPRESSION IN LIVER REGENERATION 8.1. INTRODUCTION 8.2. SYSTEMIC METABOLIC RESPONSES TO 70% HEPATECTOMY 8.3. EARLY CHANGES IN HEPATOCELLULAR METABOLISM AND GENE EXPRESSION 8.4. ENSUING MODULATION OF SYSTEMIC AND HEPATOCELLULAR METABOLISM 8.4.1. Effects of Proinflammatory Cytokines: vs. Insulin 8.4.2. Completion of Regeneration and Cell Cycle Exit 8.5. REFERENCES

184 189 190

9. PART II CONCLUSIONS

201

PART III: FATTY ACIDS, KETONE BODIES, AND BRAIN METABOLISM: IMPLICATIONS FOR NEURONAL FUNCTION AND ALZHEIMER DISEASE

205

10. INTRODUCTION TO PART III

207

11. ENERGETICS OF NEURONAL ACTIVATION 11.1. INTRODUCTION - RELEVANT PRINCIPLES OF INTERMEDIARY METABOLISM 11.2. ENERGY-CONSUMING EVENTS IN NEURONAL ACTIVATION 11.3. MITOCHONDRIAL FUNCTION AND THE GENERATION OF ATP 11.4. REFERENCES

209

12. UTILIZATION OF OXIDIZABLE SUBSTRATES IN BRAIN 12.1. INTRODUCTION 12.2. GLUCOSE 12.3. KETONE BODIES

177 177 177 180 182

209 212 213 214 217 217 218 220

Contents

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12.4. FATTY ACID UTILIZATION IN BRAIN AND THE REGULATION OF APPETITE 12.4.1. Uptake of Fatty Acids from Plasma 12.4.2. Oxidation of Fatty Acids 12.4.3. Fatty Acid Oxidation and Regulation of Feeding Behavior 12.5. CYTOKINES AND THE BRAIN 12.6. REFERENCES

13. ASTROCYTE METABOLISM AND ASTROCYTENEURON INTERACTION 13.1. INTRODUCTION 13.2. CARBOHYDRATE METABOLISM 13.3. AMINO ACID METABOLISM AND THE GLUTAMINE SHUTTLE 13.4. FATTY ACIDS, ANTIOXIDANT DEFENSE, HIV DEMENTIA, AND CELL PROLIFERATION 13.4.1. Astrocyte-Neuron Interaction and Oxidative Stress 13.4.2. HIV Dementia 13.4.3. Cell Proliferation and Tumorigenesis 13.5. REFERENCES 14. NEURONAL ENERGY METABOLISM IN BRAIN: ASTROCYTE AS METABOLIC “BUFFER” AND MEDIATOR OF NEURONAL INJURY 14.1 INTRODUCTION 14.2. ACTIVATION-RELATED GLUCOSE AND OXYGEN CONSUMPTION 14.3. INCREASES IN PLASMA CONCENTRATIONS OF KETONE BODIES AND FFA 14.4. KETOGENIC DIET AND CONTROL OF SEIZURE ACTIVITY 14.5. SYNAPTIC TRANSMISSION, LONG-TERM POTENTIATION, AND EXCITOTOXICITY 14.5.1. Energetic Considerations in Synaptic Transmission 14.5.2. Augmented Glutamate Exposure: Long-Term Potentiation and Excitotoxicity 14.5.2.1. Long-Term Potentiation 14.5.2.2. Excitotoxicity – Origins of Oxidative Stress 14.6. REFERENCES 15. PATHOGENESIS OF ALZHEIMER DISEASE: METABOLIC FACTORS 15.1. INTRODUCTION 15.2. GENETIC DETERMINANTS 15.2.1. Apolipoprotein E Isoforms 15.2.2. Presenilin and Amyloid Precursor Protein Mutations

226 226 228 233 237 241 255 255 255 259 262 264 267 268 269

277 277 278 279 280 282 282 285 285 288 293 303 303 305 305 306

Contents

15.3. ASSOCIATION OF ALZHEIMER DISEASE WITH TYPE II DIABETES MELLITUS 15.4. CEREBRAL METABOLISM AND INSULIN SIGNALING 15.4.1. Glycolysis and the Citric Acid Cycle in Alzheimer Disease 15.4.2. Wingless/Wnt-Like Signal Transduction and Effects of Lithium 15.4.2.1. Wingless/Wnt-Like Signaling 15.4.2.2. Neuroprotection and Cell CycleLinked Neuronal Death 15.4.2.3. Neurochemical Effects of Lithium 15.5. METABOLIC DETERMINANTS IN ALZHEIMER DISEASE: SYNTHESIS 15.5.1. Oxidative Stress and Amyloid: A “Vicious Cycle” of Multiple Interacting Determinants 15.5.1.1. Amyloid 15.5.1.2. Oxidative Stress 15.5.1.3. Genetic Factors 15.5.1.4. Changes in Fatty Acid and Ketone Body Metabolism 15.5.1.5. Neuroprotective Responses and the “Vicious Cycle” 15.5.2. Hypothesized Role of Fatty Acid Metabolism in Neurodegeneration 15.5.3. Integrated Model of Alzheimer Disease Pathogenesis 15.6. REFERENCES

xvii 307 309 310 314 315 316 320 321 321 322 323 323 324 325 326 329 330

16. PART III CONCLUSIONS

355

PART IV: EPILOGUE

359

17. LOOKING BACK, LOOKING AHEAD

361

INDEX

369

PART I: PROLOGUE

Chapter 1 Disparate Themes: Origins and Integration

Consider life as it may have existed a billion or so years ago. Our unicellular eukaryotic forebears drifted in a primordial “sea”. Life was precarious‚ and survival was dictated to a large extent by chance events. Prominent among these was dependence on an unpredictable ebb and flow that might bring the cell into the vicinity of nutrient that was appropriate and sufficient not merely to sustain life but — equally important — enable the cell to divide and give rise to an expanding progeny. It is reasonable to assume that under such favorable circumstances some organisms would have activated a proliferative response more rapidly and efficiently than others‚ and that they would prove to be the more successful in terms of survival and growth in numbers. Similarly‚ when the environment became depleted of nutrient‚ unable to sustain growth‚ or otherwise unfavorable‚ the successful organisms would have been those that rapidly arrested growth‚ surviving in an energy conservation mode while awaiting better times. In short‚ the successful organisms — i.e.‚ those genetically programmed to respond rapidly and appropriately to sudden and unpredictable environmental changes — would write the history of the period. In contrast‚ untold numbers of organisms would fail to execute the dramatic changes in metabolism‚ energetics‚ and growth necessary to “navigate” that perilous and unpredictable sea. These unsuccessful genetic “experiments” would vanish without a trace‚ at the hands of natural selection. What was the distinguishing characteristic that conferred such indispensable versatility and timeliness upon the survivors? To a large extent‚ speculation about this distant scene would be impossible without a profoundly important and informative contemporary window through which we are able to view it. That window is provided by direct descendants of those early eukaryotes‚ exemplified by fungi such as the yeast Saccharomyces cerevisiae (S. cerevisiae). S. cerevisiae is genetically 3

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Chapter 1

programmed to proliferate rapidly if its environment is suitable‚ i.e.‚ if it contains glucose or an acceptable alternative. Conversely‚ when glucose is no longer available S. cerevisiae activates a program encoded by the SNF1 gene that immediately arrests growth. Significantly‚ this growth arrest is linked directly to the inactivation of two major glucose-dependent energyconsuming pathways that are needed to support cell proliferation‚ i.e.‚ those that effect biosynthesis of the fatty acids and cholesterol that are required for assembly of daughter cell membranes. The signals that activate this growtharresting survival-promoting response reflect the direct and unavoidable consequences of nutrient depletion: a drop in cellular [ATP] and the tightly coupled reciprocal rise in [AMP]. As a further consequence of this programmed cessation of growth and of fatty acid biosynthesis‚ fatty acid oxidation becomes the cell’s dominant source of energy. The effectiveness of this mechanism‚ and its importance to survival over more than one billion years since the dawn of eukaryotic life‚ is dramatically demonstrated by its persistence to the present‚ not merely in S. cerevisiae‚ but as expressed through homologous genes and regulatory programs in most higher plants and animals as well. Thus‚ in the cells of human beings and other mammals‚ a nutrient depletion-induced rise in cellular [AMP] activates a kinase cascade that is mediated by the SNF1 homologue AMP-activated protein kinase (AMPK). AMPK arrests growth‚ inactivates fatty acid and cholesterol biosynthesis‚ and augments mitochondrial fatty acid oxidation. Accordingly‚ SNF1/AMPK provides a mechanism for mediating the selection between alternative fuels‚ i.e.‚ between glucose or its equivalent and fatty acids‚ as circumstances may dictate. When this ancient regulatory system first appeared‚ however‚ the choices confronting the primitive unicellular organism were simple in the extreme: consume glucose and proliferate; or‚ failing to find glucose‚ stop proliferating and consume stored fatty acids. Life is more complex now. But as we will see in the following paragraphs and pages‚ this once primitive selection between alternative fuels has acquired in mammals a vastly increased and more versatile repertoire — the reward for its successful evolutionary adaptation to numerous regulatory processes‚ many of which are only faintly suggestive of its remote‚ and exclusively growth-related‚ origins. How is this single example of gene conservation — one that provides survival advantage by rapidly arresting growth in response to an unfavorable and unpredictable change of circumstances — related to the present treatise and to the issues concerning both basic science and human disease that it examines? Briefly answered‚ it provides direct (and to say the least timetested!) evidence supporting several key concepts upon which the treatise is based. These concepts serve to integrate a number of cellular processes which‚ perhaps understandably‚ have generally been regarded as being

1. Disparate Themes: Origin and Integration

5

related only distantly or not at all. Summarized in the following paragraphs‚ these concepts will be developed in greater detail in subsequent chapters: First‚ cellular nutrient metabolism and energetics are linked directly and intimately to the regulation of cell proliferation. In general‚ cell proliferation depends upon aerobic or anaerobic glycolysis to generate the enormous and rapid increases in ATP that are required for completion of the cell (mitotic) cycle and its obligatory accompaniments. The latter include not only biosynthesis of lipids‚ proteins‚ and nucleic acids‚ but also numerous translocational‚ regulatory‚ and signaling processes. Although mitochondrial oxidation of fatty acids also generates ATP‚ this process has certain disadvantages: 1) it may suppress glucose utilization; 2) it may compromise efficiency of mitochondrial energetics and foster cell injury by promoting generation of reactive oxygen species (ROS); and 3) it consumes what otherwise would become the lipid constituents of daughter cell membranes. Therefore‚ it is hypothesized that in proliferating cells‚ fatty acid oxidation is counterproductive and of necessity suppressed. This inverse relationship between cell proliferation and mitochondrial fatty acid oxidation is of paramount importance‚ as are its metabolic and energetic ramifications. More broadly‚ it is consistent with‚ and linked to‚ the effects of signaling cascades and other diverse regulators of cell function. In particular‚ the time- and energy-critical process of cell proliferation utilizes components of a signal transduction cascade‚ i.e.‚ wingless/Wnt (wg/Wnt)‚ that first appeared in multicellular organisms several hundred million years after the dawn of SNF1-expressing eukaryotes. In supporting the complex process of growth regulation‚ wg/Wnt-like signaling integrates control of cell substrate utilization and energetics‚ biosynthetic and regulatory processes‚ cell-cell attachment‚ and gene expression. Second‚ arrest of cell proliferation‚ which may be essential if the cell is to survive a period of limited nutrient supply‚ is also closely coupled to metabolic and energetic factors. Thus‚ if cell energetics are impaired‚ e.g.‚ by nutrient deprivation‚ reciprocal changes in [ATP] and [AMP] will activate AMPK and lead to growth arrest. If the cell cycle were to progress despite so inadequate a metabolic foundation‚ the energetic deficiencies would be compounded through further depletion of the ATP that is already limited in supply. AMPK activation-induced mitochondrial fatty acid oxidation may foster additional adverse consequences‚ as noted. This sequence of events illustrates the point that arrest of the cell cycle under energetically adverse conditions would actually favor survival by limiting further depletion of ATP‚ mimimizing ROS generation‚ and promoting restoration of energy balance. Incompatibility of cell cycle progression with cell energetic insufficiency may represent the key to an hypothesized novel approach to cancer treatment‚ to be discussed.

6

Chapter 1

Third, while speed and timing in cellular energetics and signaling are of overriding importance in growth initiation and growth arrest, they are also critical in other seemingly unrelated processes, the energetics of which appear to have been conserved in their evolution from that more primitive requisite of survival. For example, neuronal activation in brain also depends almost entirely upon aerobic glycolysis, and thus requires suppression of fatty acid oxidation. Moreover, this process, which like mitosis is time- and energy-critical, also utilizes components of the wg/Wnt signaling cascade. Thus wg/Wnt-like signaling appears to be particularly important in those mechanisms, exemplified by cell cycle progression and neuronal activation, that link nutrient and energetics to processes in which speed and efficiency of response may be critical determinants of cell/organism survival. Activation of wg/Wnt-like signaling by lithium accounts for this small univalent cation’s remarkably diverse effects, e.g., on neuronal function, insulin signaling, and cell proliferation. Energetics, metabolism, and function of the neuron are also profoundly influenced through interaction with its neighbor, the astrocyte. In what may be a particularly important aspect of this interaction, fatty acid oxidation in astrocyte mitochondria drives production of ketone bodies (acetoacetate and which are exported and thus available for utilization by neurons. Despite the fact that ketone bodies are a less efficient fuel than glucose, and may compromise mitochondrial energetics, they are actually utilized by neurons in preference to glucose. Accordingly, it is hypothesized that excessive rates of astrocyte ketogenesis, driven by the elevated plasma concentrations of free (unesterified) fatty acids (FFA) that may occur — e.g., in obesity, insulin resistance, type II diabetes mellitus, or human immunodeficiency virus (HIV) infection — may suppress neuronal glycolysis, impairing energetic efficiency and promoting oxidative stress. Such conditions would predispose to synaptic dysfunction, and eventually to neuronal injury and neurodegenerative disorders such as Alzheimer disease and HIV dementia. Conversely, omega-3 fatty acids, although readily oxidized, increase inner mitochondrial membrane permeability to protons; this “uncoupling” of oxidative phosphorylation would contribute to fatty acid-induced suppression of both oxidative stress and cell proliferation. Notwithstanding these principles of neuronal substrate utilization, one region of the mammalian brain exhibits what appears to represent a recapitulation of the interaction between metabolism and feeding behavior that held sway in the primordial “sea”. Thus, as discussed below (Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior), those hypothalamic neurons that are specialized for neuroendocrine and metabolic regulation differ from their synaptic and cognitive counterparts in that they apparently oxidize fatty acids. Available evidence suggests the possibility


7

that feeding behavior‚ i.e.‚ appetite‚ is stimulated by oxidation of glucose in these cells‚ whereas their oxidation of fatty acids‚ possibly leptin-driven‚ may suppress appetite. Thus‚ determinants of the interaction between organism and environmental nutrient‚ although distanced from their primitive origins by more than a billion years and by the metabolic complexity inherent in a multicellular organism‚ appear to have survived in a specialized population of brain neurons‚ and in an unexpected and more sophisticated form: oxidize glucose and activate the drive to eat; oxidize fatty acids and suppress that drive. Finally‚ and despite its potentially adverse effects‚ it is clear that fatty acid oxidation is not only utilized widely to energize cell processes‚ but in fact is mandatory in certain situations. Unlike the dramatic and unpredictable events linked to wg/Wnt-like signaling‚ the processes fueled by fatty acids generally are characterized by sustained and relatively predictable energy demands. They include maintenance of posture or cardiac contraction (red muscle fibers)‚ ketogenesis (hepatocyte‚ astrocyte)‚ or gluconeogenesis (hepatocyte‚ renal tubular epithelial cell). Unlike glycolysis‚ there exists no anaerobic “option” to fatty acid oxidation; accordingly‚ it requires not only molecular oxygen but also — except for generally minor contributions of peroxisomes and the endoplasmic reticulum — mitochondria. As a corollary‚ cells in which such sustained and predictable energy requirements are met primarily by fatty acid oxidation (e.g.‚ red muscle‚ liver‚ and kidney) are abundantly endowed with mitochondria — i.e.‚ they are red. For each cell‚ this greater endowment of mitochondria would serve to distribute the oxidative burden of a given fatty acid load among a larger number of individual mitochondria‚ thereby minimizing overload of their respective electron transport chains and thus generation of the ROS that are the recognized byproducts of excessive fatty acid oxidation. In contrast‚ in those cells in which energy demands are sudden‚ intense‚ and unpredictable‚ energy generation is fueled principally by glycolysis (e.g.‚ white muscle‚ brain neurons)‚ while fatty acid oxidation is limited. Despite the fact that most brain neurons have fewer mitochondria (and lack their color)‚ neuronal energetic efficiency normally is maximized by complete mitochondrial oxidation of the products of glycolysis. In tissues other than brain‚ including red muscle‚ sudden demands for large amounts of energy may be met by the less efficient anaerobic glycolysis. In this pathway‚ glucose is catabolized only as far as pyruvate‚ which substitutes for oxygen as electron acceptor and is thereby reduced to lactate; it is exemplified by the oxygen debt and lactic acidemia incurred by human sprinters. A homely but useful illustration of these differences is provided by the contrasts in appearance and usage between the pectoral (“breast”)

8

Chapter 1

muscles of domesticated chickens and turkeys (few mitochondria‚ white: glycolytic; usually subserving a sporadic and unpredictable predator escape or defensive function) as compared with those of migratory ducks (many mitochondria‚ red: fatty acid oxidation; usually subserving a predictable and sustained long distance flight)‚ and their similar postural (leg‚ thigh) muscles (red in all three birds). Unfortunately‚ and despite these distinctions‚ it is possible in most cells for mitochondrial fatty acid oxidation to be driven at excessive rates by sustained increases in plasma [FFA] and lipoprotein levels‚ as often characterize obesity‚ insulin resistance‚ diabetes mellitus‚ or HIV infection. Moreover‚ “trans” fatty acids‚ being less readily oxidized in mitochondria‚ are more likely to undergo peroxisomal The increases in ROS generation and oxidative stress that result in either case may contribute importantly to the pathophysiology of diverse disorders‚ among them atherosclerosis‚ myocardial failure‚ reperfusion injury‚ aging‚ carcinogenesis‚ and Alzheimer disease. In view of these potentially damaging effects of ROS‚ it may seem surprising that ROS in low concentrations may also activate survival and proliferative responses. The significance of this apparent paradox will be addressed. A major objective of this volume is to critically examine those areas that are important in normal function and disease‚ and in which metabolic and energetic processes are integrated with signal transduction pathways. It is proposed that an understanding of such integrated systems‚ “hard wired” over millennia into mammalian physiology and cell biology‚ may provide novel perspective on many complex scientific and clinical phenomena that await elucidation and remain subjects of intensive investigation. It is further suggested that such integration brings to light unexpected parallels among a diversity of cells and cellular processes that seem to bear little resemblance to cell proliferation. For example‚ the linkage of glycolysis to unpredictable‚ sudden‚ and intense ATP demands of cell cycle progression and mitosis appears to have been adapted (with parallels in signal transduction as well) in support of other evolutionarily more recent processes — e.g.‚ neuronal activation and synaptic plasticity — in which the speed of initiation‚ rate‚ and intensity of the energetic (and thus synaptic) response are also critical to survival. The text consists of two major sections (Parts II and III)‚ preceded and followed by relatively brief introductory and concluding summations‚ i.e.‚ Parts I (this) and IV. In Part II‚ the nature of the linkage of cell metabolism and energetics to cell signaling and proliferation is explored in detail‚ as is their connection with programmed cell death‚ and with oxidative stress as an agent of cell injury and genomic instability in various settings ranging from atherosclerosis to carcinogenesis. Part III addresses brain neuronal function‚


9

substrate utilization‚ and the neuron-astrocyte interaction‚ along with their relationship to selected aspects of neurophysiology‚ regulation of feeding behavior‚ and the pathogenesis of HIV dementia and Alzheimer disease. Certain elements of the treatise are necessarily hypothetical. It is hoped that those concepts that may eventually be validated will provide an improved understanding of human biology and a basis for novel and productive approaches to the prevention and treatment of human disease.

PART II: FATTY ACIDS‚ MITOCHONDRIA‚ AND SIGNAL TRANSDUCTION: INTEGRATED CONTROL OF CELL PROLIFERATION‚ INJURY‚ AND DEATH

Chapter 2 Introduction to Part II

Selective regulation of the growth of specific cell populations is required for normal development and function in multicellular organisms. Dysregulation of such cell proliferation (“growth” and “proliferation” are used interchangeably) is fundamental to the pathogenesis of neoplastic and many non-neoplastic diseases. Despite recent advances‚ important gaps remain in the understanding of the mechanisms involved. In this context‚ it is particularly significant that growth signal transduction cascades and intermediary metabolism are intersecting‚ and that in certain settings control of cell growth appears to be predominantly nutritional or metabolic. Thus: 1) dietary glucose prevents‚ arrests‚ or reverses hepatic tumorigenesis in patients with glycogen storage disease type I (glucose-6-phosphatase deficiency)1‚2; 2) xenobiotic peroxisome proliferators‚ e.g.‚ fibrate hypolipidemic agents‚ inhibit mitochondrial fatty acid oxidation and in rodents initiate hepatocyte mitosis within 24 hours‚ leading to the development of liver cancer3-5; and 3) several inhibitors of cell growth increase mitochondrial fatty acid oxidation‚ absolutely and/or relative to mitochondrial ATP biosynthesis 6 (also‚ R. Ockner‚ unpublished observations). In the present treatise a model of cell growth regulation is developed that emphasizes the central importance of two requirements for initiation and successful completion of the cell cycle: 1) sufficiency of those substrates specifically required for mitosis-associated increases in biosynthetic‚ translocational‚ signaling‚ and regulatory processes; and 2) maximally efficient conversion of metabolic fuel into the increased quantities of chemical energy (largely ATP) upon which these processes depend. Pressures for survival imposed by these requirements fostered genetically programmed responses in primitive eukaryotic unicellular organisms more than a billion years ago — responses that are conserved to the present in 13

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higher plants and animals7-9. While it has been tacitly assumed that adequacy of such metabolic determinants is non-limiting in mammalian cell growth regulation‚ available evidence indicates that through their interaction with other factors they may in fact dominate control of growth‚ growth arrest‚ and apoptosis. The requirements of a valid general model of eukaryotic cell growth regulation are formidable. It must be applicable to organisms that span the phylogenetic tree from yeast to mammals‚ and include neoplastic disease. It must reconcile the dominant role of glucose as an inducer of growth in yeast7‚ with its seemingly paradoxical inhibition of growth early in mammalian liver regeneration10‚11 and in hepatic tumorigenesis associated with glycogen storage disease type I1‚2. It must contribute to an understanding of events in early embryogenesis12 and placental vascularization13‚ the pro-apoptotic anti-tumorigenic effects of dietary caloric restriction14-17‚ the dramatic proliferative and secretory responsiveness of the pancreatic islet beta cell to glucose and fatty acids18-23‚ and an unexpected parallelism between apoptosis and cell cycle progression24-32. Finally‚ it must functionally link growth-related factors that appear overtly “nutritional” (e.g.‚ glucose and fatty acids) with those that do not (e.g.‚ humoral and paracrine mediators‚ signal transduction cascades‚ and the nonsteroidal antiinflammatory drugs — NSAIDs). Energization of cell cycle progression requires substantial increases in ATP biosynthesis and utilization. With certain exceptions as noted‚ evidence discussed in the following pages supports the concept that this energization depends primarily upon glycolytic fueling of the citric acid cycle and aerobic metabolism in order to maximize mitochondrial inner membrane potential in turn drives the accelerated oxidative phosphorylation (ATP generation) that is needed for cell cycle progression and for support of the associated biosynthetic‚ translocational‚ signaling‚ and regulatory processes upon which it depends. Mitochondrial oxidation of fatty acids‚ conversely‚ inhibits glycolysis; it also may generate reactive oxygen species (ROS) and oxidative stress‚ thereby compromising mitochondrial energetics. Moreover‚ fatty acids may uncouple oxidative phosphorylation and are required for daughter cell membrane and eicosanoid biogenesis. As a corollary‚ it is a central element of the proposed model that suppression of mitochondrial fatty acid oxidation is obligatory in aerobic cell proliferation‚ and is implicit in anaerobic and hypoxic proliferation. Consistent with this concept‚ mitochondrial oxidation of fatty acids is inhibited by diverse mitogenic stimuli‚ and is relatively or absolutely increased by various agents of growth arrest and/or apoptosis. The proposed model of growth regulation is based on the concept that initiation and completion of the cell cycle require an appropriate metabolic

2. Introduction to Part II

15

milieu‚ absence of which is sufficient to arrest growth and/or precipitate apoptosis. In cell proliferation‚ requisite integration of intermediary metabolism‚ mitochondrial energetics‚ transcriptional regulation‚ and cell cycle progression is provided in significant measure by wingless/Wnt-like signaling‚ and its interactions with diverse determinants of survival and growth. In growth arrest and apoptosis‚ conversely‚ the AMP-activated protein kinase may play a key regulatory role. Heterogeneity among cells with respect to gene expression‚ energy generating capacity‚ and adaptation to microenvironment favors cell cycle completion and lineage survival for more “competent” genotypes and gene expression patterns; thus‚ Darwinian natural selection in microcosm. Incorporating these principles‚ the proposed model of growth regulation: 1) integrates effects of structurally diverse mitogens and antiproliferative agents on intermediary metabolism and signal transduction cascades; 2) rationalizes pro-survival and adverse effects of reactive oxygen species‚ contrasting utilization of nutrients by host and malignant tumor in cancer cachexia‚ and evolving changes in metabolism and gene expression during liver regeneration; 3) reconciles differing interpretations of the effects of nonsteroidal antiinflammatory drugs and cyclooxygenase-2; and 4) identifies the diverse mechanisms through which omega-3 fatty acids‚ salicylate and certain other nonsteroidal antiinflammatory drugs‚ and the “statin” hypolipidemic agents may exert similarly broad and beneficial antiproliferative and antioxidant effects. The model holds that metabolic milieu is a major determinant of cell survival and growth‚ and it may foster new approaches to their therapeutic control in both normal and transformed cells. It also suggests a broad role for mitochondrial and extramitochondrial fatty acid oxidation-induced oxidative stress in the pathogenesis of diverse forms of cell injury‚ as may occur‚ e.g.‚ in the endothelial lesion of atherosclerosis‚ nonalcoholic steatohepatitis‚ and ischemia-reperfusion injury‚ as well as in aging‚ cancer‚ and neurodegenerative disorders. While broadly oriented‚ the present analysis emphasizes the hepatocyte as this cell is an ideal paradigm of the interactions between intermediary metabolism and growth‚ and is central to the metabolic aberrations that characterize the syndrome of cancer cachexia. Although its life span is approximately 6 months33‚ it retains the capacity to re-enter the cell cycle within hours after partial hepatectomy‚ and to undergo one or more cell divisions within 24 to 72 hours‚ thereby restoring the organ to its appropriate size33‚34. Despite substantial progress‚ an understanding of the forces that drive this properly timed initiation and termination of mitosis has been elusive. As will be demonstrated‚ this process dramatically illustrates the fact that simple abundance of carbon and nitrogen sources and essential

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cofactors is not sufficient for growth; rather‚ substrate requirements and their mode of utilization are specific. Thus‚ DNA synthesis is delayed for approximately 12-16 hours following hepatectomy‚ until hepatocytes emerge from an initial phase in which mitochondrial fatty acid oxidation‚ gluconeogenesis‚ and oxidative stress are maximal‚ mitochondrial function is compromised‚ and growth is arrested. In the ensuing proliferative phase — and in most dividing cells — glycolysis and fatty acid synthesis predominate‚ while citric acid cycle activity‚ and energy coupling efficiency are high.

REFERENCES 1. P. Parker‚ I. Burr‚ A. Slonim‚ F. Ghishan‚ and H. Greene‚ Regression of hepatic adenomas in type Ia glycogen storage disease with dietary therapy‚ Gastroenterology 81:534-536 (1981). 2. R. Ockner‚ R. Kaikaus‚ and N. Bass‚ Fatty acid metabolism and the pathogenesis of hepatocellular carcinoma: review and hypothesis‚ Hepatology 18:669-676 (1993). 3. J. Reddy‚ M. Rao‚ D. Azarnoff‚ and S. Sell‚ Mitogenic and carcinogenic effect of hypolipidemic peroxisome proliferators [4-chloro-6(2‚3-xylidino)-2-pyrimidylthio]acetic acid (Wy-14‚643) in rat and mouse liver‚ Cancer Res 39:152-161 (1979). 4. P. Eacho‚ and P. Foxworthy‚ Inhibition of hepatic fatty acid oxidation by bezafibrate and bezafibroyl CoA.‚ Biochem Biophys Res Commun 157:1148-1153 (1988). 5. P. Foxworthy‚ and P. Eacho‚ Inhibition of hepatic fatty acid oxidation at carnitine palmitoyltransferase I by the peroxisome proliferator 2-hydroxy-3-propyl-4-[6-(tetrazol-5yl) hexyloxy]acetophenone.‚ Biochem J 252:409-414 (1988). 6. R. Ockner‚ N. Lysenko‚ N. Wu‚ and N. Bass‚ Hepatocyte growth inhibitors modulate mitochondrial and extramitochondrial fatty acid oxidation‚ Hepatology 24:253A (Abstract) (1996). 7. D. Hardie‚ D. Carling‚ and M. Carlson‚ The AMP-activated/SNF1 protein kinase subfamily: metabolic sensors of the eukaryotic cell?‚ Annu Rev Biochem 67:821-855 (1998). 8. R. Sheaff‚ M. Groudine‚ M. Gordon‚ J. Roberts‚ and B. Clurman‚ Cyclin E-CDK2 is a regulator of p27Kip1‚ Genes Devel 11:1464-1478 (1997). 9. M. Pagano‚ S. Tam‚ A. Theodoras‚ et al.‚ Role of the ubiquitin-proteasome pathway in regulating abundance of the cyclin-dependent kinase inhibitor p27‚ Science 269:682-685 (1995). 10. J. Simek‚ V. Chmelar‚ J. Melka‚ J. Pazderka‚ and Z. Charvat‚ Influence of protracted infusion of glucose and insulin on the composition and regeneration activity of liver after partial hepatectomy in rats‚ Nature 213:910-911 (1967). 11. J. Ngala Kenda‚ B. De Hemptinne‚ and L. Lambotte‚ Role of metabolic overload in the initiation of DNA synthesis following partial hepatectomy in the rat‚ Eur Surg Res 16:294302(1984). 12. J. Van Blerkom‚ P. Davis‚ V. Mathwig‚ and S. Alexander‚ Domains of high-polarized and low-polarized mitochondria may occur in mouse and human oocytes and early embryos‚ Hum Reprod 17:393-406 (2002). 13. O. Genbacev‚ Y. Zhou‚ J. Ludlow‚ and S. Fisher‚ Regulation of human placental development by oxygen tension.‚ Science 277:1669-1672 (1997).

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14. B. Grasl-Kraupp‚ W. Bursch‚ B. Ruttkay-Nedecky‚ A. Wagner‚ B. Lauer‚ and R. SchulteHermann‚ Food restriction eliminates preneoplastic cells through apoptosis and antagonizes carcinogenesis in rat liver‚ Proc Natl Acad Sci USA 91:9995-9999 (1994). 15. K. Kolaja‚ K. Bunting‚ and J. Klaunig‚ Inhibition of tumor promotion and hepatocellular growth by dietary restriction in mice‚ Carcinogenesis 17:1657-1664 (1996). 16. R. Gredilla‚ G. Barja‚ and M. Lopez-Torres‚ Effect of short-term caloric restriction on production and oxidative DNA damage in rat liver mitochondria and location of the free radical source‚ J Bioenerg Biomembr 33:279-287 (2001). 17. V. D. Longo‚ and C. E. Finch‚ Evolutionary medicine: from dwarf model systems to healthy centenarians?‚ Science 299:1342-6 (2003). 18. S. Bonner-Weir‚ D. Deery‚ J. Leahy‚ and G. Wei‚ r‚ Compensatory growth of pancreatic beta-cells in adult rats after short-term glucose infusion.‚ Diabetes 38:49-53 (1989). 19. Y. Zhou‚ D. Priestman‚ P. Randle‚ and V. Grill‚ Fasting and decreased B cell sensitivity: important role for fatty acid-induced inhibition of PDH activity‚ Am J Physiol 270:E988994 (1996). 20. B. Leibiger‚ T. Moede‚ T. Schwarz‚ et al.‚ Short-term regulation of insulin gene transcription by glucose‚ Proc Natl Acad Sci USA 95:9307-9312 (1998). 21. C. Bernard‚ M. Berthault‚ C. Saulnier‚ and A. Ktorza‚ Neogenesis vs. apoptosis as main components of pancreatic beta cell mass changes in glucose-infused normal and mildly diabetic adult rats.‚ FASEB J 13:1195-1205 (1999). 22. G. Steil‚ N. Trivedi‚ J.-C. Jonas‚ et al.‚ Adaptation of beta-cell mass to substrate oversupply: enhanced function with normal gene expression.‚ AJP - Endocrinol Metab 280:E788-E796 (2002). 23. R. Tuttle‚ N. Gill‚ W. Pugh‚ et al.‚ Regulation of pancreatic beta-cell growth and survival by the serine/threonine protein kinase Akt1/PKBalpha‚ Nat Med 7:1133-1137 (2001). 24. W. Meikrantz‚ and R. Schlegel‚ Apoptosis and the cell cycle‚ J Cell Biochem 58:160-174 (1995). 25. G. Fan‚ X. Ma‚ B. Kren‚ and C. Steer‚ The retinoblastoma gene product inhibits TGFbetal induced apoptosis in primary rat hepatocytes and human HuH-7 hepatoma cells.‚ Oncogene 12:1909-1919 (1996). 26. G. Gil-Gómez‚ A. Berns‚ and H. Brady‚ A link between cell cycle and cell death: Bax and Bcl-2 modulate Cdk2 activation during thymocyte apoptosis‚ EMBO J 17:7209-7218 (1998). 27. J. Park‚ K. Kim‚ S. Kim‚ and S. Lee‚ Caspase 3 specifically cleaves p21WAF1/CIP1 in the earlier stage of apoptosis in SK-HEP-1 human hepatoma cells‚ Eur J Biochem 257:242248 (1998). 28. E. Thompson‚ The many roles of c-Myc in apoptosis‚ Annu Rev Physiol 60:575-600 (1998). 29. K. Hiromura‚ J. Pippin‚ M. Fero‚ J. Roberts‚ and S. Shankland‚ Modulation of apoptosis by the cyclin-dependent kinase inhibitor p27(Kip1)‚ J Clin Invest 103:597-604 (1999). 30. E. Lind‚ J. Wayne‚ Q.-Z. Wang‚ T. Staeva‚ A. Stolzer‚ and H. Petrie‚ Bcl-2-induced changes in E2F regulatory complexes reveal the potential for integrated cell cycle and cell death functions‚ J Immunol 162:5374-5379 (1999). 31. K. Hoeflich‚ J. Luo‚ E. Rubie‚ M. Tsao‚ O. Jin‚ and J. Woodgett‚ Requirement for glycogen synthase kinase-3beta in cell survival and NF-kappaB activation‚ Nature 406:8690 (2000). 32. R. F. Schwabe‚ and D. A. Brenner‚ Role of glycogen synthase kinase-3 in TNF-alphainduced NF-kappaB activation and apoptosis in hepatocytes‚ Am J Physiol Gastrointest Liver Physiol 283:G204-11 (2002).

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33. N. Fausto. Hepatic regeneration‚ in: Hepatology: a Textbook of Liver Disease‚ T. Boyer‚ ed.‚ Saunders‚ Philadelphia (1996)‚ pp. 32-58. 34. W. Kaufmann‚ and T. Goldsworthy‚ Regulation of the hepatocyte cell cycle‚ Prog Liver Dis 15:31-56 (1997).

Chapter 3 Nutrient and Energy Metabolism in Cell Proliferation

3.1.

INTRODUCTION

In this chapter the fundamentals of intermediary metabolism and energetics are discussed as they relate to the cell cycle and cell proliferation. Emphasis is placed on the important role of glucose in ATP generation under both aerobic and hypoxic conditions‚ and on the critical integration of the cell cycle with gene expression‚ intermediary metabolism‚ and cell-cell interaction that is provided by wingless/Wnt-like signaling. This signal cascade may be activated by growth factor receptors as well as by low-level reactive oxygen species (ROS)‚ and by the small univalent cation thus accounting for the latter’s versatility in diverse processes and conditions. Implicit in the critical role of glycolysis (aerobic or hypoxic) in energizing the cell cycle is the requirement that fatty acids be excluded as oxidized mitochondrial substrates. The complex and essential modulatory role of fatty acids in this and other settings is dealt with in subsequent chapters.

3.2.

INTERMEDIARY METABOLISM: GENERAL CONSIDERATIONS

The following two paragraphs are provided to help orient the reader to those aspects of intermediary metabolism‚ and the utilization of the predominant substrates — glucose and fatty acids — that are relevant to cell proliferation. The accompanying figures illustrate several aspects of these processes‚ and will be referred to in various portions of the text as appropriate. 19

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Chapter 3

Figure 3.1. Glycolysis- and citric acid cycle-fueled biosynthesis of ATP and fatty acid; suppressed fatty acid oxidation and gluconeogenesis. This figure represents the predominant pattern of substrate utilization in proliferating cells; enclosed areas represent mitochondria. Arrow thickness approximates relative pathway activity; dotted lines: pathways that are suppressed during cell proliferation. Fatty acids also may be derived from plasma. Abbreviations — CPT-I: carnitine palmitoyltransferase-I (shown as inhibited by malonyl CoA); EMFO: extramitochondrial fatty acid oxidation in peroxisomes‚ and in endoplasmic reticulum); F26BP: fructose-2‚6-bisphosphate; PEPCK: phosphoenolpyruvate carboxykinase; PL: glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

Glucose utilization requires phosphorylation by hexokinase and entry into the glycolytic pathway (Fig. 3.1). Completion of glycolysis with formation of pyruvate generates a net of 2 molecules of ATP per molecule of glucose. Under certain conditions the process ends here: pyruvate substitutes for oxygen as acceptor of the electrons generated by glycolysis‚ thereby forming lactate which is exported from the cell. This “anaerobic glycolysis” permits ATP generation in the absence of oxygen‚ but is far less efficient than mitochondrial utilization of glycolysis-derived pyruvate via the key regulatory enzyme pyruvate dehydrogenase and the citric acid cycle

Figure 3.2. Mitochondrial electron transport chain and oxidative phosphorylation. Pathways are shown for utilization of three principal substrates, i.e., glucose, fatty acid (FA), and glutamine. Pyruvate dehydrogenase- (PDH)-mediated entry of acetyl CoA into citric acid cycle is subject to feedback regulation by acetyl CoA/CoA, NADH/NAD, and ATP/ADP, thus minimizing electron transport chain overload and oxygen radical formation. Compared to PDH regulation, carnitine palmitoyltransferase-I- (CPT-I)-mediated entry of FA into the cycle is less tightly controlled and may be overridden by high fatty acid concentrations; as a result, acetyl CoA and citric acid cycle activity may cause inhibition of PDH, relative overloading of the electron transport chain, and increased propensity to oxygen radical formation. Conversion of glutamine and glutamate to 2-oxo-glutarate bypasses the pyruvate dehydrogenase control point and directly fuels the citric acid cycle.

3. Nutrient and Energy Metabolism in Cell Proliferation

21

(Fig. 3.1‚ and Fig. 3.2 facing this page). In this latter pathway‚ mitochondrial inner membrane potential is maintained by electron flux-energized export of protons at electron transport chain complexes I‚ III‚ and IV; in turn‚ drives ATP synthase-mediated oxidative phosphorylation. Complete oxidation of a glucose molecule through this process yields 30 molecules of ATP. Fatty acids may also undergo oxidation in mitochondria to generate ATP (Fig. 3.2 facing this page‚ and Fig. 3.3). The long chain fatty acid molecule is converted to its acyl CoA then acylcarnitine derivatives. The latter step‚ mediated by carnitine

Figure 3.3. Fatty acid oxidation and gluconeogenesis; suppressed glycolysis and fatty acid synthesis. Enclosed areas represent mitochondria. This pattern of substrate utilization is antiproliferative. Arrow thickness approximates relative pathway activity; dotted line: pathway suppressed during growth arrest. Numbers in parentheses indicate approximate yields of ATP from complete oxidation of palmitate (~100)‚ compared (in a ketogenic cell‚ e.g.‚ hepatocyte) with complete diversion to ketogenesis (~20). Fatty acids activate transcriptionally upregulating both mitochondrial and extramitochondrial fatty acid oxidation. Gluconeogenesis limited to hepatocyte‚ renal tubular epithelial cell‚ and astrocyte. When present in excess‚ fatty acids may override malonyl CoA inhibition of CPT-I. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; EMFO: extramitochondrial fatty acid oxidation in peroxisomes‚ and oxidation in endoplasmic reticulum); F26BP: fructose-2‚6-bisphosphate; PEPCK: phosphoenolpyruvate carboxykinase; PL: glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

Chapter 3

22

palmitoyltransferase-I (CPT-I)‚ permits fatty acid entry into the mitochondrial matrix where acyl CoA is regenerated and undergoes providing electrons to the electron transport chain and generating acetyl CoA. Acetyl CoA may enter the citric acid cycle or be converted to ketone bodies and exported from the cell. Importantly‚ accelerated fatty acid oxidation may drive excessive electron flux‚ predisposing to formation of reactive oxygen species (ROS) and oxidative stress. A theoretical yield of 106 ATP molecules per palmitic acid molecule is diminished by ketogenesis‚ fatty acid uncoupling of oxidative phosphorylation‚ loss of electrons from the electron transport chain in generating ROS‚ and the injurious effects of oxidative stress on components of the electron transport chain. An important aspect of these alternative pathways (glycolysis and mitochondrial fatty acid oxidation) is their relative exclusivity. Thus‚ aerobic glycolysis and the pentose phosphate pathway (Fig. 3.1) generate substrates‚ e.g.‚ NADPH‚ that are utilized in antioxidant defense and in the de novo synthesis of fatty acids‚ isoprenoids‚ and sterols. The first committed intermediate in fatty acid biosynthesis‚ malonyl CoA‚ inhibits CPT-I and mitochondrial fatty acid oxidation. Glycolysis thereby supports fatty acid synthesis; its indirect (malonyl CoA-mediated) inhibition of CPT-I and fatty acid oxidation‚ however‚ may be overridden in states of cellular fatty acid excess. Mitochondrial fatty acid oxidation‚ moreover‚ strongly inhibits glucose uptake‚ glycolysis‚ and pyruvate dehydrogenase1-4. In hepatocytes and renal tubular epithelial cells‚ mitochondrial fatty acid oxidation fuels gluconeogenesis (Fig. 3.3) and export of newly synthesized glucose. Accordingly‚ ATP generation may be: 1) primarily glycolytic (Fig. 3.1) with suppression of fatty acid oxidation; or 2) primarily dependent on fatty acid oxidation (Fig. 3.3)‚ supporting gluconeogenesis and/or ketogenesis‚ while suppressing glycolysis and predisposing to ROS generation and oxidative stress. These concepts will be revisited as cellular processes and conditions are examined in relation to growth regulation.

3.3.

ATP GENERATION‚ CARBOHYDRATE METABOLISM‚ AND CELL GROWTH

Evidence that rates of oxygen consumption and ATP generation are increased in mitosis is provided by studies of several transformed and nontransformed cell systems. Rates increase throughout the cell cycle‚ reaching high levels during G1 and S and even higher in G2/M5-9. ATP production was linearly related to growth of hybridoma cells (clonal cell lines created by fusion of antibody-producing and tumor cells)10‚ to epidermal growth factor (EGF)- and/or insulin-stimulated DNA synthesis in 3T3 fibroblast


23

cultures11‚ and to DNA repair in yeast12. In primary rat hepatocyte cultures‚ citric acid cycle intermediates (oxidation of which fuels the electron transport chain‚ driving oxidative phosphorylation) stimulated DNA synthesis and mitosis‚ equalling or exceding the mitogenic effect of EGF13. Conversely‚ depletion of nucleotide triphosphates (NTP)‚ which include the readily interconvertible ATP‚ CTP‚ GTP‚ TTP‚ and UTP‚ caused Gl arrest in the absence of DNA damage14 and in E. coli inhibited initiation of ribosomal RNA transcription15. Importantly‚ availability of ATP and oxygen at critical sites may be limited by cytoplasmic diffusion barriers16-18.

3.3.1. Aerobic Glycolysis‚ Mitochondrial Respiration‚ and Oxidative Phosphorylation Increased glycolysis‚ long recognized as characteristic of proliferating cells19-23‚ is associated with altered expression of related genes. Thus‚ in neoplastic hepatocytes the normally expressed low affinity adult plasma membrane glucose transporter‚ GLUT 2‚ is supplanted by the high affinity GLUT 1 fetal isoform24 that is expressed in many human cancers25 (Fig. 3.1). Glucose uptake and plasma membrane glucose transporter expression also increased in ras or src transfected26 or virally transformed27 rat fibroblasts‚ and hepatic glucose utilization was increased by overexpression of c-myc in Conversely‚ inhibition of streptozotocin-treated transgenic mice28. insulinoma cell growth by butyrate significantly decreased GLUT 1 expression29. After uptake‚ glucose normally is phosphorylated in hepatocytes by the low affinity cytosolic adult type IV hexokinase‚ designated glucokinase. In contrast‚ tumor cells may express a mitochondrial-bound higher affinity hexokinase23‚30‚ predominantly type II31‚ reflecting at least in part epigenetic upregulation of the HKII gene through promoter demethylation32. Mitochondrial-bound hexokinase is an integral component of a membranespanning complex that appears to be interconvertible with the apoptosisassociated permeability transition pore (Fig. 3.4; also‚ see Chapter 5.3.: Mitochondrial Function in Apoptosis). Located at the contact sites between inner and outer mitochondrial membranes‚ this complex (voltage-dependent anion channel‚ or VDAC) also includes the adenine nucleotide translocase‚ porin‚ creatine kinase‚ as well as anti-apoptotic members of the Bcl-2 protein family33-35. VDAC itself is associated with the outer mitochondrial membrane‚ but the mechanism by which this highly permeable membrane generates the potential (“voltage”) presumably required to regulate the voltage-dependent channel (VDAC) is unknown36. Accordingly‚ the schematic shown in Fig. 3.4‚ while consistent with a proposed model that in effect provides for direct‚ regulated communication across the profound

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Chapter 3

potential difference between cytosol and the mitochondrial matrix36‚ is not established. In any case‚ this location accounts for hexokinase-mediated suppression of Bax-induced cytochrome c release37‚38 (also‚ see Chapter 5.3.: Mitochondrial Function in Apoptosis) and kinetically favors both hexokinase utilization of mitochondrially generated ATP23‚39 and the return of ADP so produced to the ATP synthase. Transfection of this hexokinase isoform into 3T3 fibroblasts increased both overall hexokinase activity and cell growth40. Several human and rat tumors and tumor cell lines express a hexokinase variant which was specifically inhibited by mannoheptulose but not by glucose-6-phosphate‚ the physiological inhibitor of the normally expressed hexokinase; mannoheptulose decreased both glucose uptake and cell growth41. Expression of type II hexokinase is amplified in AS30D hepatoma and other cancer cell lines42‚ reflecting increased expression of structurally normal VDAC43‚ and contributing to an aerobic glycolytic phenotype correlated with cell growth42.

Figure 3.4. Mitochondrial voltage-dependent anion channel (VDAC) components and relationship to electron transport chain; schematic representation. VDAC complex-mediated juxtaposition of hexokinase II and the adenine nucleotide translocase in proliferating cells optimizes energetics of glucose phosphorylation to glucose-6-phosphate and entry into the glycolytic pathway‚ leading to pyruvate formation. Detailed structural relationships remain incompletely defined (see text). Increased mitochondrial fatty acid oxidation inhibits glucose uptake‚ hexokinase‚ and pyruvate dehydrogenase‚ and predisposes to electron “leakage” from the electron transport chain; the latter results in one-electron reduction of oxygen and formation of and derivative ROS. Excess cellular fatty acids may disrupt hexokinase II‚ VDAC‚ and the contact sites. Unlike anti-apoptotic Bcl-2 family members‚ pro-apoptotic members may associate with‚ and perturb‚ the mitochondrial membrane after activation of the apoptotic cascade. Abbreviations — CYT C: cytochrome c; inner mitochondrial membrane potential; superoxide anion; UBIQ: ubiquinone; ROS: reactive oxygen 44 species; VDAC: voltage-dependent anion channel. With permission .


25

Further modulation of glycolysis is effected by 6-phosphofructokinase2/fructose-2‚6-bisphosphatase (PFK2/F-2‚6-BPase) and the activity of its PFK2 component (Fig. 3.1). The latter is increased in proliferating cells‚ resulting in accumulation of the key regulatory intermediate fructose-2‚6bisphosphate (F-2‚6-BP)45. F-2‚6-BP‚ an allosteric activator of phosphofructokinase and inhibitor of fructose- 1‚6-bisphosphatase‚ augments glycolysis and inhibits the opposing gluconeogenesis pathway. PFK2 activity and expression — and glycolysis — are increased by EGF‚ hepatocyte growth factor (HGF)‚ and insulin in primary hepatocyte cultures and several cell lines‚ including HT-29 human colon carcinoma‚ chick embryo and human fibroblasts‚ and A431 hepatoma45’46. Conversely‚ the growth inhibitor transforming growth factor decreases hepatocyte PFK2 activity and F-2‚6-BP abundance46‚ and increases the mRNA abundance of phosphoenolpyruvate carboxykinase (PEPCK)47‚ a key determinant of gluconeogenesis (see Chapter 6.3.: Enhanced glycolysis during tumorigenesis is also favored by the mitogens insulin 48 and EGF49‚ by increased expression and activity of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and pyruvate kinase (PK)24‚ and by decreased PEPCK expression24 (Fig. 3.1). Increased GAPDH expression was correlated with the transformed state and glucose transporter expression in several cell lines50. In EGF receptor-expressing human MDA468 breast cancer cells‚ EGF caused both a near doubling of the rate of glucose uptake and phosphorylation and an increase in cell growth51. Limited production of lactate in the latter experiments suggests that glucosederived pyruvate entered the citric acid cycle‚ consistent with the importance of oxidative phosphorylation during cell growth. This critical step depends on the conversion of phosphoenolpyruvate (PEP) to pyruvate by PK‚ the isoform of which is increased in virtually all tumors52. The insulin-like response which these changes represent is consistent with the metabolic changes early in experimental and human hepatocarcinogenesis53‚ in which expression of insulin receptor substrate-1 (IRS-1) is increased54. In addition to glycolysis‚ glucose-6-phosphate may enter the pentose phosphate pathway via the key enzyme glucose-6-phosphate dehydrogenase (Fig. 3.1). This pathway generates NADPH‚ required both for biosynthesis of fatty acids and sterols‚ and for maintenance of cell redox balance55. The implied dependence on oxidative phosphorylation (Fig. 3.2 facing page 21) during cell cycle progression56 is demonstrated more directly by parallel increases in oxygen consumption‚ ATP biosynthesis‚ and ATP abundance5-8‚57. In human leukemia cells‚ cell cycle progression required augmented biogenesis of mitochondria and of mitochondrial proteins involved in oxidative phosphorylation. Inhibition of the latter suppressed mitochondrial ATP production and cell cycle progression‚ and was not

26

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compensated for by increased generation of ATP by glycolysis alone9. In hybridoma cells‚ ATP generation depended chiefly on glucose consumption and citric acid cycle activity‚ the latter indicative of aerobic metabolism10. Four types of transplanted tumors in mice utilized glucose‚ as well as other mitochondrial substrates including ketone bodies‚ glutamine‚ and lactate58. Conversely‚ inhibition of aerobic glycolysis appears to account for the dramatic therapeutic effect of STI571 against chronic myelogenous leukemia59. The importance of mitochondrial ATP generation in tumor cells is further suggested by the aforementioned mitochondria-associated hexokinase (Fig. 3.4) and its kinetically preferential access to newly synthesized ATP‚ permitting accelerated glucose-6-phosphate formation23‚30‚39. Utilization of glucose and/or oxygen was correlated with growth of xenografts of human breast and lung cancers and tumor-derived cell lines in nude mice21‚ and in adriamycin-resistant vs. adriamycinsensitive wild type Ehrlich ascites tumor cells60. In the presence of glucose‚ approximately 50% of the ATP requirement of most rapidly growing tumor cells was derived from glycolysis20; when glucose was replaced as a carbon source by glutamine (a precursor of the citric acid cycle intermediate 2-oxoglutarate)‚ ATP was produced entirely via oxidative phosphorylation 61 . Significantly‚ glutamine in this circumstance‚ and in L929 mouse fibrosarcoma cells62‚ was able to sustain ATP generation at rates equal to those with glucose alone. This further indicates the central importance of the citric acid cycle and oxidative phosphorylation in cell proliferation‚ and may account for the dominant role of glutamine as a nutrient in several types of normal and neoplastic cells during rapid growth61-64‚ including intestinal epithelium as discussed below (see Chapter 6.2.: Butyrate). Finally‚ the importance of mitochondrial energization and oxidative phosphorylation is emphasized by the increase in that is characteristic of proliferating cells‚ both normal and transformed65‚ and the predisposition by hypoxia to p53-dependent and p53-independent apoptosis. Despite these considerations‚ it is important to note that in selected circumstances cell proliferation may be supported under hypoxic conditions by anaerobic glycolysis. This occurs‚ e.g.‚ in tumor angiogenesis66‚67‚ placental vascularization 68 ‚ and mitogen-stimulated t h y m o c y t e proliferation69. Moreover‚ survival and growth of cells in less well perfused and oxygenated regions of a tumor mass is critically linked to their response to hypoxia‚ largely dependent on activation of hypoxia-inducible factor Active transcriptionally upregulates genes the products of which promote erythropoiesis‚ angiogenesis‚ and glycolysis67‚70‚ including mitochondrial VDAC-associated hexokinase II23’31 (also‚ see 3.3.2.3.: Reactive Oxygen Species). The latter’s perhaps unexpected expression – under conditions in which limited oxygen availability


27

constrains mitochondrial respiration — may provide for an anticipated and accelerated glycolytic response to increases in oxygen availability mediated by erythropoietic and angiogenic factors. may also induce expression of the cell cycle inhibitors and leading 71 to G1 cell cycle activation and/or G1/S arrest (also‚ see 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). In addition‚ through its activation of wg/Wnt-like signaling (see below)‚ erythropoietin promotes endothelial cell survival72. Furthermore‚ hypoxia may serve as a growth selection factor‚ favoring survival and proliferation of tumor cells in which p53 activity is deleted and p53-mediated apoptosis is therefore defective‚ e.g.‚ as the result of p53 mutation73. Under normoxic conditions‚ mitochondrial respiration may be suppressed by glucose in normal and cancer cells through the incompletely understood “Crabtree effect”36‚74-76 (also‚ see Chapter 7.3.: Origins of Intramitochondrial Oxidative Stress).

3.3.2. Wingless/Wilt-Like Signaling: Convergence of Antecedents and the Unpredictable A major regulatory linkage between mitosis and glycolysis is provided by wingless/Wnt-like (wg/Wnt-like) signaling pathways and their pivotal 77-80 negative regulator glycogen synthase . These cascades are of critical importance in development as well as in the pathogenesis of human familial adenomatous polyposis (FAP) and‚ untreated‚ the latter’s virtually invariable culmination in colon cancer. They are commonly dysregulated (constitutively activated) in human cancer‚ often the result of inactivating mutations in the gene encoding the inhibitory lipid phosphatase PTEN81. A critical component of wg/Wnt-like signaling is phosphatidyl inositol-3-kinase- (PI3K)-dependent activation of the protein kinase Akt‚ also designated protein kinase B (Akt/PKB)82‚83. In this sequence‚ active PI3K generates phosphoinositide activators of phosphoinositide-dependent kinase 1 (PDK1)‚ which in turn activates Akt/PKB. Activated Akt/PKB promotes both cell survival and proliferation (Fig. 3.5)‚ inducing the secretion of insulin by the pancreatic islet beta cell84‚ and of insulin-like growth factors by 3T3-L1 preadipocytes85. The survival-enhancing function of Akt/PKB reflects in part its enhancement of glucose utilization‚ activation of mitochondrial hexokinase II and glycolysis generally‚ hexokinase II-mediated suppression of Baxinduced cytochrome c release37‚38‚ and maintenance of high and an open conformation of VDAC80‚86‚87. It is actively expressed in embryonic stem cells88‚ and essential for proliferative self-renewal in hematopoietic stem cells89. Moreover‚ Akt/PKB phosphorylates and thereby inactivates:

28

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Figure 3.5. Wingless/Wnt-like signaling: cell growth and suppression of cell death. Activation of PI3K-Akt/PKB via diverse receptor and non-receptor tyrosine kinases (including EGF, HGF, insulin, IGF-1, PDGF, and receptors) and/or ROS leads to Akt/PKB phosphorylation, activation of SREBPs, MDM2, and telomerase, and inactivation of the apoptosis-promoting and/or growth inhibiting proteins Bad, caspase 9, Forkhead transcription factors, p53, and (not shown) Inactivation of abrogates its inhibition of proteins important in storage and utilization of glucose (glycogen synthase, pyruvate dehydrogenase), cell cycle progression (cyclin D1), and gene transcription The resulting increase in c-myc expression leads, among its other effects, to transcriptional activation of the gene encoding the cyclin-dependent kinase cdk4. Abbreviations — COX 2: cyclo-oxygenase 2 (prostaglandin G/H synthase 2); EGF: epidermal growth factor; HCC: hepatocellular carcinoma; HGF: hepatocyte growth factor; IGF-1: insulin-like growth factor-1; PDGF: platelet-derived growth factor; PDH: pyruvate dehydrogenase; SREBPs: sterol regulatory element binding proteins; tumor necrosis

1) Bad‚ a pro-apoptotic member of the Bcl-2 family90; 2) caspase 9‚ a key cysteine protease in the apoptotic cascade91; and 3) the cytosolic 92 inhibitor of the transcriptional regulator nuclear factor . Inactivation of Bad and caspase 9‚ and activation of promote cell survival by suppressing specific mitochondrial and extramitochondrial events in apoptosis (see Fig. 3.5‚ and Chapter 5.3.: Mitochondrial Function in Apoptosis). Akt/PKB also suppresses Fas-mediated apoptosis by inhibiting interaction of procaspase 8 with the death-inducing signaling

3.Nutrient and Energy Metabolism in Cell Proliferation

29

complex (DISC)93. As a corollary of its linkage to aerobic metabolism‚ however‚ Akt/PKB is unable to protect against oxygen deprivation80‚94. Support of cell proliferation by Akt/PKB‚ in contrast to its survivalenhancing effects‚ reflects to a large extent its inactivation of several antiproliferative factors (Fig. 3.5). These include the Forkhead family of transcription factors95‚ the G2 arrest-inducing Chk1 kinase96‚ and Forkhead transcription factors (AFX‚ FKHR‚ FKHRL-1‚ and FOXO-3 in mammals; Daf 16 in C. elegans) induce G1 arrest97‚ 98 Gadd45-mediated G2 arrest and DNA repair ‚ antioxidant defenses including (mitochondrial) Mn-superoxide dismutase99‚ and apoptosis-inducing ligand (TRAIL)100. Inactivation of is a major targeted effect of Akt/PKB. Active suppresses mitosis by phosphorylating key elements in cell proliferation‚ leading to their inactivation (glycogen synthase‚ PDH) or degradation and cyclin D1)101-104 (Fig. 3.5); as a result‚ Ecadherin-mediated cell-cell adhesion is stabilized‚ fostering cell differentiation101‚102. Thus‚ inactivation of contributes to mitosis by providing for several requirements of this process: 1) inactivation prevents phosphorylation of thereby increasing the cytosolic abundance of unphosphorylated and favoring its participation as a transcriptional regulator in conjunction with Tcf/lef101‚102. Among the targets of transactivation are 105 the genes encoding cyclin D1 ‚ the matrix metalloproteinase matrilysin106‚ and c-myc‚ an inducer of cell cycle progression107‚108. C-myc‚ in turn‚ transcriptionally activates expression of the cyclin D-dependent kinase cdk4109‚ and a member of the peroxiredoxin family important in maintenance of mitochondrial function110‚ while suppressing that of the cdk inhibitor 2) inactivation prevents its phosphorylation and resulting degradation of cyclin D1104‚ thereby increasing the latter’s abundance‚ its activation of cyclin-dependent kinases cdk4 and cdk6‚ and cell cycle progression. The cell cycle is also subject to control by the interaction between and Akt/PKB in regulating the Cip/Kip family of cdk inhibitors and Although these cell cycle regulators serve to control the G1/S transition‚ both are sequestered by‚ and at low abundance activate‚ cyclins D1 and D2112-114. The importance of and to cell survival (see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer) is suggested by the rarity with which they undergo inactivating mutations in human cancer113‚ as well as by the complexity of their regulation. For example‚ phosphorylation of at T57 decreases its stability115. This action is opposed by Akt/PKB‚ which increases stability by inactivating

Chapter 3

30 115

, and by directly phosphorylating at S146116. In addition, Akt/PKB-mediated phosphorylation of at T145 decreases binding of to PCNA116 and/or promotes its cytosolic localization117; both of the latter actions would tend to diminish the antiproliferative effects of thus promoting cell cycle progression. Similarly, Akt/PKB-mediated phosphorylation of promotes the latter’s cytosolic retention118,119. Thus, Akt/PKB influences cell cycle regulation in part through its effects on nuclear abundance of both of the major Cip/Kip proteins, which in low abundance promote activation of the cyclin D kinases cdk4 and cdk 6, and in high abundance inhibit cyclin E/cdk 2 and the G1/S transition. 3) inactivation, in addition to augmenting cell cycle progression, abrogates its inactivating phosphorylation of glycogen synthase and pyruvate dehydrogenase103 (also, see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). Further, in addition to activating mitochondrial hexokinase II80,86, Akt/PKB directly upregulates GLUT 4 translocation and glucose uptake120, and activates PFK2121. Together, these direct and indirect effects of Akt/PKB would support increased cellular uptake of glucose, its storage (as glycogen), and its utilization in aerobic or anaeobic (hypoxic) glycolysis. Suppression of c-myc expression in rat liver by fasting122, which inhibits glycogen storage and glycolysis, further supports the linkage between wg/Wnt-like induction of mitosis and glucose metabolism. In addition, Akt/PKB’s inactivation of abrogates the latter’s activation of MEKK1 and the stress-activated (JNK/SAPK) protein kinase pathway123. Thus, predisposition to colon cancer in the FAP syndrome can be understood as the result of constitutively augmented wg/Wnt-like signaling, reflecting mutations in the adenomatous polyposis coli (APC) or gene. Such mutations prevent formation of the complex upon which the phosphorylation and degradation of and abrogation of wg/Wnt-like signaling, depend. As a corollary, increased hepatocyte death and embryonic lethality caused by 124 absence of may reflect failure of a protective 125 cell cycle arrest under conditions inappropriate for mitosis (see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer). In the normal postmitotic differentiating colonocyte, conversely, absence of wg/Wnt-like signaling permits to remain unphosphorylated and enzymatically active. The mutually inhibitory interactions between IRS-1 and and the latter’s ubiquitous expression126-128, suggest that insulin and other mitogenic factors serve as activators of a wg/Wnt-like pathway (Fig. 3.5)128, as has been demonstrated, e.g., for insulin-like growth factor-1 (IGF-1)129, EOF130,


31

92 HGF131‚132‚ ‚ the B cell CD40 receptor133‚ and the recently characterized Wnt itself134. Direct evidence for such an interaction is also provided by the observation that insulin stimulates the integrin-linked kinase (ILK) in a PI3K-dependent manner135‚ leading to activation of Akt/PKB‚ inactivation of and increased nuclear translocation of Although wg/Wnt-like signaling-dependent phosphorylation of was previously regarded as protein kinase C- (PKC)-mediated and therefore independent of PI3K and Akt/PKB136‚ its functional equivalent can in fact be initiated by this more recently identified mechanism‚ and by others as well. Moreover‚ it is now clear that several classes of PKC are activated in a PI3K-dependent manner137‚ and that these two pathways may interact in the induction of colonocyte proliferation and colon carcinogenesis138. Inhibition of by insulin during glycogen synthesis has long been recognized‚ and is required early in human and experimental hepatocarcinogenesis as noted above53‚54. On the basis of these recent observations‚ however‚ it now is clear that the interaction of growth factors such as HGF and insulin with wg/Wnt-like signaling activates the critically important convergence of four distinct and essential antecedents to cell proliferation‚ i.e.‚ altered cell-cell and cell-matrix interaction‚ enhanced availability and utilization of energy-generating substrate (glycogen‚ glucose)‚ activation of transcriptional regulation‚ and cell cycle progression. Moreover‚ it is of considerable importance in the present context that insulin-initiated wg/Wnt-like signaling is inhibited by fatty acids3‚139-148 and in obesity149‚ as well as by cyclic AMP-mediated prevention of PDK1 membrane localization150. In addition‚ as discussed above‚ the tumor suppressor gene PTEN‚ the product of which is a lipid phosphatase that inhibits PI3K signaling‚ induces apoptosis in Jurkat T cells151 and is defective in many human cancers81.

3.3.2.1. Broader Significance of wg/Wnt-Like Signaling

That the interaction between growth factors and may transcend cell proliferation is suggested by evidence that components of the wg/Wntlike signaling pathway are operative in brain neurons (see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). It is reasonable to question what advantage might be served by expression of a pathway so intimately linked to proliferation control in a cell in which proliferation is precluded‚ such as the terminally differentiated neuron. Consideration of the energy requirements of cell growth and neuronal activation provides insight into this seeming paradox. In cortical neurons‚ presenilin-1 (PS1) may function as a scaffold protein‚ in a manner superficially resembling that of APC or axin152‚153. Thus‚ in the absence of wg/Wnt-like signaling PS1

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promotes formation of a complex in which it combines with the microtubule-associated protein and active (unphosphorylated) Hyperphosphorylation of by and other kinases leads ultimately to formation of the neurofibrillary tangles that are characteristic of Alzheimer Although these protein-protein interactions are well disease103. substantiated‚ there is recent evidence that the arm of wg/Wnt-like signaling that is related to nuclear abundance‚ turnover‚ and gene transcription may not be operative in neurons154. This would suggest that‚ perhaps better suited to its functioning in a post-mitotic cell‚ neuronal wg/Wnt-like signaling may be related predominantly or exclusively to metabolic regulation‚ reflecting the interaction among PI3K‚ Akt/PKB‚ and This “truncated” version of wg/Wnt-like signaling might also be regarded as functionally analogous to the non-proliferative effect of enterocyte trefoil factor described below. In both neuronal activation and mitosis‚ the rate at which ATP is generated is critical and is optimally supported by insulin or insulin-like enhancement of glycolytic fueling of the citric acid cycle and oxidative phosphorylation (see Chapter 11: Energetics of Neuronal Activation). A key element is that in both the activated neuron and the cell entering the mitotic cycle‚ active wg/Wnt-like signaling is necessary to phosphorylate and thus inactivate This would prevent phosphorylation not only of but also of pyruvate dehydrogenase103‚ the pivotal determinant of glycolytic fueling of the citric acid cycle that is required in both settings. In contrast‚ in the (non-neuronal) cell committed to a differentiation pathway‚ and/or exposed to increased amounts of fatty acids‚ the associated decrease in wg/Wnt-like signaling and therefore in phosphorylation would augment the latter’s activity. The resulting phosphorylation of would suppress transcriptional activation and stabilize E-cadherin-dependent cellcell adhesion. According to this formulation‚ then‚ the wg/Wnt-like signaling pathway‚ among its other effects‚ may be viewed as a general and perhaps ubiquitous mechanism that contributes to the regulation of ATP availability in a variety of circumstances. In addition to cell cycle progression‚ it may be harnessed for this purpose in support of other time- and energy-critical processes that are unpredictable‚ and in which maximal rates of ATP synthesis via oxidative phosphorylation are required; these include neuronal activation amd synaptogenesis in brain155 (also‚ see chapter 15.4.2.1. Wingless/WntLike Signaling)‚ and the rapid‚ non-proliferative‚ EGF- and PI3K/Akt/PKBmediated‚ trefoil factor-dependent epithelial restitution that follows intestinal mucosal injury130. In the latter‚ trefoil factor exerts an antiproliferative effect156. Accordingly‚“truncated” (used here to indicate non-mitogenic)

3.Nutrient and Energy Metabolism in Cell Proliferation

33

wg/Wnt-like signaling may in general subserve activation of protective‚ reparative‚ or other unpredictable cellular processes that are energy intensive but not necessarily linked to cell proliferation. In contrast‚ in cells which have exited the mitotic cycle and/or otherwise have relatively low energy demands‚ suppression of wg/Wnt permits maximal activation of phosphorylation reactions and their downstream effects. Operation of the pathway may be disturbed in either direction. Thus‚ may be inappropriately suppressed (i.e.‚ wg/Wnt active) in the APC mutant colonocyte‚ leading to impaired cell-cell adhesion and increased transcriptional activation‚ mitosis‚ and tumorigenesis. Conversely‚ may be inappropriately activated (i.e.‚ wg/Wnt suppressed) in the Alzheimer neuron‚ leading to inactivation of pyruvate dehydrogenase‚ impairment of glycolysis and pyruvate-fueled oxidative phosphorylation‚ hyperphosphorylation of with generation of neurofibrillary tangles‚ oxidative stress‚ and increased amyloid formation (see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). 3.3.2.2. Lithium The broad importance of wg/Wnt-like signaling in both neuronal and non-neuronal cells is further suggested by the central role and diverse effects of the small univalent cation lithium both directly157‚158 and indirectly159 inhibits thereby augmenting Akt/PKB signaling. By this mechanism‚ 1) mimics aspects of insulin/PI3K/Akt/PKB-stimulated glucose metabolism160; 2) exerts broad cellular protective effects159‚161‚162; 3) increases cytosolic abundance of and Tcf/lef transcriptional activity in fibroblasts158‚163; and 4) augments proliferation of non-neural cells164‚165. Individually and jointly‚ these interactions strongly support the concept that activation/inactivation is of central importance in both normal cell function and disease pathogenesis. 3.3.2.3. Reactive Oxygen Species The foregoing observations emphasize a critical role for aerobic glycolysis and oxidative phosphorylation in cell growth. It therefore seems surprising that‚ despite their potentially damaging effects‚ reactive oxygen species (ROS‚ e.g.‚ and superoxide anion — may be generated when mitochondrial energization and are high (see Chapter 7: Fatty Acids and Mitochondria; Cell Growth and Injury: Broader Implications)‚ and may also participate in the mediation of cell proliferation166-168. At least in part‚ this may reflect ROS inhibition of both the Forkhead family of transcriptional regulators168 (also‚ see 3.3.2.: Wingless/Wnt-Like Signaling:

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Convergence of Antecedents and the Unpredictable), as well as of protein tyrosine phosphatase1B169,170. The latter effect permits sustained tyrosine phosphorylation and augmented signaling by both receptor and non-receptor kinases including PI3K67,171,172 and Syk173, as well as as stabilization of Akt/PKB by both Hsp 27174 and Hsp90175,176. Moreover, hypoxia activates two mutually-reinforcing limbs of the cellular response to diminished oxygen availability67,70,177. First, electron transport chain generation of ROS indirectly activates PI3K/Akt/PKB signaling (see previous paragraph). Second, suppressed oxygen-dependent proline hydroxylation of the von Hippel-Lindau protein (VHL) activates thereby increasing transcription of genes related to cellular glucose uptake (e.g., GLUT 1, 3), glycolysis, and oxygen supply (e.g., erythropoietin, VEGF)67,70,177. Activation of PI3K/Akt/PKB signaling synergizes with the increase in expression of glycolytic genes. Moreover, it 92 activates , possibly accounting for a recognized ROS-induced activation of transcriptional regulation by and AP1178,180. Thus, a twopronged cellular response to hypoxia effects synergistic promotion and integration of cytoprotective patterns of metabolism, energetics, and gene expression. In addition, may be activated under normoxic 181 conditions by insulin , and in a neuronal cell defensive response to amyloid182 (also, see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). These and other factors may activate a multifaceted upregulation of and expression of its transcriptional targets, via wg/Wnt-like PI3K/Akt/PKB signaling70; it may include an Akt/PKB183 mediated VHL-independent activation of VEGF expression by 184 and suppression of forkhead transcriptional activity . Such observations provide important insight into metabolic aspects of growth control and raise equally important questions concerning the diverse effects of ROS, which may range from necrotic death to cell proliferation185. This diversity appears to reflect at least in part variations in ROS abundance185, persistence67, and interaction with cellular antioxidant defenses that are necessary to effect modulation of critical functions such as caspase activity186 and transcriptional regulation166,187,188. However, except for 67,177,189 hypoxia-induced electron transport chain generation of , and phagocyte or vascular NADPH oxidase, the origins of ROS involved in growth signaling are largely undefined166,167. Evidence to be considered below suggests that mitochondrial and extramitochondrial fatty acid oxidation contribute importantly to their generation (see Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). In summary, most dividing cells may utilize a range of nutrients but optimal growth depends primarily on increased oxidative phosphorylationmediated ATP biosynthesis. The increase in required to support this is


35

fueled largely by glycolysis and the citric acid cycle, reflecting at least in part wg/Wnt-like signaling and transactivation of glycolysis-related genes. The principal exceptions appear to be: 1) those cells in which the citric acid cycle is fueled directly by glutamine-derived 2oxo-glutarate, e.g., in small intestine in vivo and in certain tumor cells in vitro63,64; also, see Chapter 6.2.: Butyrate); and 2) glycolytic support of hypoxic proliferation in selected cell populations as noted above (see 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation). Even in the latter, however, hypoxia-induced mitochondrial generation of ROS indirectly promotes cell cycle progression, as discussed in the preceding paragraphs. In any event, implicit in all dividing cells including these exceptions is the relative exclusion of fatty acids as mitochondrial oxidative substrate (Fig. 3.1). Thus, cell proliferation may be the resultant of a constantly changing interaction among: 1) cell cycle energy demands; 2) genetic, metabolic, and energetic heterogeneity among cells, and among mitochondria within a given cell190-193; and 3) variation in microenvironmental abundance of nutrients, oxygen, and growth modulators. In this microcosm of natural selection, these variables would favor cell cycle completion and lineage survival for only the “fittest” of genotypes and gene expression patterns.

3.4.

REFERENCES

1. P. J. Randle, Regulatory interactions between lipids and carbohydrates: the glucose fatty acid cycle after 35 years, Diabetes Metab Rev 14:263-83 (1998). 2. G. Boden, and G. I. Shulman, Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction, Eur J Clin Invest 32 Suppl 3:14-23 (2002). 3. J. McGarry, Banting lecture, 2001: dysregulation of fatty acid metabolism in the etiology of type 2 diabetes, Diabetes 51:7-18 (2002). 4. T. A. Hopkins, M. C. Sugden, M. J. Holness, R. Kozak, J. R. Dyck, and G. D. Lopaschuk, Control of cardiac pyruvate dehydrogenase activity in peroxisome proliferator-activated receptor-alpha transgenic mice, Am J Physiol Heart Circ Physiol 285:H270-6 (2003). 5. J. Scaife, and H. Brohee, An investigation of factors influencing mitotic G2 delay in synchronous cultures of human kidney cells and X-irradiation, Can J Biochem 47:237-249 (1969). 6. S. Ishiguro, H. Yamaguchi, Y. Oka, and H. Miyamoto, Change in energy metabolism in the cell cycle of mouse L cells, Cell Structure Function 3:331-340 (1978). 7. S. Skog, B. Tribukait, and G. Sundius, Energy metabolism and ATP turnover time during the cell cycle of Ehrlich ascites tumour cells, Exp Cell Res 141:23-29 (1982). 8. S. Skog, B. Tribukait, and G. Sundius, Energy metabolism and ATP turnover time during the cell cycle in roentgen irradiated Ehrlich ascites tumour cells, Acta Radiol Oncol 22:369-379 (1983).

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9. C. Van den Bogert, P. Muus, C. Haanen, A. Pennings, T. Melis, and A. Kroon, Mitochondrial biogenesis and mitochondrial activity during the progression of the cell cycle of human leukemic cells, Exp Cell Res 178:143-153 (1988). 10. L. Xie, and D. Wang, Energy metabolism and ATP balance in animal cell cultivation using a stoichiometrically based reaction network, Biotech Bioengineering 521:591-601 (1996). 11. S. Talha, and L. Harel, Early stimulation of ATP turnover induced by growth factors: Synergistic effect of EGF and insulin and correlation with DNA synthesis, Exp Cell Res 158:311-320 (1985). 12. V. Jain, I. Gupta, and K. Lata, Energetics of cellular repair processes in a respiratorydeficient mutant of yeast, Rad Res 92:463-473 (1982). 13. Y. Li, G. Sattler, and H. Pitot, Oxaloacetate induces DNA synthesis and mitosis in primary cultured rat hepatocytes in the absence of EGF, Biochem Biophys Res Commun 193:1339-1346 (1993). 14. S. Linke, K. Clarkin, L. Di, A. Tsou, and G. Wahl, A reversible, p53-dependent G0/G1 cell cycle arrest induced by ribonucleotide depletion in the absence of detectable DNA damage, Genes Devel 10:934-947 (1996). 15. T. Gaal, M. Bartlett, W. Ross, C. Turnbough, Jr, and R. Gourse, Transcription regulation by intiating NTP concentration: rRNA synthesis in bacteria., Science 278:2092-2097 (1997). 16. D. Jones, Intracellular diffusion gradients of O2 and ATP, Am J Physiol 250:C663-C675 (1986). 17. E. Fontaine, C. Keriel, S. Lantuejoul, M. Rigoulet, X. Leverve, and V. Saks, Cytoplasmic cellular structures control permeability of outer mitochondrial membrane for ADP and oxidative phosphorylation in rat liver cells., Biochem Biophys Res Commun 213:138-146 (1995). 18. E. Takahashi, H. Endoh, and K. Doi, Intracellular gradients of supply to mitochondria in actively respiring single cardiomyocyte of rats, Am J Physiol 276:H718-724 (1999). 19. O. Warburg. Uber den stoffwechsel den tumoren: Arbeiten aus dem Kaiser WilhelmInstitut fur biologie, ed., J Springer, Berlin (1926), pp. 115-149. 20. P. Pedersen, Tumor mitochondria and the bioenergetics of cancer cells, Prog Exp Tumor Res 22:190-274 (1978). 21. F. Kallinowski, K. Schlenger, M. Kloes, M. Stohrer, and P. Vaupel, Tumor blood flow: the principal modulator of oxidative and glycolytic metabolism, and of the metabolic micromilieu of human tumor xenografts in vivo, Intl J Cancer 44: 266-272 (1989). 22. S. Mazurek, C. Boschek, and E. Eigenbrodt, The role of phosphometabolites in cell proliferation, energy metabolism, and tumor therapy, J Bioenerg Biomembranes 29:315330 (1997). 23. P. L. Pedersen, S. Mathupala, A. Rempel, J. F. Geschwind, and Y. H. Ko, Mitochondrial bound type II hexokinase: a key player in the growth and survival of many cancers and an ideal prospect for therapeutic intervention, Biochim Biophys Acta 1555:14-20 (2002). 24. P. Bannasch, Pathogenesis of hepatocellular carcinoma: sequential cellular, molecular, and metabolic changes., Prog Liver Dis 14:161-197 (1996). 25. M. Younes, L. Lechago, J. Somoano, M. Mosharaf, and J. Lechago, Wide expression of the human erythrocyte glucose transporter Glut1 in human cancers, Cancer Res 56:11641167 (1996). 26. J. Flier, M. Mueckler, P. Usher, and H. Lodish, Elevated levels of glucose transport and transporter messenger RNA are induced by ras or src oncogenes., Science 235:1492-1495 (1987).


37

27. M. Birnbaum, H. Haspel, and O. Rosen, Transformation of rat fibroblasts by FSV rapidly increases glucose transporter gene transcription., Science 235:1495-1498 (1987). 28. E. Riu, F. Bosch, and A. Valera, Prevention of diabetic alterations in transgenic mice overexpressing Myc in the liver, Proc Natl Acad Sci USA 93:2198-2202 (1996). 29. M. Tiedge, and S. Lenzen, Effects of sodium butyrate on glucose transporter and glucosephosphorylating enzyme gene expression in RINm5F insulinoma cells, J Molec Endocrinol 17:19-26 (1996). 30. B. Gelb, V. Adams, S. Jones, L. Griffin, G. Macgregor, and E. Mccabe, Targeting of hexokinase 1 to liver and hepatoma mitochondria., Proc Natl Acad Sci USA 89:202-206 (1992). 31. S. Mathupala, A. Rempel, and P. Pedersen, Glucose catabolism in cancer cells: identification and characterization of a marked activation response of the type II hexokinase gene to hypoxic conditions, J Biol Chem 276:43407-43412 (2001). 32. A. Goel, S. P. Mathupala, and P. L. Pedersen, Glucose metabolism in cancer. Evidence that demethylation events play a role in activating type II hexokinase gene expression, J Biol Chem 278:15333-40 (2003). 33. G. Beutner, A. Rück, B. Riede, and D. Brdiczka, Complexes between porin, hexokinase, mitochondrial creatine kinase and adenylate translocator display properties of the permeability transition pore: Implication for regulation of permeability transition by the kinases., Biochim Biophys Acta 1368:7-18 (1998). 34. N. Zamzami, C. Brenner, I. Marzo, S. Susin, and G. Kroemer, Subcellular and submitochondrial mode of action of Bcl-2-like oncoproteins, Oncogene 16:2265-2282 (1998). 35. I. Marzo, C. Brenner, N. Zamzami, et al., The permeability transition pore complex: a target for apoptosis regulation by caspases and bcl-2-related proteins, J Exper Med 187:1261-1271 (1998). 36. V. Lemeshko, Model of the outer membrane potential generation by the inner membrane of mitochondria, Biophys J 82:684-692 (2002). 37. J. Pastorino, N. Shulga, and J. Hoek, Mitochondrial binding of hexokinase II inhibits Baxinduced cytochrome c release and apoptosis, J Biol Chem 277:7610-7618 (2002). 38. M. Y. Vyssokikh, and D. Brdiczka, The function of complexes between the outer mitochondrial membrane pore (VDAC) and the adenine nucleotide translocase in regulation of energy metabolism and apoptosis, Acta Biochim Pol 50:389-404 (2003). 39. E. McCabe, Microcompartmentation of energy metabolism at the outer mitochondrial membrane: role in diabetes mellitus and other diseases, J Bioenerg Biomembranes 26:317325 (1994). 40. M. Fanciulli, M. Paggi, T. Bruno, et al., Glycolysis and growth rate in normal and in hexokinase-transfected NIH-3T3 cells., Oncol Res 6:405-409 (1994). 41. M. Board, A. Colquhoun, and E. Newsholme, High Km glucose-phosphorylating (glucokinase) activities in a range of tumor cell lines and inhibition of rates of tumor growth by the specific enzyme inhibitor mannoheptulose., Cancer Res 55:3278-3285 (1995). 42. D. Mayer, F. Klimek, A. Rempel, and P. Bannasch, Hexokinase expression in liver preneoplasia and neoplasia (Review), Biochem Soc Trans 25:122-127 (1997). 43. Y. Shinohara, T. Ishida, M. Hino, N. Yamazaki, Y. Baba, and H. Terada, Characterization of porin isoforms expressed in tumor cells, Eur J Biochem 267:6067-6073 (2000). 44. R. Ockner, Apoptosis and liver diseases: Recent concepts of mechanisms and significance, J Gastroenterol Hepatol 16:248-260 (2001). 45. L. Hue, and G. Rousseau, Fructose 2,6-bisphosphate and the control of glycolysis by growth factors, tumor promoters and oncogenes, Adv Enzyme Regul 33:97-110 (1993).

38

Chapter 3

46. M. Joaquin, J. Rosa, C. Salvado, et al., Hepatocyte growth factor and transforming growth factor beta regulate 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase gene expression in rat hepatocyte primary cultures, Biochem J 314:235-240 (1996). 47. G. Thoresen, and T. Christoffersen, Transforming growth factor beta 1 increases the phosphoenolpyruvate carboxykinase mRNA level in cultured rat hepatocytes, Cell Biol Intl 18:171-175 (1994). 48. K. Sasaki, T. Cripe, S. Koch, et al., Multihormonal regulation of phosphoenolpyruvate carboxykinase gene transcription: The dominant role of insulin, J Biol Chem 259:1524215251 (1984). 49. C. Fillat, A. Valera, and F. Bosch, Epidermal growth factor inhibits phosphoenolpyruvate carboxykinase gene expression in rat hepatocytes in primary culture., FEBS Lett. 318: 287-291 (1993). 50. D. Persons, N. Schek, B. Hall, and O. Finn, Increased expression of glycolysis-associated genes in oncogene-transformed and growth-accelerated states, Mol Carcinog 2:88-94 (1989). 51. O. Kaplan, J. Jaroszewski, P. Faustino, et al., Toxicity and effects of epidermal growth factor on glucose metabolism of MDA-468 human breast cancer cells, J Biol Chem 265:13641-13649(1990). 52. K. Yamada, and T. Noguchi, Regulation of pyruvate kinase M gene expression, Biochem Biophys Res Commun 256:257-262 (1999). 53. P. Bannasch, F. Klimek, and D. Mayer, Early bioenergetic changes in hepatocarcinogenesis: preneoplastic phenotypes mimic responses to insulin and thyroid hormone., J Bioenergetics Biomembranes 29:303-313 (1997). 54. D. Nehrbass, F. Klimek, and P. Bannasch, Overexpression of insulin receptor substrate-1 emerges early in hepatocarcinogenesis and elicits preneoplastic hepatic glycogenosis, Am J Pathol 152:341-345 (1998). 55. W. Tian, L. Braunstein, J. Pang, et al., Importance of glucose-6-phosphate dehydrogenase activity for cell growth, J Biol Chem 273:10609-10617 (1998). 56. S. Soboll, Regulation of energy metabolism in liver, J Bioenergetics Biomembranes 27:571-582 (1995). 57. S. Sweet, and G. Singh, Accumulation of human promyelocytic leukemic (HL-60) cells at two energetic cell cycle checkpoints, Cancer Res 55:5164-5167 (1995). 58. L. Sauer, and R. Dauchy, Ketone body, glucose, lactic acid, and amino acid utilization by tumors in vivo in fasted rats, Cancer Res 43:3497-3503 (1983). 59. J. Boren, M. Cascante, S. Marin, et al., Gleevec (STI571) influences metabolic enzyme activities and glucose carbon flow toward nucleic acid and fatty acid synthesis in myeloid tumor cells., J Biol Chem 276:37747-37753 (2001). 60. S. Miccadei, M. Fanciulli, T. Bruno, M. Paggi, and A. Floridi, Energy metabolism of adriamycin-sensitive and -resistant Ehrlich ascites tumor cells, Oncol Res 8:27-35 (1996). 61. R. Nakashima, M. Paggi, and P. Pedersen, Contributions of glycolysis and oxidative phosphorylation to adenosine 5’-triphosphate production in AS-30D hepatoma cells, Cancer Res 44:5702-5706 (1984). 62. V. Goossens, J. Grooten, and W. Fiers, The oxidative metabolism of glutamine: A modulator of reactive oxygen intermediate-mediated cytotoxicity of tumor necrosis factor in L929 fibrosarcoma cells, J Biol Chem 271:192-196 (1996). 63. Z. Kovacevic, and J. Mcgivan, Mitochondrial metabolism of glutamine and glutamate and its physiological significance, Physiol Rev 63:547-605 (1983). 64. M. Board, S. Humm, and E. Newsholme, Maximum activities of key enzymes of glycolysis, glutaminolysis, pentose phosphate pathway and tricarboxylic acid cycle in normal, neoplastic and suppressed cells., Biochem J 265:503-509 (1990).


39

65. L. Chen, Mitochondrial membrane potential in living cells., Annu Rev Cell Biol 4:155181 (1988). 66. P. Maxwell, G. Dachs, J. Gleadle, et al., Hypoxia-inducible factor-1 modulates gene expression in solid tumors and influences both angiogenesis and tumor growth, Proc Natl Acad Sci USA 94:8104-8109 (1997). 67. N. Chandel, D. McClintock, C. Feliciano, et al., Reactive oxygfen species generated at mitochondrial complex III stabilize hypoxia-inducible factor-1 alpha during hypoxia: a mechanism of O2 sensing., J Biol Chem 275:25130-25138 (2000). 68. O. Genbacev, Y. Zhou, J. Ludlow, and S. Fisher, Regulation of human placental development by oxygen tension., Science 277:1669-1672 (1997). 69. E. Greiner, M. Guppy, and K. Brand, Glucose is essential for proliferation and the glycolytic enzyme induction that provokes a transition to glycolytic energy production, J Biol Chem 269:31484-31490 (1994). 70. G. Semenza, HIF-1 and tumor progression: pathophysiology and therapeutics., Trends Molec Med 8:S62-S67 (2002). 71. N. Goda, H. Ryan, B. Khadivi, W. McNulty, R. Rickert, and R. Johnson, Hypoxiainducible factor 1a is essential for cell cycle arrest during hypoxia, Molec Cell Biol 23:359-369 (2003). 72. Z. Z. Chong, J. Q. Kang, and K. Maiese, Erythropoietin is a novel vascular protectant through activation of Akt1 and mitochondrial modulation of cysteine proteases, Circulation 106:2973-9 (2002). 73. T. Graeber, C. Osmanian, T. Jacks, et al., Hypoxia-mediated selection of cells with diminished apoptotic potential in solid tumours, Nature 379:88-91 (1996). 74. I. Sussman, M. Erecinska, and D. F. Wilson, Regulation of cellular energy metabolism: the Crabtree effect, Biochim Biophys Acta 591:209-23 (1980). 75. L. Wojtczak, V. Teplova, K. Bogucka, et al., Effect of glucose and deoxyglucose on the redistribution of calcium in ehrlich ascites tumour and Zajdela hepatoma cells and its consequences for mitochondrial energetics: Further arguments for the role of Ca(2+) in the mechanism of the crabtree effect, Eur J Biochem 263:495-501 (1999B). 76. S. Rodriguez-Enriquez, O. Juarez, J. Rodriguez-Zavala, and R. Moreno-Sanchez, Multisite control of the Crabtree effect in ascites hepatoma cells, Eur J Biochem 268:2512-2519 (2001). 77. M. Peifer, Beta-catenin as oncogene: the smoking gun, Science 275:1752-1753 (1997). 78. M. Hart, R. De Los Santos, I. Albert, B. Rubinfeld, and P. Polakis, Downregulation of beta-catenin by human Axin and its association with the APC tumor suppressor, betacatenin and GSK3 beta, Current Biol 8:573-581 (1998). 79. F. McCormick, Signalling networks that cause cancer, Trends Biochem Sci 24:M53-M56 (1999). 80. D. Plas, S. Talapatra, A. Edinger, J. Rathmell, and C. Thompson, Akt and Bcl-xL promote growth factor-independent survival through distinct effects on mitochondrial physiology, J Biol Chem 276:12041-12048 (2001). 81. L. Cantley, and B. Neel, New insights into tumor suppression: PTEN suppresses tumor formation by restraining the phosphoinositide 3-kinase/AKT pathway., Proc Natl Acad Sci USA 96:4240-4245 (1999). 82. M. M. Hill, and B. A. Hemmings, Inhibition of protein kinase B/Akt. implications for cancer therapy, Pharmacol Ther 93:243-51 (2002). 83. B. Vanhaesebroeck, and D. Alessi, The PI3K-PDK1 connection: more than just a road to PKB, Biochem J 346:561-576 (2000). 84. R. Tuttle, N. Gill, W. Pugh, et al., Regulation of pancreatic beta-cell growth and survival by the serine/threonine protein kinase Akt1/PKBalpha, Nat Med 7:1133-1137 (2001).

40

Chapter 3

85. K. A. Longo, J. A. Kennell, M. J. Ochocinska, S. E. Ross, W. S. Wright, and O. A. MacDougald, Wnt signaling protects 3T3-L1 preadipocytes from apoptosis through induction of insulin-like growth factors, J Biol Chem 277:38239-44 (2002). 86. K. Gottlob, N. Majewski, S. Kennedy, E. Kandel, R. Robey, and N. Hay, Inhibition of early apoptotic events by Akt/PKB is dependent on the first committed step of glycolysis and mitochondrial hexokinase, Genes Devel 15:1406-1418 (2001). 87. A. Ahmad, S. Ahmad, B. K. Schneider, C. B. Allen, L. Y. Chang, and C. W. White, Elevated expression of hexokinase II protects human lung epithelial-like AS49 cells against oxidative injury, Am J Physiol Lung Cell Mol Physiol 283:L573-84 (2002). 88. T. Burdon, A. Smith, and P. Savatier, Signalling, cell cycle and pluripotency in embryonic stem cells, Trends Cell Biol 12:432-8 (2002). 89. T. Reya, A. W. Duncan, L. Ailles, et al., A role for Wnt signalling in self-renewal of haematopoietic stem cells, Nature 423:409-14 (2003). 90. S. Datta, H. Dudek, X. Tao, et al., Akt phosphorylation of BAD couples survival signals to the cell-intrinsic death machinery., Cell 91:231-241 (1997). 91. M. Cardone, N. Roy, H. Stennicke, et al., Regulation of cell death protease caspase-9 by phosphorylation., Science 282:1318-1321 (1998). 92. O. Ozes, L. Mayo, J. Gustin, S. Pfeffer, L. Pfeffer, and D. Donner, NF-kappaB activation by tumour necrosis factor requires the Akt serine-threonine kinase, Nature 401:82-85 (1999). 93. R. G. Jones, A. R. Elford, M. J. Parsons, et al., CD28-dependent activation of protein kinase B/Akt blocks Fas-mediated apoptosis by preventing death-inducing signaling complex assembly, J Exp Med 196:335-48 (2002). 94. D. McClintock, M. Santore, V. Lee, et al., Bcl-2 family members and functional electron transport chain regulate oxygen deprivation-induced cell death, Mol Cell Biol 22:94-104 (2002). 95. A. Brunet, A. Bonni, M. Zigmond, et al., Akt promotes cell survival by phosphorylating and inhibiting a Forkhead transcription factor., Cell 96:857-868 (1999). 96. E. Shtivelman, J. Sussman, and D. Stokoe, A role for PI 3-kinase and PKB activity in the G2/M phase of the cell cycle, Curr Biol 12:919-24 (2002). 97. R. Medema, G. Kops, J. Bos, and B. Burgering, AFX-like Forkhead transcription factors mediate cell-cycle regulation by Ras and PKB through p27kip1, Nature 404:782-788 (2000). 98. H. Tran, A. Brunet, J. Grenier, et al., DNA repair pathway stimulated by the forkhead transcription factor FOXO3a through the Gadd45 protein, Science 296:530-534 (2002). 99. Y. Honda, and S. Honda, The daf-2 gene network for longevity regulates oxidative stress resistance and Mn-superoxide dismutase gene expression in Caenorhabditis elegans, FASEB J 13:1385-1393 (1999). 100. S. Ghaffari, Z. Jagani, C. Kitidis, H. F. Lodish, and R. Khosravi-Far, Cytokines and BCR-ABL mediate suppression of TRAIL-induced apoptosis through inhibition of forkhead FOXO3a transcription factor, Proc Natl Acad Sci U S A 100:6523-8 (2003). 101. B. Gumbiner, Carcinogenesis: a balance between beta-catenin and APC, Current Biol 7:R443-R446 (1997). 102. A. Yap, W, Brieher, and B. Gumbiner, Molecular and functional analysis ofcadherinbased adherens junctions, Ann Rev Cell Dev Biol 13:119-146 (1997). 103. M. Hoshi, A. Takashima, K. Noguchi, et al., Regulation of mitochondrial pyruvate dehydrogenase activity by tau protein kinase I/glycogen synthase kinase 3beta in brain, Proc Natl Acad Sci USA 93:2719-2723 (1996). 104. J. Diehl, M. Cheng, M. Roussel, and C. Sherr, Glycogen synthase kinase-3beta regulates cyclin D1 proteolysis and subcellular localization., Genes Devel 12:3499-3511 (1998).


41

105. O. Tetsu, and F. Mccormick, Beta-catenin regulates expression of cyclin D1 in colon carcinoma cells, Nature 398:422-426 (1999). 106. H. C. Crawford, B. M. Fingleton, L. A. Rudolph-Owen, et al., The metalloproteinase matrilysin is a target of beta-catenin transactivation in intestinal tumors, Oncogene 18:2883-91 (1999). 107. T. He, A. Sparks, C. Rago, et al., Identification of c-MYC as a target of the APC pathway, Science 281:1509-1512 (1998). 108. G. Christofori, and H. Semb, The role of the cell-adhesion molecule E-cadherin as a tumour-suppressor gene., Trends Biochem Sci 24:73-76 (1999). 109. H. Hermeking, C. Rago, M. Schuhmacher, et al., Identification of CDK4 as a target of c-MYC, Proc Natl Acad Sci USA 97:2229-2234 (2000). 110. D. R. Wonsey, K. I. Zeller, and C. V. Dang, The c-Myc target gene PRDX3 is required for mitochondrial homeostasis and neoplastic transformation, Proc Natl Acad Sci U S A 99:6649-54 (2002). 111. P. Staller, K. Peukert, A. Kiermaier, et al., Repression of p15INK4b expression by Myc through association with Miz-1, Nature Cell Biol 3:392-399 (2001). 112. I. Perez-Roger, S. Kim, B. Griffiths, A. Sewing, and H. Land, Cyclins D1 and D2 mediate Myc-induced proliferation via sequestration of p27(Kip1) and p21(Cip1), EMBO J 18:5310-5320 (1999). 113. C. Sherr, and J. Roberts, CDK inhibitors: positive and negative regulators of Gl-phase progression, Genes Dev 13:1501-12 (1999). 114. O. Coqueret, New roles for p21 and p27 cell-cycle inhibitors: a function for each cell compartment?, Trends Cell Biol 13:65-70 (2003). 115. L. Rossig, C. Badorff, Y. Holzmann, A. M. Zeiher, and S. Dimmeler, Glycogen synthase kinase-3 couples AKT-dependent signaling to the regulation of p21Cip1 degradation, J Biol Chem 277:9684-9 (2002). 116. Y. Li, D. Dowbenko, and L. A. Lasky, AKT/PKB phosphorylation of p21Cip/WAF1 enhances protein stability of p21Cip/WAF1 and promotes cell survival, J Biol Chem 277:11352-61 (2002). 117. B. Zhou, Y. Liao, W. Xia, B. Spohn, M. Lee, and M. Hung, Cytoplasmic localization of p21Cip1/WAF1 by Akt-induced phosphorylation in HER-2/neu-overexpressing cells, Nat Cell Biol 3:245-252 (2001). 118. I. Shin, F. M. Yakes, F. Rojo, et al., PKB/Akt mediates cell-cycle progression by phosphorylation of p27(Kip1) at threonine 157 and modulation of its cellular localization, Nat Med 8:1145-52 (2002). 119. J. Liang, J. Zubovitz, T. Petrocelli, et al., PKB/Akt phosphorylates p27, impairs nuclear import of p27 and opposes p27-mediated G1 arrest, Nat Med 8:1153-60 (2002). 120. A. Kohn, S. Summers, M. Birnbaum, and R. Roth, Expression of a constitutively active Akt Ser/Thr kinase in 3T3-L1 adipocytes stimulates glucose uptake and glucose transporter 4 translocation, J Biol Chem 271:31372-31378 (1996). 121. J. Deprez, D. Vertommen, D. Alessi, L. Hue, and M. Rider, Phosphorylation and activation of heart 6-phosphofructo-2-kinase by protein kinase B and other protein kinases of the insulin signaling cascades., J Biol Chem 272:17269-17275 (1997). 122. D. Corcos, S. Vaulont, N. Denis, et al., Expression of c-myc is under dietary control in rat liver, Oncogene Res 1:193-199 (1987). 123. J. W. Kim, J. E. Lee, M. J. Kim, E. G. Cho, S. G. Cho, and E. J. Choi, Glycogen synthase kinase 3 beta is a natural activator of mitogen-activated protein kinase/extracellular signal-regulated kinase kinase kinase 1 (MEKK1), J Biol Chem 278:13995-14001 (2003).

42

Chapter 3

124. K. Hoeflich, J. Luo, E. Rubie, M. Tsao, O. Jin, and J. Woodgett, Requirement for glycogen synthase kinase-3beta in cell survival and NF-kappaB activation, Nature 406:8690 (2000). 125. R. F. Schwabe, and D. A. Brenner, Role of glycogen synthase kinase-3 in TNF-alphainduced NF-kappaB activation and apoptosis in hepatocytes, Am J Physiol Gastrointest Liver Physiol 283:G204-11 (2002). 126. C. Sutherland, I. Leighton, and P. Cohen, Inactivation of glycogen synthase kinase-3 beta by phosphorylation: new kinase connections in insulin and growth-factor signalling, Biochem J 296:15-19 (1993). 127. H. Eldar-Finkelman, and E. Krebs, Phosphorylation of insulin receptor substrate 1 by glycogen synthase kinase 3 impairs insulin action., Proc Natl Acad Sci USA 94:9660-9664 (1997). 128. P. Shepherd, D. Withers, and K. Siddle, Phosphoinositide 3-kinase: the key switch mechanism in insulin signalling, Biochem J 333:471-490 (1998). 129. D. Decraene, P. Agostinis, R. Bouillon, H. Degreef, and M. Garmyn, Insulin-like growth factor-1-mediated AKT activation postpones the onset of ultraviolet B-induced apoptosis, providing more time for cyclobutane thymine dimer removal in primary human keratinocytes, J Biol Chem 277:32587-95 (2002). 130. D. Taupin, K. Kinoshita, and D. Podolsky, Intestinal trefoil factor confers colonic epithelial resistance to apoptosis, Proc Natl Acad Sci USA 97:799-804 (2000). 131. G. Skouteris, and E. Georgakopoulos, Hepatocyte growth factor-induced proliferation of primary hepatocytes is mediated by activation of phosphatidylinositol 3-kinase, Biochem Biophys Res Commun 218:229-233 (1996). 132. J. Papkoff, and M. Aikawa, WNT-1 and HGF regulate GSK3- beta activity and betacatenin signaling in mammary epithelial cells, Biochem Biophys Res Comm 247:851-858 (1998). 133. S. Andjelic, C. Hsia, H. Suzuki, T. Kadowaki, S. Koyasu, and H.-C. Liou, Phosphatidylinositol 3-kinase and NF-kB/Rel are at the divergence of CD40-mediated proliferation and survival pathways., J Immunol 165: 3860-3867 (2000). 134. K. Willert, J. D. Brown, E. Danenberg, et al., Wnt proteins are lipid-modified and can act as stem cell growth factors, Nature 423:448-52 (2003). 135. M. Delcommenne, C. Tan, V. Gray, L. Rue, J. Woodgett, and S. Dedhar, Phosphoinositide-3-OH kinase-dependent regulation of glycogen synthase kinase 3 and protein kinase B/AKT by the integrin-linked kinase., Proc Natl Acad Sci USA 95:1121111216(1998). 136. D. Cook, M. Fry, K. Hughes, R. Sumathipala, J. Woodgett, and T. Dale, Wingless inactivates glycogen synthase kinase-3 via an intracellular signalling pathway which involves a protein kinase C., Embo J 15:4526-4536 (1996). 137. E. Dempsey, A. Newton, D. Mochly-Rosen, et al., Protein kinase C isozymes and the regulation of diverse cell responses., Am J Physiol Lung Cell Mol Physiol 279:L429-L438 (2000). 138. N. Murray, L. Davidson, R. Chapkin, W. Gustafson, D. Shattenberg, and A. Fields, Overexpression of protein kinase C betaII induces colonic hyperproliferation and increased sensitivity to colon carcinogenesis, J Cell Biol 145:699-711 (1999). 139. X. Chen, N. Iqbal, and G. Boden, The effects of free fatty acids on gluconeogenesis and glycogenolysis in normal subjects., J Clin Invest 103:365-372 (1999). 140. A. Dresner, D. Laurent, M. Marcucci, et al., Effects of free fatty acids on glucose transport and IRS-1-associated phosphatidylinositol 3-kinase activity., J Clin Invest 103:253-259 (1999).


43

141. M. E. Griffin, M. J. Marcucci, G. W. Cline, et al., Free fatty acid-induced insulin resistance is associated with activation of protein kinase C theta and alterations in the insulin signaling cascade, Diabetes 48:1270-4 (1999). 142. K. Cusi, K. Maezono, A. Osman, et al., Insulin resistance differentially affects the PI 3kinase- and MAP kinase-mediated signaling in human muscle., J Clin Invest 105:311-320 (2000). 143. H. Steinberg, G. Paradisi, G. Hook, K. Crowder, J. Cronin, and A. Baron, Free fatty acid elevation impairs insulin-mediated vasodilation and nitric oxide production, Diabetes 49:1231-1238(2000). 144. P. Arner, Insulin resistance in type 2 diabetes: role of fatty acids, Diabetes Metab Res Rev 18 Suppl 2:S5-9 (2002). 145. Y. Kruszynska, D. Worrall, J. Ofrecio, J. Frias, G. Macaraeg, and J. Olefsky, Fatty acidinduced insulin resistance: decreased muscle PI3K activation but unchanged Akt phosphorylation, J Clin Endocrinol Metab 87:226-234 (2002). 146. T. K. Lam, H. Yoshii, C. A. Haber, et al., Free fatty acid-induced hepatic insulin resistance: a potential role for protein kinase C-delta, Am J Physiol Endocrinol Metab 283:E682-91 (2002). 147. C. L. Soltys, L. Buchholz, M. Gandhi, A. S. Clanachan, K. Walsh, and J. R. Dyck, Phosphorylation of cardiac protein kinase B is regulated by palmitate, Am J Physiol Heart Circ Physiol 283:H1056-64 (2002). 148. C. Yu, Y. Chen, G. W. Cline, et al., Mechanism by which fatty acids inhibit insulin activation of insulin receptor substrate-1 (IRS-1)-associated phosphatidylinositol 3-kinase activity in muscle, J Biol Chem 277:50230-50236 (2002). 149. Z. Jiang, Y. Lin, A. Clemont, et al., Characterization of selective resistance to insulin signaling in the vasculature of obese Zucker (fa/fa) rats, J Clin Invest 104:447-457 (1999). 150. S. Kim, K. Jee, D. Kim, H. Koh, and J. Chung, Cyclic AMP inhibits Akt activity by blocking the membrane localization of PDK1, J Biol Chem 276:12864-70 (2001). 151. Z. Xu, D. Stokoe, L. P. Kane, and A. Weiss, The inducible expression of the tumor suppressor gene PTEN promotes apoptosis and decreases cell size by inhibiting the PI3K/Akt pathway in Jurkat T cells, Cell Growth Differ 13:285-96 (2002). 152. A. Takashima, M. Murayama, O. Murayama, et al., Presenilin 1 associates with glycogen synthase kinase-3beta and its substrate tau, Proc Natl Acad Sci USA 95:96379641 (1998). 153. M. Murayama, S. Tanaka, J. Palacino, et al., Direct association of presenilin-1 with betacatenin, FEBS Letters 433:73-77 (1998). 154. F. Kirschenbaum, S.-C. Hsu, B. Cordell, and J. McCarthy, Substitution of a glycogen synthase kinase-3b phosphorylation site in presenilin 1 separates presenilin function from b-catenin signaling, J Biol Chem 276:7266-7375 (2001). 155. M. Packard, D. Mathew, and V. Budnik, Wnts and TGF beta in synaptogenesis: old friends signalling at new places, Nat Rev Neurosci 4:113-20 (2003). 156. C. Bossenmeyer-Pourie, R. Kannan, S. Ribieras, et al., The trefoil factor 1 participates in gastrointestinal cell differentiation by delaying G1-S phase transition and reducing apoptosis, J Cell Biol 157:761-70 (2002). 157. P. Klein, and D. Melton, A molecular mechanism for the effect of lithium on development, Proc Natl Acad Sci USA 93:8455-8459 (1996). 158. V. Stambolic, L. Ruel, and J. Woodgett, Lithium inhibits glycogen synthase kinase-3 activity and mimics wingless signalling in intact cells, Curr Biol 6:1664-1668 (1996). 159. E. Chalecka-Franaszek, and D. Chuang, Lithium activates the serine/threonine kinase Akt-1 and suppresses glutamate-induced inhibition of Akt-1 activity in neurons., Proc Natl Acad Sci USA 96:8745-8750 (1999).

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160. S. Summers, A. Kao, A. Kohn, et al., The role of glycogen synthase kinase 3beta in insulin-stimulated glucose metabolism, J Biol Chem 274:17934-17940 (1999). 161. S. Nonaka, C. Hough, and D. Chuang, Chronic lithium treatment robustly protects neurons in the central nervous system against excitotoxicity by inhibiting N-methyl-Daspartate receptor-mediated calcium influx, Proc Natl Acad Sci USA 95: 2642-2647 (1998). 162. H. Wei, P. Leeds, Y. Qian, W. Wei, R.-W. Chen, and D.-M. Chuang, Beta-amyloid peptide-induced death of PC 12 cells and cerebellar granule cell neurons is inhibited by long-term lithium treatment, Eur J Pharmacol 392:117-123 (2000). 163. F. Staal, B. Burgering, M. van de Wetering, and H. Clevers, Tcf-1-mediated transcription in T lymphocytes: differential role for glycogen synthase kinase-3 in fibroblasts and T cells, Intl Immunol 11:317-323 (1999). 164. K. Ptashne, F. Stockdale, and S. Conlon, Initiation of DNA synthesis in mammary epithelium and mammary tumors by lithium ions, J Cellular Physiol 103:41-6 (1980). 165. H. Cui, Y. Meng, and R. Bulleit, Inhibition of glycogen synthase kinase 3beta activity regulates proliferation of cultured cerebellar granule cells., Brain Research Devel Brain Res 111:177-188 (1998). 166. T. Finkel, Redox-dependent signal transduction., FEBS Letters 476:52-54 (2000A). 167. V. Thannickal, R. Day, S. Klinz, M. Bastien, J. Larios, and B. Fanburg, Ras-dependent and -independent regulation of reactive oxygen species by mitogenic growth factors and TGF-b1, FASEB J 14:1741-1748 (2000). 168. S. Nemoto, and T. Finkel, Redox regulation of forkhead proteins through a p66shcdependent signaling pathway, Science 295:2450-2452 (2002). 169. S. Lee, K. Kwon, S. Kim, and S. Rhee, Reversible inactivation of protein-tyrosine phosphatase 1B in A431 cells stimulated with epidermal growth factor, J Biol Chem 273:15366-15372(1998). 170. W. Barrett, J. DeGnore, Y. Keng, Z. Zhang, M. Yim, and P. Chock, Roles of superoxide radical anion in signal transduction mediated by reversible regulation of protein-tyrosine phosphatase 1B., J Biol Chem 274:34543-34546 (1999). 171. Y. Bae, J. Sung, O. Kim, et al., Platelet-derived growth factor-induced production requires the activation of phosphatidylinositol 3-kinase., J Biol Chem 275:10527-10531 (2000). 172. S. Qin, E. Stadtman, and P. Chock, Regulation of oxidative stress-induced calcium release by phosphatidylinositol 3-kinase and Bruton’s tyrosine kinase in B cells, Proc Natl Acad Sci USA 97:7118-7123 (2000). 173. J. Ding, T. Takano, S. Gao, et al., Syk is required for the activation of Akt survival pathway in B cells exposed to oxidative stress., J Biol Chem 275:30873-30877 (2000). 174. K. M. Mearow, M. E. Dodge, M. Rahimtula, and C. Yegappan, Stress-mediated signaling in PC12 cells - the role of the small heat shock protein, Hsp27, and Akt in protecting cells from heat stress and nerve growth factor withdrawal, J Neurochem 83:452-62 (2002). 175. S. Sato, N. Fujita, and T. Tsuruo, Modulation of Akt kinase activity by binding to Hsp90, Proc Natl Acad Sci USA 97:10832-10837 (2000). 176. A. D. Basso, D. B. Solit, G. Chiosis, B. Giri, P. Tsichlis, and N. Rosen, Akt forms an intracellular complex with heat shock protein 90 (Hsp90) and Cdc37 and is destabilized by inhibitors of Hsp90 function, J Biol Chem 277:39858-66 (2002). 177. C. Schroedl, D. S. McClintock, G. R. Budinger, and N. S. Chandel, Hypoxic but not anoxic stabilization of HIF-1 alpha requires mitochondrial reactive oxygen species, Am J Physiol Lung Cell Mol Physiol 283:L922-31 (2002).


45

178. C. Sen, and L. Packer, Antioxidant and redox regulation of gene transcription, FASEB J 10:709-720 (1996). 179. V. Lakshminarayanan, E. Drab-Weiss, and K. Roebuck, and tumor necrosis factoralpha induce differential binding of the redox-responsive transcription factors AP-1 and NF-kappaB to the interleukin-8 promoter in endothelial and epithelial cells, J Biol Chem 273:32670-32678 (1998). 180. E. Shaulian, and M. Karin, AP-1 as a regulator of cell life and death, Nat Cell Biol 4:E131-6 (2002). 181. E. Zelzer, Y. Levy, C. Kahana, B. Shilo, M. Rubinstein, and B. Cohen, Insulin induces transcripition of target genes through the hypoxia-inducible factor HIF-1alpha/ARNT, EMBO J 17:5085-5094 (1998). 182. T. Soucek, R. Cumming, R. Dargusch, P. Maher, and D. Schubert, The regulation of glucose metabolism by HIF-1 mediates a neuroprotective response to amyloid beta peptide, Neuron 39:43-56 (2003). 183. V. Easwaran, S. H. Lee, L. Inge, et al., beta-Catenin regulates vascular endothelial growth factor expression in colon cancer, Cancer Res 63:3145-53 (2003). 184. T. T. Tang, and L. A. Lasky, The forkhead transcription factor FOXO4 induces the down-regulation of hypoxia-inducible factor 1 alpha by a von Hippel-Lindau proteinindependent mechanism, J Biol Chem 278:30125-35 (2003). 185. J. Dypbukt, M. Ankarcrona, M. Burkitt, et al., Different prooxidant levels stimulate growth, trigger apoptosis, or produce necrosis of insulin-secreting RINm5F cells: The role of intracellular polyamines., J Biol Chem 269:30553-30560 (1994). 186. V. Borutaite, and G. Brown, Caspases are reversibly inactivated by hydrogen peroxide., FEBS Lett. 500:114-118 (2001). 187. R. Ockner, R. Kaikaus, and N. Bass, Fatty acid metabolism and the pathogenesis of hepatocellular carcinoma: review and hypothesis, Hepatology 18:669-676 (1993). 188. Q. Zhang, D. Piston, and R. Goodman, Regulation of corepressor function by nuclear NADH, Science 295:1895-1897 (2002). 189. F. H. Agani, M. Puchowicz, J. C. Chavez, P. Pichiule, and J. LaManna, Role of nitric oxide in the regulation of HIF-1 alpha expression during hypoxia, Am J Physiol Cell Physiol 283:C178-86 (2002). 190. T. J. Collins, M. J. Berridge, P. Lipp, and M. D. Bootman, Mitochondria are morphologically and functionally heterogeneous within cells, Embo J 21:1616-27 (2002). 191. M. B. Elowitz, A. J. Levine, E. D. Siggia, and P. S. Swain, Stochastic gene expression in a single cell, Science 297:1183-6 (2002). 192. N. Fedoroff, and W. Fontana, Genetic networks. Small numbers of big molecules, Science 297:1129-31 (2002). 193. J. M. Levsky, S. M. Shenoy, R. C. Pezo, and R. H. Singer, Single-cell gene expression profiling, Science 297:836-40 (2002).

Chapter 4 Fatty Acids and Growth Regulation

4.1.

INTRODUCTION

To this point, emphasis has been placed on the importance of aerobic and anaerobic glycolysis in the fueling of cell cycle progression, and the obligatory exclusion of mitochondrial fatty acid oxidation. Perhaps somewhat unjustly, this focus has painted fatty acids as a whole with a broad brush as either non-contributory to the process or, worse, deleterious. In fact, as will be discussed in the ensuing paragraphs, it is quite clear that fatty acids are essential to cell proliferation. Although unlike glucose in their exclusion from its energization, they are obligatory for membrane and eicosanoid biogenesis, and typically are synthesized de novo in proliferating cells, both normal and transformed. They also play an important modulatory role in the proliferative process, reflecting substantial differences among the subclasses of fatty acids. Their influence in this regard has been documented in several settings, discussed below, in which their distinctive effects on cell growth are directly related to corresponding differences in their metabolism. These differences are exemplified by the role of fatty acids in growth promotion and in the biogenesis of membranes and eicosanoids, as contrasted with the propensity of fatty acids to undergo mitochondrial oxidation and inhibit growth, yet also to ameliorate intramitochondrial oxidative stress. The relationship between fatty acid metabolism and growth regulation is further highlighted by striking and unexpected parallels between host and tumor metabolism in cancer cachexia and the evolving metabolic patterns that characterize, and control, liver regeneration.

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48

4.2.

OMEGA-6

FATTY ACIDS

Early cell culture studies showed that linoleic acid (18:2, and certain other unsaturated fatty acids stimulated the growth of rat mammary carcinoma and normal cells in culture, and accounted entirely for the growth stimulatory effect of serum1-3. More recent evidence suggests that fatty acids increase ROS generation and activation of the transcriptional regulators and AP-1 in vitro4 and in vivo5,6. Although excessive levels of ROS result in cell injury, low/moderate levels may activate PI3K/Akt/PKB signaling and thereby contribute in this manner to the linkage between ROS and cell growth discussed above (see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable, and Chapter 7.4.2.2.: Paradoxical Benefits of Adversity). Other growth-related in vivo effects of fatty acids also have been demonstrated. Thus, fasting induced Jensen rat sarcoma growth — — incorporation) within 7 hours 7 , paralleling increases in plasma free fatty acids (FFA), glycerol, ketone bodies, and triglycerides, single pass tumor uptake of which averaged 4060% (ketone body uptake not quantified). Refeeding suppressed incorporation to control levels within 2 hours. The fasting response was attributed to an “adipose-derived nutrient”, consistent with its occurrence in adults but not young in which adipose tissue is sparse. Furthermore, in adult but not young rats, acute streptozotocin-induced diabetes mellitus increased incorporation in both Jensen sarcomas and Morris hepatomas 7288CTC within two to four hours; over the ensuing 3 days, tumor weight increased as host weight decreased8. Thus, contrasting effects on adult and young animals served to dissociate tumor growth from plasma glucose which decreased in all fasted and increased in all diabetic animals, and from plasma insulin which decreased in all diabetic animals. Tumor growth despite the antiproliferative effect of ketone bodies9,10, and see below) suggests modulation of this effect by growthpromoting factors, e.g., demonstrated increases in plasma concentrations of glucose and fatty acids. These studies were extended to in situ perfusion of 7288CTC Morris hepatomas in Buffalo rats with blood from variously treated donor animals11. Perfusion of tumors in fed rats with blood from starved animals (presumably FFA-rich) increased tumor incorporation, whereas perfusion of tumors in starved rats with normolipemic blood from fed donors suppressed incorporation. Based on reconstitution experiments, it was concluded that essential fatty acids in the perfusate, i.e., linoleic (18:2, and arachidonic (20:4, were

Fatty Acids and Growth Regulation

49

“rate-limiting” for stimulation of tumor incorporation; the nonessential saturated and monounsaturated fatty acids were inactive. Singlepass FFA uptake was similar in Morris hepatomas 7288CTC and 7777 and Jensen sarcomas12. Fasting also increased the number of mitoses in hepatocytes and pancreatic acinar cells in intact normal adult rats13,14. In vitro stimulation of cell growth by linoleic acid (18:2, was related to expression of liver fatty acid binding protein (L-FABP), expression of which is also increased during liver regeneration and in proliferating hyperplastic and neoplastic hepatocytes15,16. The role of fatty acids in tumor growth was further elucidated in studies of two transplantable colon carcinomas, i.e., cachexia-inducing MAC-16 and cachexia-non-inducing MAC-1317-19. Despite contrasting tumor effects on host nutrition, MAC-16- and MAC-13-bearing animals exhibited similar in vitro fatty acid synthase activities and lipogenesis rates in liver, kidney, and epididymal fat, and whole body glucose oxidation rates. These findings argued against changes in host glycolysis and lipogenesis as the basis for cachexia in MAC-16 mice. Whole body fatty acid oxidation, however, and oxidation in liver and heart homogenates, were significantly greater in the cachexia-prone MAC-16-bearing mice, consistent with increased expression of uncoupling proteins UCP1 in adipose tissue and UCP 2 and 3 in skeletal muscle20; in contrast, lipid accumulated in the MAC-16 tumor itself. This suggested that increased whole body fatty acid oxidation in MAC-16bearing mice reflected increased fatty acid oxidation in normal host tissues but not in the tumor. Consistent with this apparently low rate of fatty acid oxidation in the tumor, glucose utilization in both MAC-13 and MAC-16 tumors was high, and second only to that in host brain18. This metabolic pattern of the tumors was in marked contrast to that in normal tissues of the tumorbearing host, which utilized glucose at lower rates and oxidized FFA at higher rates than those of non-tumor-bearing controls18. In MAC-16 mice, the peroxisome proliferator bezafibrate accelerated both host weight loss and tumor growth21, the latter associated with increased FFA concentration in plasma and uptake by tumor. Bezafibrate had no effect on body weight or tumor growth in MAC-13 mice. Dietary medium chain triglyceride (MCT) representing 80% of caloric intake ameliorated host weight loss and suppressed tumor growth, associated with elevated plasma ketone bodies and diminished long chain FFA as compared with controls22. Insulin administration also increased host weight, but unlike MCT enhanced tumor growth9. This adverse (tumor growth-enhancing) effect of insulin was reversed by interpreted as suggesting that the anti-tumor effects of

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MCT may have resulted from enhanced ketogenesis; the mechanism of the anti-tumor effect of ketone bodies was not defined. However, an inverse relationship has been demonstrated between hepatoma cell growth rate and the activity of dehydrogenase 10 ; this mitochondrial enzyme either converts to acetoacetate, thereby initiating its subsequent or catalyzes the reverse reaction during ketogenesis. The apparent antiproliferative effect of ketone bodies9,10 may be mechanistically similar to that of short chain fatty acids (see Chapter 6.2.: Butyrate). Thus, in tumor-bearing mice, host utilization of glucose decreased while fatty acid utilization increased, as the host lost weight. The tumor, in contrast, preferentially oxidized glucose and gained weight. As discussed below, these contrasts between cancer and host metabolism also characterize the human cancer cachexia syndrome, and the sequence of changes in metabolism that take place during liver regeneration (see this Chapter 4.5.: Metabolic Interaction between Tumor and Host: Cancer Cachexia; Chapter 8: Liver Regeneration; and Fig. 4.1).

Fig. 4.1. Contrasting patterns of intermediary metabolism in liver regeneration and cancer cachexia. In early liver regeneration and in hepatocytes in the setting of cancer cachexia, increased mitochondrial fatty acid oxidation is associated with increased gluconeogenesis and oxidative stress, and decreased and ATP. In contrast, in late regeneration (proliferative) hepatocytes and in cancer cells, glycolysis, ATP abundance, and fatty acid synthesis are increased, while fatty acid oxidation is suppressed. Under hypoxic conditions, glycolysis in normal and cancer cells is largely anaerobic, survival mechanisms are activated, and suppressed fatty acid oxidation implicit (see Chapter 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation). Abbreviations — mitochondrial inner membrane potential; FA: fatty acid; NA: data not available.


4.3.

OMEGA-3

51

FATTY ACIDS

4.3.1. Metabolic Effects Fatty acids modulate hepatocellular metabolism in vivo, in perfused liver, and in hepatocyte culture23. In general, dietary eicosapentaenoic acid (EPA — 20:5, and docosahexaenoic acid (DHA — 22:6, increase mitochondrial, and dose-dependently increase peroxisomal fatty acid oxidation24-26. They also decrease plasma FFA27, hepatocellular incorporation of fatty acid into triacylglycerol (TG), secretion of TG and of apoB-containing lipoproteins, hepatocellular 28 concentrations of TG23, and lipogenesis24,25. Independent of , these effects result from preferential mitochondrial oxidation of fatty acids, especially EPA29, and increased ketogenesis, reflecting enhanced activity of the rate-determining enzyme CPT-I30 and its diminished sensitivity to inhibition by malonyl CoA24-26. Moreover, incorporation of dietary fatty acids into mitochondrial membrane lipids increases inner membrane proton leakage, thereby uncoupling oxidative 31 phosphorylation. and attenuating . Despite the greater propensity of fatty acids to undergo mitochondrial therefore, their uncoupling effect suppresses intramitochondrial ROS generation32-36. This antioxidant property likely accounts for fatty acid-mediated protection against oxidative stress in human lymphocytes37, reduced expression of inducible NOS, and proinflammatory genes in colon 38 cancer cells , and diminished macrophage ICAM-1 and scavenger receptor expression39 (see also Chapter 6.4.: Salicylates and Other Nonsteroidal Anti-inflammatory Drugs, and Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). Insulin increased the incorporation of fatty acids into perfusate lipoprotein TG by livers from rats fed safflower oil but not from those fed fish oil25, suggesting that mitochondrial fatty acid oxidation and ketogenesis, when utilizing fatty acids, were less subject to suppression by insulin. In isolated rat hepatocytes, mitochondrial oxidation of eicosapentaenoic acid (20:5, greatly 40 exceeded that of arachidonic acid (20:4, . In hamsters, the fatty acid-induced decrease in TG biosynthesis was associated with decreased activity of both phosphatidate phosphohydrolase and diacylglycerol acyltransferase41, i.e., changes which would divert a proportionately greater fraction of esterified fatty acid away from TG biosynthesis and toward that of phospholipids. In primary rat hepatocyte cultures, EPA and DHA inhibited secretion of TG-rich apo-B-containing

52

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lipoproteins and increased intracellular degradation of apoB100 and apoB4842.

Larger quantities of fatty acids in vivo24,26,43 or in cell culture 44 increased hepatocellular TG accumulation and peroxisomal fatty acid oxidation; these effects were more prominent with DHA than EPA27,29,45,46. Decreases in plasma FFA were also observed47. Conversely, when relatively small amounts of EPA were fed, plasma TG decreased27,41,46; moreover, plasma TG levels were inversely related to CPT-I activity and mitochondrial which increased without significant change in 41,46. peroxisomal fatty acid

4.3.2. Growth Effects Fatty acids inhibit tumorigenesis and the growth of normal and transformed epithelial cells, compared with and other long chain fatty acids, in human subjects, as well as in human cell lines in culture and after implantation into nude mice38,48-54. For example, feeding of fish oil rich) to human volunteers diminished BrdU labelling, ornithine decarboxylase activity, and prostaglandin generation in normal rectal mucosa as compared with corn oil rich) or placebo feeding55. Similar effects were observed on the labelling index of sporadic colorectal adenomas in subjects fed fish oil56, and on colorectal epithelial cell proliferation in patients with a history of colon polyp or stage 1 or 2 colon carcinoma fed fatty acids57. Phytohemagglutinin-induced monocyte proliferation and cytokine production ex vivo were inhibited in cells from normal subjects fed fatty acid supplements58. Some studies of cells in culture (especially non-epithelial) have produced negative or equivocal results59-61. Animal cell and tumor experiments have generated results similar to those obtained in the aforementioned human studies62,63. Thus, in the in vivo perfused 7288CTC Morris hepatomas in Buffalo rats64, linoleic, arachidonic, and other fatty acids stimulated incorporation, whereas the fatty acids EPA and (18:3, competitively inhibited both linoleic acid uptake by the tumor and its stimulation of incorporation. EPA blocked tumor growth and host cachexia in MAC-16 colon carcinoma-bearing mice65,66, inhibited stimulation of adipose tissue lipolysis by a tumor-derived lipidmobilizing factor67,68, and suppressed other adverse effects of cancer and cancer-derived cachexia-promoting factors, in vivo and in vitro (see this Chapter 4.5.: Metabolic Interaction between Tumor and Host: Cancer Cachexia). EPA also decreased cell proliferation in normal rat colon69 and liver70, in transplanted Morris hepatomas70, and in Solt-Farber


53

carcinogen-induced liver cancers71. A pre-operative fish oil diet decreased post-hepatectomy mitotic activity (BrdU incorporation) in hepatic acinar zones 1 and 2 by 60%43. Fatty acids inhibited transgenic murine colon carcinogenesis, and reversed suppression of in vivo and in vitro72,73. Thus, the available data indicate that fatty acids, especially EPA, increase mitochondrial fatty acid oxidation, decrease TG biosynthesis and VLDL secretion, and increase the intracellular degradation of apoB, as compared with other long chain fatty acids. The uncoupling action of fatty acids in feeding experiments likely reflects increased inner mitchondrial membrane proton permeability resulting from their incorporation into membrane lipids31; the antioxidant effects of this and other uncoupling agents are discussed below (see Chapter 6.4.: Salicylates and Other Nonsteroidal Anti-inflammatory Drugs; and Chapter 7.3.: Origins of Intramitochondrial Oxidative Stress). However, the fact that metabolic effects are evident in short term in vitro studies of preparations from subjects fed control diets and in Hep G2 cells26,74, suggests that their distinctive metabolic characteristics also reflect properties of the unesterified fatty acids, i.e., independent of prior incorporation into membrane phospholipids. The fatty acids, especially EPA, also inhibit the growth of normal and transformed epithelial cells, in contrast to the fatty acids, especially linoleic, which promote growth. These effects are clearly associated with changes in cellular fatty acid metabolism. As EPA inhibits arachidonate utilization by prostaglandin H synthase (cyclooxygenase — COX1, COX2) and lipoxygenase75, differences in eicosanoid biosynthesis may contribute to these changes76,77.

4.4.

FATTY ACIDS, MODULATION OF CELL GROWTH, AND THE AMP-ACTIVATED PROTEIN KINASE

Consistent with the pattern of augmented glucose utilization and lipogenesis in tumor cells discussed above, cell lines derived from human breast, colorectal, and prostate cancers express fatty acid synthase and actively synthesize fatty acids de novo; inhibition of fatty acid synthase arrested tumor cell growth and induced apoptosis78-80. A similarly widespread expression and relationship to tumorigenesis has been described for C-FABP, the cutaneous isoform of FABP81. It is important to note that fatty acid synthesis requires increased acetyl CoA

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Chapter 4

Figure 4.2. Metabolism of fatty acids in rodent hepatocyte. Both mitochondrial and extramitochondrial FA oxidation pathways are transcriptionally activated by Direct inhibition of carnitine palmitoyl transferase-I by malonyl CoA, cytokeratins 8 and 18, or peroxisome proliferators favors diversion of FA toward esterification and extramitochondrial oxidation. Active AMPK abrogates both malonyl CoA-dependent and malonyl CoA-independent (cytokeratin-mediated) inhibition of CPT-I, thereby promoting mitochondrial fatty acid oxidation. Activation of the transcriptional regulator may reflect cellular abundance of FA. Abbreviations — AMPK: adenosine monophosphate-activated protein kinase; CPT-I: carnitine palmitoyltransferase-I; FA: fatty acid; FA-CoA: fatty acid-coenzyme A thioester; PL: glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

carboxylase activity and malonyl CoA abundance. As a result, mitochondrial fatty acid oxidation is suppressed, reflecting inhibition of the rate-determining enzyme CPT-I by malonyl CoA30, as well as by malonyl CoA-independent mechanisms82-84 (Figs. 4.1 and 4.2). wg/Wntlike activity also increases glucose utilization85 and de novo lipogenesis; the latter has been demonstrated directly86, and is implied by cell proliferation (which requires fatty acids) in the absence of exogenous lipid87,88. In addition, and in contrast to the antilipolytic effect of insulin,


55

90,91 HGF89 and augment serum FFA concentrations after partial hepatectomy (also, see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin). This reflects increased adipocyte lipolysis, thereby providing to the proliferating hepatocytes additional fatty acids, including the essential fatty acids which generate ROS and are utilized in membrane and eicosanoid biosynthesis (see this Chapter 4.2.:Omega-6 Fatty Acids). Interaction of acetyl CoA carboxylase with wild type but not mutant BRCA192 suggests that tumor suppression by BRCA1 may reflect in part control of lipogenesis. In addition to fatty acids, cell growth also depends on de novo biosynthesis of mevalonate, isoprenoids including sterols93-95, and phospholipid96. Although growth-related post-translational regulation of the fatty acid and isoprenoid/sterol biosynthetic pathways through the AMPK mechanism (see this Chapter 4.4.1.: AMP-Activated Protein Kinase) is well established97, coordinate transcriptional regulation of their respective genes, and those related to glycolysis, would also be advantageous, e.g., as mediated by the sterol regulatory element binding proteins (SREBPs) and by the oxysterol-activated liver X receptor (LXR)98-100. Thus, the LXR-SREBPlc lipogenesis pathway is activated by insulin99,101-104 and the key PI3K/Akt/PKB-mediated wg/Wnt-like signaling pathway80,105, and is inhibited by polyunsaturated fatty acids106,107 and by the HIV-1 protease inhibitor indinavir108. Moreover, HGF-induced expression of the low density lipoprotein receptor gene in HepG2 cells109 would serve to enhance delivery to the cell of both fatty acids and sterols; this suggests that cell proliferation induced by HGF-initiated wg/Wnt-like signaling110,111 is linked to the SRE/SREBP mechanism, which plays a major role in cellular regulation of both isoprenoid (including sterols) and fatty acid biosynthesis. Direct evidence for such linkages in tumor cells has been provided recently80,105,112,113. Moreover, recent evidence indicates that LXR, while activating GLUT4 expression and thereby supporting glycolysis and insulin responsiveness, suppresses the opposing (fatty acid oxidation-energized) pathway of gluconeogenesis114. Furthermore, LXR not only promotes lipogenesis via its interaction with SREBP1c, but also suppresses increases in fatty acid oxidation115; (also, see Chapter 5.2.1.: Together, these observations provide important support for the concept of coordinate linkage of glycolytic and lipogenic patterns of carbohydrate, fatty acid, and isoprenoid (sterol) regulation to cell proliferation. Additionally, a potentially critical role of these linkages in the atherosclerosissuppressing effect of statin hypocholesterolemic agents is discussed below (Chapter 7.4.: Consequences of Intramitochondrial Oxidative Stress: Atherosclerosis and Beyond).

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Figure 4.3. Glycolysis- and citric acid cycle-fueled biosynthesis of ATP and fatty acid; suppressed fatty acid oxidation and gluconeogenesis. This figure represents the predominant pattern of substrate utilization in proliferating cells; enclosed areas represent mitochondria. Arrow thickness approximates relative pathway activity; dotted lines: pathways that are suppressed during cell proliferation. Fatty acids also may be derived from plasma. Abbreviations — CPT-I: carnitine palmitoyltransferase-I (shown as inhibited by malonyl CoA); EMFO: extramitochondrial fatty acid oxidation in peroxisomes, and in endoplasmic reticulum); F26BP: fructose-2,6bisphosphate; PEPCK: phosphoenolpyruvate carboxykinase; PL: glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

Taken together, available evidence strongly suggests that non-fatty acid fuels, principally glucose, are utilized in cells undergoing mitosis (Figs. 4.1 and 4.3). This supports the obligatory increases in both energy generation and biosynthesis of fatty acids and sterols that characterize cell proliferation. To the extent that fatty acids and sterols can be obtained from plasma, consumption of ATP and diversion of carbohydrate-derived substrates away from ATP generation and toward lipogenesis are minimized. It is of interest in this context that while the mechanism


57

underlying the mitogenic and survival effects of lysophosphatidic acid (LPA) remains incompletely defined116, these effects are augmented by insulin117 and mediated by Akt/PKB118.

In any case, however, exogenous essential fatty acids, which generate growth-promoting ROS, eicosanoids, and membrane lipids, are required. As a corollary, fatty acids that most effectively inhibit cell growth, i.e., medium chain, and short chain (e.g., butyrate; see Chapter 6.2.) are those that most readily undergo mitochondrial oxidation, by attenuating or circumventing (medium and short chain) a critical control point in the process, i.e., the physiological inhibition of CPT-I by malonyl CoA-dependent and malonyl CoAindependent mechanisms. The evidence that increased mitochondrial fatty acid oxidation is associated with growth inhibition implies that growth stimulation by essential fatty acids reflects their utilization not as substrate for ATP generation but rather for the biogenesis of ROS, membranes, and eicosanoids and the synthesis of glycerophosphatides important in signal transduction119.

4.4.1. AMP-Activated Protein Kinase A direct link among fatty acid and isoprenoid (e.g., sterol) biosynthesis, mitochondrial fatty acid oxidation, energy metabolism, and the regulation of cell growth is provided by the AMP-activated protein kinase (AMPK)97. AMPK is the mammalian homolog of the SNF1 gene product in yeast, a key determinant of growth arrest in response to glucose starvation. In mammals, AMPK is activated by increased [AMP] or [AMP]/[ATP]; in yeast, SNF1 is activated in association with, and possibly by, similar changes in adenine nucleotides97,120. The active enzyme phosphorylates and inactivates the rate-determining steps in fatty acid and sterol biosynthesis, i.e., acetyl CoA carboxylase and CoA reductase (HMG CoA reductase), respectively, and in yeast arrests growth. Recent evidence indicates that AMPK plays a critical and central role in both malonyl CoA-dependent and malonyl CoA-independent regulation of CPT-I and, thus, mitochondrial fatty acid oxidation (Fig. 4.2). Malonyl CoA-dependent regulation of CPT-I reflects the decreased cellular abundance of malonyl CoA that results from: 1) decreased malonyl CoA synthesis as the result of an inhibitory AMPK phosphorylation of acetyl CoA carboxylase97; and 2) increased malonyl CoA degradation (in liver, adipose, and muscle) via AMPK-mediated activation of malonyl CoA decarboxylase121. The resulting drop in cellular abundance of malonyl CoA derepresses CPT-I activity and thus

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augments mitochondrial fatty acid oxidation97. Malonyl CoAindependent regulation of CPT-I reflects AMPK-mediated alteration of the interation between CPT-I and intermediate filaments, specifically cytokeratins 8 and 1882,83. Active AMPK phosphorylates these cytoskeletal components, disrupting their mitochondrial connection and releasing CPT-I activity from the constraint that they otherwise impose. In addition, fatty acids of varied chain length lead to hepatocyte AMPK activation122. Thus, in a multifaceted stress-induced and antiproliferative cascade, fatty acids and/or altered adenine nucleotide balance activate AMPK; in turn, active AMPK augments mitochondrial fatty acid oxidation by suppressing both malonyl CoA-dependent and malonyl CoA-independent (cytokeratin-mediated) inhibition of CPT-I, and (glucose-dependent) lipogenesis. The resulting promotion of fatty acid oxidation by AMPK is additionally fostered in contracting cardiac myocytes through its activation of fatty acid translocase (FAT)/CD36mediated fatty acid uptake123. It appears likely that these consequences of altered abundance of adenine nucleotides and fatty acids would have broad phylogenetic significance in the linkage of cell metabolism to cell proliferation, growth arrest, and apoptosis97,120,124 (also, see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer). Moreover, AMPK mediation of mitochondrial fatty oxidation may be activated by leptin 125-127 (also, see Chapter 7.3.: Origins of Intramitochondrial Oxidative Stress; and, 7.4.: Consequences of Oxidative Stress). This action may contribute importantly to leptin’s systemic effects, including hypothalamic regulation of appetite and feeding behavior (see Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior). Importantly, AMPK activation may be antagonized by insulin, in an adenine nucleotide-independent and PI3K-dependent manner128, and similarly is subject to modulation by other agents (see Chapter 5.3.4.: Apoptosis and the Cell Cycle).

4.4.2. Stoichiometric Considerations It is useful to consider the stoichiometry of ATP generation in relation to utilization of glucose and fatty acids, and the influence of ketogenesis. During complete citric acid cycle-mediated oxidation of glucose to and the theoretical maximum net yield of ATP molecules per molecule of consumed can be calculated to be 5.0129. The corresponding values for complete oxidation of glutamine as a respiratory substrate, reflecting its conversion to glutamate and entry into the citric acid cycle as 2-oxo-glutarate, can be calculated as


59

approximately 20-23 molecules of ATP per molecule of glutamine, or 5.0 to 5.6 per

molecule129.

In contrast, during complete oxidation of palmitate the theoretical maximum is 4.6 ATP molecules per molecule of consumed, but the yield may be decreased in ketogenic cells by conversion of palmitate to ketone bodies, i.e., to 3.6 (complete conversion to acetoacetate) or 3.9 (complete conversion to and even further by long chain fatty acid-induced uncoupling of oxidative phosphorylation130-134. Viewed from a different perspective, the theoretical potential net yield of 106 ATP molecules derived per molecule of palmitate oxidized (complete oxidation, no ketogenesis, no uncoupling), would be reduced by more than 75% by complete conversion to ketone bodies, i.e., to 26 (all acetoacetate), or to 18 (all and further diminished by the uncoupling effect. Moreover, control of CPT-I by malonyl CoA can be overridden by high fatty acid abundance84,135-138 and/or suppressed (in liver, adipose tissue, and skeletal muscle) by fatty acid- and/or AMPKinduced malonyl CoA decarboxylase expression121,139. The resulting increase in mitochondrial fatty acid oxidation may augment “leakage” of electrons from the electron transport chain and generation of thus further diminishing the efficiency of fatty acids as fuel for oxidative phosphorylation. This unsuitability of fatty acid oxidation for support of cell proliferation may account in large part for the recent observation that cannabinoid receptor activation, which promotes astrocyte fatty acid oxidation and ketogenesis (see Chapter 13.4.: Fatty Acids, Antioxidant Defense, HIV Dementia, and Cell Proliferation), inhibits growth of gliomas and skin tumors, and angiogenesis140. It is clear that mitochondrial oxidation of fatty acids, with or without ketogenesis, is energetically less efficient than is fueling of the citric acid cycle by either glycolysis or glutamine. For this reason, and because of the fatty acid requirement for membrane biogenesis, fatty acid oxidation as the basis for support of cell cycle-related energetics would be less favored in terms of survival. Significantly, enzymes controlling both the entry of glucose into the glycolytic pathway141-143, and the entry of fatty acid into the mitochondrial pathway144,145, are localized to the contact sites between inner and outer mitochondrial membranes (Fig. 4.4). There, the VDAC complex links cytosol and mitochondrial matrix, permitting adenine nucleotide translocase-mediated transfer of ATP and ADP between these compartments. Fatty acids in excess perturb this critical regulatory locus by causing: 1) desorption of hexokinase from the VDAC complex, thus further limiting utilization of glucose and favoring both opening of the permeability transition pore141 and Bax-

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Figure 4.4. Mitochondrial voltage-dependent anion channel (VDAC) components and relationship to electron transport chain; schematic representation. VDAC complexmediated juxtaposition of hexokinase II and the adenine nucleotide translocase in proliferating cells optimizes energetics of glucose phosphorylation to glucose-6phosphate and entry into the glycolytic pathway, leading to pyruvate formation. Detailed structural relationships remain incompletely defined (see Chapter 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation). Increased mitochondrial fatty acid oxidation inhibits glucose uptake, hexokinase, and pyruvate dehydrogenase, and predisposes to electron “leakage” from the electron transport chain; the latter results in one-electron reduction of oxygen and formation of and derivative ROS. Excess cellular fatty acids may disrupt hexokinase II, VDAC, and the contact sites. Unlike anti-apoptotic Bcl-2 family members, pro-apoptotic members may associate with, and perturb, the mitochondrial membrane after activation of the apoptotic cascade. Abbreviations — CYT C: cytochrome c; inner mitochondrial membrane potential; superoxide anion; UBIQ: ubiquinone; ROS: reactive oxygen species; VDAC: voltage-dependent anion channel. With permission (see Fig. 3.4.)

induced release of cytochrome c146,147; and 2) disruption of the contact

sites themselves148. Evolutionary pressures for survival are likely to have favored not only maximal efficiency in energy generation, but also maximal precision in its control. The latter may be provided most effectively by an otherwise seemingly contradictory activation of opposing pathways. For example, whereas fatty acids inhibit transgenic signaling73, fatty acids activate certain isoforms of PKC149-151. Similarly, expression of a growth inhibitor, is increased in hepatocytes at various stages during hepatic regeneration152,153, although receptor signaling is 154 suppressed . expression is also increased in many tumor cells


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despite their accelerated growth155,156. In addition, increases in cytosolic mediated by EGF and other growth factors lead to corresponding increases in mitochondrial thereby enhancing the activity of several citric acid cycle-related mitochondrial dehydrogenases (pyruvate, isocitrate, and 2-oxo-glutarate dehydrogenases). This favors glycolytic fueling of oxidative phosphorylation157, representing an important “effector arm” of signaling.

4.5. METABOLIC INTERACTION BETWEEN TUMOR AND HOST: CANCER CACHEXIA Cancer cachexia — the most frequent and obvious indication of the impact of tumor on host, and of their metabolic interaction — has been extensively reviewed158-161. Variable in its manifestations, it is generally characterized by progressive loss of fat and muscle, is largely unrelated to anorexia and unresponsive to nutritional supplementation, and is a major determinant of poor prognosis and mortality. In addition to its clinical significance, this syndrome provides insight into the metabolism of proliferating cells, in this case those comprising the malignant neoplasm. Hypotheses concerning the pathogenesis of this syndrome include increased resting energy expenditure, a starvation-like response, a parasite-like effect of the tumor on host nutrition, and an effect of proinflammatory cytokines, especially tumor necrosis However, various inconsistencies exclude each of these as a sole or dominant basis for the syndrome.. Recently, two cancer cachexiaassociated factors have been demonstrated in the urine of patients with cancer cachexia and in the tumor and urine of cachectic MAC-16 tumorbearing mice 67,68. They are not demonstrable in normal or tumor-bearing but non-cachectic patients or mice, nor in wasting related to trauma or sepsis. One of these, designated lipid-mobilizing factor (LMF), has an apparent molecular mass of 43 kDa67, has been further characterized as a 68 , and has been shown to be structurally related to the class I major histocompatibility heavy chain162. When administered to normal mice, LMF rapidly induced adipose tissue lipolysis, and loss of muscle mass and body weight; in vitro it degraded skeletal muscle protein. The second cachexia-associated factor, designated proteolysisinducing factor (PIF)163, has an apparent molecular mass of 24 kDa. PIF effects fatty acid- (EPA)-suppressible apoptosis in murine myotubes 164 in vitro . In vivo, fatty acids in high concentrations suppress the catabolic effects of both factors in mice65,66; their possible usefulness in

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treatment of weight-losing pancreatic cancer patients is under investigation165. LMF and PIF may be important causes of cancer cachexia, and may provide further insight into its pathogenesis. Increased expression of uncoupling proteins may also contribute20. Metabolic disturbances in cachectic cancer patients often are similar to those associated with diabetes mellitus. Thus, they may resemble either the insulin resistance and increased insulin secretory response typical of type II, or the diminished fasting insulin levels, insulin sensitivity, and insulin response typical of type I. In either case, lipolysis and FFA mobilization are increased166-170. Impaired synthesis and/or accelerated degradation of muscle protein leads to loss of lean body mass160-171. Utilization of glucose by the tumor is often increased at a time when the host is glucose intolerant, the latter reflecting impaired glucose utilization in skeletal muscle and increased hepatic glucose production166-169-172. In 173 type II diabetes mellitus, increased adipocyte release of and FFA leads to impaired insulin regulation of both peripheral glucose utilization and hepatic glucose production 30,174-181, and it is likely that these factors would contribute to the similar insulin resistance that characterizes cancer cachexia. In contrast to this more general pattern, some patients with hepatocellular carcinoma and severe hypoglycemia may occupy a metabolically intermediate position between fasting and maximal insulinization, reflecting circulation of variant insulin-like growth factorII peptides182,183. At a cellular level, the metabolic interaction between tumor and host is reflected to a large extent by competition between substrates for mitochondrial oxidation, i.e., fatty acids vs. pyruvate, the latter generated via glycolysis or derived from lactate or deamination of alanine. Mitochondrial oxidation of pyruvate depends on pyruvate dehydrogenase184, activity of which is inhibited by fatty acid oxidationinduced increases in the ratios acetyl CoA/CoA, NADH/NAD, and ATP/ADP in the mitochondrial matrix. Increased cellular lipid and mitochondrial fatty acid oxidation in host tissues184,185 inhibit glucose uptake, glycolysis, and pyruvate dehydrogenase, contributing to insulin resistance30,174-181,186,187, and driving increased gluconeogenesis in liver and kidney (Figs. 4.1 and 4.5). Further evidence of inefficiency in host energetics is provided by the increased expression of host tissue uncoupling proteins in the MAC16 mouse colon cancer model20. Significantly, hepatocellular ATP is diminished in cancer cachexia188,189. This perhaps unexpected finding also characterizes another systemic catabolic crisis in which increased FFA mobilization and a fatty acid oxidation-fueled gluconeogenic response are activated, namely, the


63

Figure 4.5. Fatty acid oxidation and gluconeogenesis; suppressed glycolysis and fatty acid synthesis. Enclosed areas represent mitochondria. This pattern of substrate utilization is antiproliferative. Arrow thickness approximates relative pathway activity; dotted line: pathway suppressed during growth arrest. Numbers in parentheses indicate approximate yields of ATP from complete oxidation of palmitate (~100), compared (in a ketogenic cell, e.g., hepatocyte) with complete diversion to ketogenesis (~20). Fatty acids activate transcriptionally upregulating both mitochondrial and extramitochondrial fatty acid oxidation. When present in excess, fatty acids may override malonyl CoA inhibition of CPT-I. Abbreviations — ATP: adenosine triphosphate; CPTI: carnitine palmitoyltransferase-I; EMFO: extramitochondrial fatty acid oxidation in peroxisomes, and in endoplasmic reticulum); F26BP: fructose-2,6-bisphosphate; PEPCK: phosphoenolpyruvate carboxykinase; PL: phospholipids, i.e., glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

several hour-long period that immediately follows 70% hepatectomy (see Fig. 4.1 and Chapter 8: Liver Regeneration). The substantial differences in nutrient utilization between host liver and tumor (Fig. 4.1) suggest that tumor growth requirements are well served by cancer cachexia-enhanced lipolysis in adipose tissue and release of amino acids from muscle. Thus, the resulting increases in plasma [FFA] and contribute to insulin resistance in host tissues

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by inhibiting systemic glucose utilization and promoting hepatic and renal gluconeogenesis. This increases the availability to the tumor of both glucose (for energy and the biosynthesis of fatty acids and isoprenoids), and fatty acids (for membrane and eicosanoid biogenesis). Increased amino acid release into plasma would contribute to tumor cell protein biosynthesis and in some instances energy metabolism (glutamine), as well as to host gluconeogenesis (alanine and glutamine). Tumor cell proliferation and its relative growth advantage, therefore, are supported at the expense to the host of both substrate and energy, the latter “wastefully” expended both by its consumption in support of gluconeogenesis and by the relative inefficiency of augmented fatty aciddependent mitochondrial energy generation.

4.6.

REFERENCES

1. W. Kidwell, M. Monaco, M. Wicha, and G. Smith, Unsaturated fatty acid requirements for growth and survival of a rat mammary tumor cell line, Cancer Res 38:4091-4100 (1978). 2. M. Wicha, L. Liotta, and W. Kidwell, Effects of free fatty acids on the growth of normal and neoplastic rat mammary epithelial cells, Cancer Res 39:426-435 (1979). 3. S. Abraham, and L. Hillyard, Effect of dietary 18-carbon fatty acids on growth of transplantable mammary adenocarcinomas in mice., J Natl Cancer Inst 71:601-605 (1983). 4. C. Maziere, M.-A. Conte, J. Degonville, D. Ali, and J.-C. Maziere, Cellular enrichment with polyunsaturated fatty acids induces an oxidative stress and activates the transcription factors AP1 and NF-kappaB, Biochem Biophys Res Comm 265:116122 (1999). 5. I. Rusyn, C. Bradham, L. Cohn, et al., Corn oil rapidly activates nuclear factor-kappaB in hepatic Kupffer cells by oxidant-dependent mechanisms, Carcinogenesis 20:20952100 (1999). 6. I. Rusyn, H. Tsukamoto, and R. Thurman, WY-14 643 rapidly activates nuclear factor kappaB in Kupffer cells before hepatocytes, Carcinogenesis 19:1217-1222 (1998). 7. L. Sauer, and R. Dauchy, Blood nutrient relationships during the onset of an acute fast nt concentrations and tumor growth in vivo in rats:, Cancer Res 47:1065-1068 (1987A). 8. L. Sauer, and R. Dauchy, Stimulation of tumor growth in adult rats in vivo during acute streptozotocin-induced diabetes, Cancer Res 47:1756-1761 (1987B). 9. S. Beck, and M. Tisdale, Effect of insulin on weight loss and tumour growth in a cachexia model., Br J Cancer 59:677-681 (1989). 10. W. Zhang, S. Churchill, R. Lindahl, and P. Churchill, Regulation of D-betahydroxybutyrate dehydrogenase in rat hepatoma cell lines, Cancer Res 49:2433-2437 (1989). 11. L. Sauer, and R. Dauchy, Identification of linoleic and arachidonic acids as the factors in hyperlipemic blood that increase [3H]thymidine incorporation in hepatoma 7288CTC perfused in situ, Cancer Res 48:3106-3111 (1988). 12. L. Sauer, and R. Dauchy, Uptake of plasma lipids by tissue-isolated hepatomas 7288CTC and 7777 in vivo, Br J Cancer 66:290-296 (1992A).


65

13. R. Daoust, The passage of G2 hepatocytes into mitosis during fasting., Chem Biol Interact 62:99-103 (1987). 14. R. Daoust, Stimulation of G2-arrested rat liver and pancreas cells by fasting., Gastroenterology 97:219-221 (1989). 15. J. Bassuk, P. Tsichlis, and S. Sorof, Liver fatty acid binding protein is the mitosisassociated polypeptide target of a carcinogen in rat hepatocytes., Proc Natl Acad Sci USA 84:7547-7551 (1987). 16. S. Sorof, Modulation of mitogenesis by liver fatty acid binding protein, Cancer Metast Rev 13:317-336 (1994). 17. H. Mulligan, and M. Tisdale, Lipogenesis in tumour and host tissues in mice bearing colonic adenocarcinomas, Br J Cancer 63:719-722 (1991A). 18. H. Mulligan, and M. Tisdale, Metabolic substrate utilization by tumour and host tissues in cancer cachexia, Biochem J 277:321-326 (1991B). 19. H. Mulligan, S. Beck, and M. Tisdale, Lipid metabolism in cancer cachexia, Br J Cancer 66:57-61 (1992). 20. C. Bing, M. Brown, P. King, P. Collins, M. J. Tisdale, and G. Williams, Increased gene expression of brown fat uncoupling protein (UCP)1 and skeletal muscle UCP2 and UCP3 in MAC16-induced cancer cachexia, Cancer Res 60:2405-10 (2000). 21. H. Mulligan, and M. Tisdale, Effect of the lipid-lowering agent bezafibrate on tumour growth rate in vivo, Br J Cancer 64:1035-1038 (1991C). 22. M. Tisdale, R. Brennan, and K. Fearon, Reduction of weight loss and tumour size in a cachexia model by a high fat diet, Br J Cancer 56:39-43 (1987). 23. A. Moir, B. Park, and V. Zammit, Quantification in vivo of the effects of different types of dietary fat on the loci of control involved in hepatic triacylglycerol secretion, Biochem J 308:537-542 (1995). 24. S. Wong, P. Nestel, R. Trimble, G. Storer, R. Illman, and D. Topping, The adaptive effects of dietary fish and safflower oil on lipid and lipoprotein metabolism in perfused rat liver, Biochim Biophys Acta 792:103-109 (1984). 25. D. Topping, R. Trimble, and G. Storer, Failure of insulin to stimulate lipogenesis and triacylglycerol secretion in perfused livers from rats adapted to dietary fish oil, Biochim Biophys Acta 927:423-428 (1987). 26. T. Ide, M. Murata, and M. Sugano, Stimulation of the activities of hepatic fatty acid oxidation enzymes by dietary fat rich in a-linolenic acid in rats, J Lipid Res 37:448463 (1996). 27. A. Demoz, D. Asiedu, O. Lie, and R. Berge, Modulation of plasma and hepatic oxidative status and changes in plasma lipid profile by n-3 (EPA and DHA), n-6 (corn oil) and a 3-thia fatty acid in rats., Biochim Biophys Acta 1199:238-244 (1994). 28. J. Dallongeville, E. Baugé, A. Tailleux, et al., Peroxisome proliferator-activated receptor alpha is not rate-limiting for the lipoprotein-lowering action of fish oil., J Biol Chem 16 276:4634-4639 (2001). 29. L. Madsen, L. Frøyland, E. Dyrøy, K. Helland, and R. Berge, Docosahexaenoic and eicosapentaenoic acids are differently metabolized in rat liver during mitochondria and peroxisome proliferation, J Lipid Res 39:583-593 (1998). 30. J. McGarry, Banting lecture, 2001: dysregulation of fatty acid metabolism in the etiology of type 2 diabetes, Diabetes 51:7-18 (2002). 31. D. Rolfe, A. Hulbert, and M. Brand, Characteristics of mitochondrial proton leak and control of oxidative phosphorylation in the major oxygen-consuming tissues of the rat, Biochim Biophys Acta 1188:405-416 (1994).

66

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32. I. Reynolds, and T. Hastings, Glutamate induces the production of reactive oxygen species in cultured forebrain neurons following NMDA receptor activation, J Neurosci 15:3318-3327 (1995). 33. V. Skulachev, Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants, Quart Rev Biophys 29:169-202 (1996). 34. A. Nègre-Salvayre, C. Hirtz, G. Carrera, et al., A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation, FASEB J 11:809-815 (1997). 35. A. Stout, H. Raphael, B. Kanterewicz, E. Klann, and I. Reynolds, Glutamate-induced neuron death requires mitochondrial calcium uptake, Nature Neurosci 1:366-373 (1998). 36. S. Korshunov, O. Kokina, E. Ruuge, V. Skulachev, and A. Starkov, Fatty acids as natural uncouplers preventing generation of and by mitochondria in the resting state, FEBS Letts 435:215-218 (1998). 37. S. Bechoua, M. Dubois, Z. Dominguez, et al., Protective effect of docosahexaenoic acid against hydrogen peroxide-induced oxidative stress in human lymphocytes., Biochem Pharmacol 57:1021-1030 (1999). 38. B. A. Narayanan, N. K. Narayanan, B. Simi, and B. S. Reddy, Modulation of inducible nitric oxide synthase and related proinflammatory genes by the omega-3 fatty acid docosahexaenoic acid in human colon cancer cells, Cancer Res 63:972-9 (2003). 39. E. Miles, F. Wallace, and P. Calder, Dietary fish oil reduces intercellular adhesion molecule 1 and scavenger receptor expression on murine macrophages, Athersclerosis 152:43-50 (2000). 40. T. N. Tran, and B. O. Christophersen, Partitioning of polyunsaturated fatty acid oxidation between mitochondria and peroxisomes in isolated rat hepatocytes studied by HPLC separation of oxidation products, Biochim Biophys Acta 1583:195-204 (2002). 41. M. Surette, J. Whelan, K. Broughton, and J. Kinsella, Evidence for mechanisms of the hypotriglyceridemic effect of n-3 polyunsaturated fatty acids, Biochim Biophys Acta 1126:199-205 (1992). 42. H. Wang, X. Chen, and E. Fisher, N-3 fatty acids stimulate intracellular degradation of apoprotein B in rat hepatocytes, J Clin Invest 91:1380-1389 (1993). 43. C. Van Noorden, Effects of n-3 and n-6 polyunsaturated fatty acid-enriched diets on lipid metabolism in periportal and pericentral compartments of female rat liver lobules and the consequences for cell proliferation after partial hepatectomy, J Lipid Res 36:1708-1720 (1995). 44. P. Tollet, M. Stromstedt, L. Froyland, R. Berge, and J. Gustafsson, Pretranslational regulation lf cytochrome P4504A1 by free fatty acids in primary cultures of rat hepatocytes, J Lipid Res 35:248-254 (1994). 45. N. Willumsen, S. Hexeberg, J. Skorve, M. Lundquist, and R. Berge, Docosahexaenoic acid shows no triglyceride-lowering effects but increases the peroxisomal fatty acid oxidation in liver of rats, J Lipid Res 34:13-22 (1993). 46. L. Frøyland, L. Madsen, H. Vaagenes, et al., Mitochondrion is the principal target for nutritional and pharmacological control of triglyceride metabolism., J Lipid Res 38:1851-1858 (1997). 47. A. Rustan, B. Hustvedt, and C. Drevon, Postprandial decrease in plasma unesterified fatty acids during n-3 fatty acid feeding is not caused by accumulation of fatty acids in adipose tissue, Biochim Biophys Acta 1390:245-257 (1998).


67

48. A. Dannenberg, and M. Reidenberg, Dietary fatty acids are also drugs., Clin Pharmacol Ther 55: 5-9 (1994). 49. D. Rose, and J. Connolly, Effects of fatty acids and inhibitors of eicosanoid synthesis on the growth of a human breast cancer cell line in culture, Cancer Res 50:7139-7144 (1990). 50. D. Rose, J. Connolly, and X. Liu, Effects of linoleic acid and gamma-linolenic acid on the growth and metastasis of a human breast cancer cell line in nude mice and on its growth and invasive capacity in vitro, Nutr Cancer 24:33-45 (1995). 51. V. Chajes, W. Sattler, A. Stranzl, and G. Kostner, Influence of n-3 fatty acids on the growth of human breast cancer cells in vitro: relationship to peroxides and vitaminE., Breast Cancer Res Treat 34:199-212 (1995). 52. L. Maehle, E. Eilertsen, S. Mollerup, S. Schonberg, H. Krokan, and A. Haugen, Effects of n-3 fatty acids during neoplastic progression and comparison of in vitro and in vivo sensitivity of two human tumour cell lines, Br J Cancer 71:691-696 (1995). 53. P. Lai, J. Ross, K. Fearon, J. Anderson, and D. Carter, Cell cycle arrest and indection of apoptosis in pancreatic cancer cells exposed to eicosapentaenoic acid in vitro, Br J Cancer 74:1375-1383 (1996). 54. H. Finstad, M. Myhrstad, H. Heimli, et al., Multiplication and death-type of leukemia cell lines exposed to very long-chain polyunsaturated fatty acids., Leukemia 12:921929 (1998). 55. H. Bartram, A. Gostner, W. Scheppach, et al., Effects of fish oil on rectal cell proliferation, mucosal fatty acids, and prostaglandin E2 release in healthy subjects., Gastroenterology 105:1317-1322 (1993). 56. M. Anti, F. Armelao, G. Marra, et al., Effects of different doses of fish oil on rectal cell proliferation in patients with sporadic colonic adenomas., Gastroenterology 107:1709-1718 (1994). 57. Y.-C. Huang, J. Jessup, R. Forse, et al., n-3 Fatty acids decrease colonic epithelial cell proliferation in high-risk bowel mucosa, Lipids 31:S313-S317 (1996). 58. S. Endres, S. Meydani, R. Ghorbani, R. Schindler, and C. Dinarello, Dietary supplementation with n-3 fatty acids suppresses interleukin-2 production and mononuclear cell proliferation., J Leukocyte Biol 54:599-603 (1993). 59. G. Johanning, and T. Lin, Unsaturated fatty acid effects on human breast cancer cell adhesion, Nutr Cancer 24:57-66 (1995). 60. J. Gore, P. Besson, C. Hoinard, and P. Bougnoux, Na+-H+ antiporter activity in relation to membrane fatty acid composition and cell proliferation, Am J Physiol 266:C110-C120 (1994). 61. H. Obermeier, N. Hrboticky, and A. Sellmayer, Differential effects of polyunsaturated fatty acids on cell growth and differentiation of premonocytic U937 cells, Biochim Biophys Acta 1266:179-185 (1995). 62. H. Gabor, L. Hillyard, and S. Abraham, Effect of dietary fat on growth kinetics of transplantable mammary carcinoma in BALB/c mice., J Natl Cancer Inst 74:12991305 (1985). 63. H. Gabor, and S. Abraham, Effect of dietary menhaden oil on tumour cell loss and the accumulation of mass of a transplantable mammary adenocarcinoma in BALB/c mice., J Natl Cancer Inst 76:1223-1229 (1986). 64. L. Sauer, and R. Dauchy, The effect of omega-6 and omega-3 fatty acids on 3Hthymidine incorporation in hepatoma 7288CTC perfused in situ, Br J Cancer 66:297303 (1992B).

68

Chapter 4

65. S. Price, and M. Tisdale, Mechanism of inhibition of a tumor lipid-mobilizing factor by eicosapentaenoic acid, Cancer Res 58:4827-4831 (1998). 66. H. Hussey, and M. Tisdale, Effect of a cachectic factor on carbohydrate metabolism and attenuation by eicosapentaenoic acid, Br J Cancer 80:1231-1235 (1999). 67. P. Todorov, T. Mcdevitt, D. Meyer, H. Ueyama, I. Ohkubo, and M. Tisdale, Purification and characterization of a tumor lipid-mobilizing factor, Cancer Res 58:2353-2358 (1998). 68. K. Hirai, H. Hussey, M. Barber, S. Price, and M. Tisdale, Biological evaluation of a lipid-mobilizing factor isolated from the urine of cancer patients, Cancer Res 58:2359-2365 (1998). 69. J. Pell, J. Brown, and I. Johnson, Polyunsaturated fatty acids of the n - 3 series influence intestinal crypt cell production in rats, Carcinogenesis 15:1115-1119 (1994). 70. G. Calviello, P. Palozza, E. Piccioni, et al., Dietary supplementation with eicosapentaenoic and docosahexaenoic acid inhibits growth of Morris hepatocarcinoma 3924A in rats: effects on proliferation and apoptosis., IntI J Cancer 75:699-705 (1998). 71. G. Calviello, L. Tessitore, E. Piccioni, P. Palozza, P. Franceschelli, and G. Bartoli, EPA inhibits the development of focal proliferative lesions during hepatocarcinogenesis in rat., Gastroenterology 110:A1162 (1996). 72. N. Murray, L. Davidson, R. Chapkin, W. Gustafson, D. Shattenberg, and A. Fields, Overexpression of protein kinase C betaII induces colonic hyperproliferation and increased sensitivity to colon carcinogenesis, J Cell Biol 145:699-711 (1999). 73. N. R. Murray, C. Weems, L. Chen, et al., Protein kinase C betaII and TGFbetaRII in omega-3 fatty acid-mediated inhibition of colon carcinogenesis, J Cell Biol 157:915-20 (2002). 74. Y. Lin, M. Smit, R. Havinga, H. Verkade, R. Vonk, and F. Kuipers, Differential effects of eicosapentaenoic acid on glycerolipid and apolipoprotein B metabolism in primary human hepatocytes compared to HepG2 cells and primary rat hepatocytes, Biochim Biophys Acta 1256:88-96 (1995). 75. P. Needleman, A. Raz, M. Minkes, J. Ferrendelli, and H. Sprecher, Triene prostaglandins: prostacyclin and thromboxane biosynthesis and unique biological properties, Proc Natl Acad Sci USA 76:944-948 (1979). 76. W. Lands, Biochemistry and physiology of n-3 fatty acids, FASEB J 6:2530-2536 (1992). 77. L. Sauer, R. Dauchy, and D. Blask, Mechanism for the antitumor and anticachectic effects of n-3 fatty acids, Cancer Res 60:5289-5295 (2000). 78. E. Pizer, S. Lax, F. Kuhajda, G. Pasternack, and R. Kurman, Fatty acid synthase correlation with cell proliferation and hormone receptors expression in endometrial carcinoma:, Cancer 83:528-537 (1998). 79. J. Swinnen, F. Vanderhoydonc, A. Elgammal, et al., Selective activation of the fatty acid synthesis pathway in human prostate cancer, Intl J Cancer 88:176-179 (2000B). 80. Y. A. Yang, W. F. Han, P. J. Morin, F. J. Chrest, and E. S. Pizer, Activation of fatty acid synthesis during neoplastic transformation: role of mitogen-activated protein kinase and phosphatidylinositol 3-kinase, Exp Cell Res 279:80-90 (2002). 81. J. Adamson, E. A. Morgan, C. Beesley, et al., High-level expression of cutaneous fatty acid-binding protein in prostatic carcinomas and its effect on tumorigenicity, Oncogene 22:2739-49 (2003).


69

82. G. Velasco, D. Gómez, Pulgar, T, D. Carlin, and M. Guzmán, Evidence that the AMPactivated protein kinase stimulates rat liver carnitine palmitoyltransferase I by phosphorylating cytoskeletal components, FEBS Lett 439:317-320 (1998A). 83. G. Velasco, M. Geelen, T. Gomez del Pulgar, and M. Guzmán, Malonyl-CoAindependent acute control of hepatic carnitine palmitoyltransferase I activity: Role of Ca2+/calmodulin-dependent protein kinase II and cytoskeletal components, J Biol Chem 273:21497-21504 (1998B). 84. J. Sleboda, K. Risan, O. Spydevold, and J. Bremer, Short-term regulation of carnitine palmitoyltransferase I in cultured rat hepatocytes: spontaneous inactivation and reactivation by fatty acids, Biochim Biophys Acta 1436:541-549 (1999). 85. M. Joaquin, J. Rosa, C. Salvado, et al., Hepatocyte growth factor and transforming growth factor beta regulate 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase gene expression in rat hepatocyte primary cultures, Biochem J 314:235-240 (1996). 86. Q. Shao, N. Arakaki, T. Ohnishi, O. Nakamura, and Y. Daikuhara, Effect of hepatocyte growth factor/scatter factor on lipogenesis in adult rat hepatocytes in primary culture, J Biochem 119:940-946 (1996). 87. M. Kaibori, A. Kwon, M. Oda, Y. Kamiyama, N. Kitamura, and T. Okumura, Hepatocyte growth factor stimulates synthesis of lipids and secretion of lipoproteins in rat hepatocytes, Hepatology 27:1354-1561 (1998). 88. S. Presnell, M. Hooth, K. Borchert, W. Coleman, J. Grisham, and G. Smith, Establishment of a functional HGF/C-MET autocrine loop in spontaneous transformants of WB-F344 rat liver stem-like cells, Hepatology 28:1253-1259 (1998). 89. M. Kaibori, A. Kwon, M. Nakagawa, et al., Stimulation of liver regeneration and function after partial hepatectomy in cirrhotic rats by continuous infusion of recombinant human hepatocyte growth factor, J Hepatol 27:381-390 (1997). 90. K. Feingold, I. Staprans, R. Memon, et al., Endotoxin rapidly induces changes in lipid metabolism that produce hypertriglyceridemia: low doses stimulate hepatic triglyceride production while high doses inhibit clearance., J Lipid Res 33:17651776 (1992). 91. M. Navasa, K. Feingold, and C. Grunfeld, Effects of endotoxin and cytokines on hepatic lipid metabolism, Prog Liver Dis 15:147-170 (1997). 92. C. Magnard, R. Bachelier, A. Vincent, et al., BRCA1 interacts with acetyl-CoA carboxylase through its tandem of BRCT domains, Oncogene 21:6729-39 (2002). 93. R. Chakrabarti, and E. Engleman, Interrelationships between mevalonate metabolism and the mitogenic signaling pathway in T lymphocyte proliferation., J Biol Chem 266:12216-12222 (1991). 94. O. Larsson, and H. Blegen, Regulatory role of mevalonate in the growth of normal and neoplastic human mammary epithelial cells, Anticancer Res 13:1075-1079 (1993). 95. B. Agarwal, C. Rao, S. Bhendwal, et al., Lovastatin augments sulindac-induced apoptosis in colon cancer cells and potentiates chemopreventive effects of sulindac., Gastroenterology 117:838-847 (1999). 96. S. Jackowski, Coordination of membrane phospholipid synthesis with the cell cycle, J Biol Chem 269:3858-3867 (1994). 97. D. Hardie, D. Carting, and M. Carlson, The AMP-activated/SNF1 protein kinase subfamily: metabolic sensors of the eukaryotic cell?, Annu Rev Biochem 67:821-855 (1998).

70

Chapter 4

98. K. Tobin, H. Steineger, S. Alberti, et al., Cross-talk between fatty acid and cholesterol metabolism mediated by liver X receptor-a, Molec Endocrinol 14:741752 (2000). 99. J. D. Horton, J. L. Goldstein, and M. S. Brown, SREBPs: activators of the complete program of cholesterol and fatty acid synthesis in the liver, J Clin Invest 109:112531 (2002). 100. S. B. Joseph, B. A. Laffitte, P. H. Patel, et al., Direct and indirect mechanisms for regulation of fatty acid synthase gene expression by liver X receptors, J Biol Chem 277:11019-25 (2002A). 101. M. Foretz, C. Pacot, I. Dugail, et al., ADD1/SREBP-1c is required in the activation of hepatic lipogenic gene expression by glucose, Mol Cell Biol 19:3760-8 (1999A). 102. M. Foretz, C. Guichard, P. Ferré, and F. Foufelle, Sterol regulatory element binding protein-1c is a major mediator of insulin action on the hepatic expression of glucokinase and lipogenesis-related genes., Proc Natl Acad Sci USA 96:12737-12742 (1999B). 103. I. Shimomura, Y. Bashmakov, S. Ikemoto, J. D. Horton, M. S. Brown, and J. L. Goldstein, Insulin selectively increases SREBP-1c mRNA in the livers of rats with streptozotocin-induced diabetes, Proc Natl Acad Sci U S A 96:13656-61 (1999). 104. X. Deng, L. M. Cagen, H. G. Wilcox, E. A. Park, R. Raghow, and M. B. Elam, Regulation of the rat SREBP-1c promoter in primary rat hepatocytes, Biochem Biophys Res Commun 290:256-62 (2002). 105. Y. A. Yang, P. J. Morin, W. F. Han, et al., Regulation of fatty acid synthase expression in breast cancer by sterol regulatory element binding protein-1c, Exp Cell Res 282:132-7 (2003). 106. J. Ou, H. Tu, B. Shan, et al., Unsaturated fatty acids inhibit transcription of the sterol regulatory element-binding protein-1c (SREBP-1c) gene by antagonizing ligand-dependent activation of the LXR, Proc Natl Acad Sci USA 98:6027-6032 (2001). 107. A. Pawar, D. Botolin, D. J. Mangelsdorf, and D. B. Jump, The role of liver X receptor-alpha in the fatty acid regulation of hepatic gene expression., J Biol Chem 278:40736-40743 (2003). 108. A. R. Miserez, P. Y. Muller, and V. Spaniol, Indinavir inhibits sterol-regulatory element-binding protein-1c-dependent lipoprotein lipase and fatty acid synthase gene activations, Aids 16:1587-94 (2002). 109. Y. Pak, M. Kanuck, D. Berrios, M. Briggs, A. Cooper, and J. Ellsworth, Activation of LDL receptor gene expression in HepG2 cells by hepatocyte growth factor, J Lipid Res 37:985-998 (1996). 110. G. Skouteris, and E. Georgakopoulos, Hepatocyte growth factor-induced proliferation of primary hepatocytes is mediated by activation of phosphatidylinositol 3-kinase, Biochem Biophys Res Commun 218:229-233 (1996). 111. J. Papkoff, and M. Aikawa, WNT-1 and HGF regulate GSK3- beta activity and betacatenin signaling in mammary epithelial cells, Biochem Biophys Res Comm 247:851-858 (1998). 112. J. Swinnen, H. Heemers, L. Deboel, F. Foufelle, W. Heyns, and G. Verhoeven, Stimulation of tumor-associated fatty acid synthase expression by growth factor activation of the sterol regulatory element-binding protein pathway, Oncogene 19:5173-5181 (2000A).


71

113. J. Li, M. Mahmoud, W. Han, M. Ripple, and E. Pizer, Sterol regulatory elementbinding protein-1 participates in the regulation of fatty acid synthase expression in colorectal neoplasia, Exp Cell Res 261:159-165 (2000). 114. B. A. Laffitte, L. C. Chao, J. Li, et al., Activation of liver X receptor improves glucose tolerance through coordinate regulation of glucose metabolism in liver and adipose tissue, Proc Natl Acad Sci U S A 100:5419-24 (2003). 115. T. Ide, H. Shimano, T. Yoshikawa, et al., Cross-talk between peroxisome proliferator-activated receptor (PPAR) alpha and liver X receptor (LXR) in nutritional regulation of fatty acid metabolism. II. LXRs suppress lipid degradation gene promoters through inhibition of PPAR signaling, Mol Endocrinol 17:1255-67 (2003). 116. S. Hooks, W. Santos, D. Im, C. Heise, T. Macdonald, and K. Lynch, Lysophosphatidic acid-induced mitogenesis is regulated by lipid phosphate phosphatases and is Edg-receptor independent, J Biol Chem 276:4611-4621 (2001). 117. J. Chappell, J. Leitner, S. Solomon, I. Golovchenko, M. Goalstone, and B. Draznin, Effect of insulin on cell cycle progression in mcf-7 breast cancer cells. Direct and potentiating influence., J Biol Chem 276:38023-38028 (2001). 118. Y. Sautin, J. Crawford, and S. Svetlov, Enhancement of survival by LPA via Erk1/Erk2 and PI 3-kinase/Akt pathways in a murine hepatocyte cell line, Am J Physiol Cell Physiol 281:C2010-C2019 (2001). 119. S. Ghosh, S. J, and B. R, Lipid biochemistry: functions of glycerolipids and sphingolipids in cellular signaling, FASEB J 11: 45-50 (1997). 120. G. A. Rutter, G. da Silva Xavier, and I. Leclercq, Roles of 5'-AMP-activated protein kinase (AMPK) in mammalian glucose homeostasis, Biochem J 375:1-16 (2003). 121. H. Park, V. K. Kaushik, S. Constant, et al., Coordinate regulation of malonyl-CoA decarboxylase, sn-glycerol-3-phosphate acyltransferase, and acetyl-CoA carboxylase by AMP-activated protein kinase in rat tissues in response to exercise, J Biol Chem 277:32571-7 (2002). 122. T. Kawaguchi, K. Osatomi, H. Yamashita, T. Kabashima, and K. Uyeda, Mechanism for fatty acid “sparing” effect on glucose-induced transcription: regulation of carbohydrate-responsive element-binding protein by AMP-activated protein kinase, J Biol Chem 277:3829-3835 (2002). 123. J. J. Luiken, S. L. Coort, J. Willems, et al., Contraction-induced fatty acid translocase/CD36 translocation in rat cardiac myocytes is mediated through AMPactivated protein kinase signaling, Diabetes 52:1627-34 (2003). 124. R. Ockner, Apoptosis and liver diseases: Recent concepts of mechanisms and significance, J Gastroenterol Hepatol 16:248-260 (2001). 125. Z. Dagher, N. Ruderman, K. Tornheim, and Y. Ido, Acute regulation of fatty acid oxidation and AMP-activated protein kinase in human umbilical vein endothelial cells., Circ Res 88:1276-82 (2001). 126. G. Steinberg, A. Bonen, and D. Dyck, Fatty acid oxidation and triacylglycerol hydrolysis are enhanced after chronic leptin treatment in rats, AJP - Endocrinol Metab 282:E593-E600 (2002). 127. Y. Minokoshi, Y. Kim, O. Peroni, et al., Leptin stimulates fatty-acid oxidation by activating AMP-activated protein kinase, Nature 415:339-343 (2002). 128. C. Beauloye, A. S. Marsin, L. Bertrand, et al., Insulin antagonizes AMP-activated protein kinase activation by ischemia or anoxia in rat hearts, without affecting total adenine nucleotides, FEBS Lett 505:348-52 (2001). 129. L. Stryer, Biochemistry, W.H. Freeman and Co, New York (1995).

72

Chapter 4

130. V. Skulachev, Fatty acid circuit as a physiological mechanism of uncoupling of oxidative phosphorylation, FEBS Lett 294:158-162 (1991). 131. L. Wojtczak, and P. Schönfeld, Effect of fatty acids on energy coupling processes in mitochondria, Biochim Biophys Acta 1183:41-57 (1993). 132. L. Wojtczak, and M. Wieckowski, The mechanisms of fatty acid-induced proton permeability of the inner mitochondrial membrane, J Bioenerg Biomembranes 31:447-455 (1999A). 133. P. Schönfeld, M. Wiêckowski, and L. Wojtczak, Thyroid hormone-induced expression of the ADP/ATP carrier and its effect on fatty acid-induced uncoupling of oxidative phosphorylation, FEBS Lett 416:19-22 (1997A). 134. P. Schönfeld, and R. Bohnensack, Fatty acid-promoted mitochondrial permeability transition by membrane depolarization and binding to the ADP/ATP carrier, FEBS Lett 420:167-170 (1997B). 135. S. Mills, D. Foster, and J. Mcgarry, Interaction of malonyl-CoA and related compounds with mitochondria from different rat tissues: Relationship between ligand binding and inhibition of carnitine palmitoyltransferase I, Biochem J 214:83-91 (1983). 136. L. Drynan, P. Quant, and V. Zammit, Flux control exerted by mitochondrial outer membrane carnitine palmitoyltransferase over beta-oxidation, ketogenesis and tricarboxylic acid cycle activity in hepatocytes isolated from rats in different metabolic states., Biochem J 317:791-795 (1996A). 137. L. Drynan, P. Quant, and V. Zammit, The role of changes in the sensitivity of hepatic mitochondrial overt carnitine palmitoyltransferase in determining the onset of the ketosis of starvation in the rat., Biochem J 318:767-770 (1996B). 138. J. McGarry, and N. Brown, Reconstitution of purified, active and malonyl-CoAsensitive rat liver carnitine palmitoyltransferase I: relationship between membrane environment and malonyl-CoA sensitivity, Biochem J 349:179-187 (2000). 139. M. Young, G. Goodwin, J. Ying, et al., Regulation of cardiac and skeletal muscle malonyl-CoA decarboxylase by fatty acids, Am J Physiol Endocrinol Metab 280: E471-E479 (2001 A). 140. M. L. Casanova, C. Blazquez, J. Martinez-Palacio, et al., Inhibition of skin tumor growth and angiogenesis in vivo by activation of cannabinoid receptors, J Clin Invest 111:43-50 (2003). 141. G. Beutner, A. Ruck, B. Riede, and D. Brdiczka, Complexes between porin, hexokinase, mitochondrial creatine kinase and adenylate translocator display properties of the permeability transition pore: Implication for regulation of permeability transition by the kinases., Biochim Biophys Acta 1368:7-18 (1998). 142. N. Zamzami, C. Brenner, I. Marzo, S. Susin, and G. Kroemer, Subcellular and submitochondrial mode of action of Bcl-2-like oncoproteins, Oncogene 16:22652282 (1998). 143. I. Marzo, C. Brenner, N. Zamzami, et al., The permeability transition pore complex: a target for apoptosis regulation by caspases and bcl-2-related proteins, J Exper Med 187:1261-1271 (1998). 144. F. Fraser, R. Padovese, and V. Zammit, Distinct kinetics of carnitine palmitoyltransferase I in contact sites and outer membranes of rat liver mitochondria., J Biol Chem 276:20182-20185 (2001). 145. F. Fraser, C. Corstorphine, and V. Zammit, Topology of carnitine palmitoyltransferase I in the mitochondrial outer membrane., Biochem J 323:711718 (1997).


73

146. J. Pastorino, N. Shulga, and J. Hoek, Mitochondrial binding of hexokinase II inhibits Bax-induced cytochrome c release and apoptosis, J Biol Chem 277:76107618 (2002). 147. M. Y. Vyssokikh, and D. Brdiczka, The function of complexes between the outer mitochondrial membrane pore (VDAC) and the adenine nucleotide translocase in regulation of energy metabolism and apoptosis, Acta Biochim Pol 50:389-404 (2003). 148. G. Klug, J. Krause, A. Ostlund, G. Knoll, and D. Brdiczka, Alterations in liver mitochondrial function as a result of fasting and exhaustive exercise, Biochim Biophys Acta 764:272-282 (1984). 149. M. Diaz-Guerra, M. Junco, and L. Bosca, Oleic acid promotes changes in the subcellular distribution of protein kinase C in isolated hepatocytes., J Biol Chem 266:23568-23576 (1991). 150. Y. Nishizuka, Intracellular signaling by hydrolysis of phospholipids and activation of protein kinase C, Science 258:607-614 (1992). 151. E. Dempsey, A. Newton, D. Mochly-Rosen, et al., Protein kinase C isozymes and the regulation of diverse cell responses., Am J Physiol Lung Cell Mol Physiol 279:L429-L438 (2000). 152. L. Braun, J. Mead, M. Panzica, R. Mikumo, G. Bell, and N. Fausto, Transforming growth factor beta mRNA increases during liver regeneration: a possible paracrine mechanism of growth regulation., Proc Natl Acad Sci USA 85:1539-1543 (1988). 153. D. Bissell, S. Wang, W. Jarnagin, and F. Roll, Cell-specific expression of transforming growth factor-beta in rat liver: Evidence for autocrine regulation of hepatocyte proliferation., J Clin Invest 96:447-455 (1995). 154. M. Macias-Silva, W. Li, J. I. Leu, M. A. Crissey, and R. Taub, Up-regulated transcriptional repressors SnoN and Ski bind Smad proteins to antagonize transforming growth factor-beta signals during liver regeneration, J Biol Chem 277:28483-90 (2002). 155. T. Fynan, and M. Reiss, Resistance to inhibition of cell growth by transforming growth factor-beta and its role in oncogenesis., Crit Rev Oncog 4:493-540 (1993). 156. H. Tsushima, S. Kawata, S. Tamura, et al., High levels of transforming growth factor beta1 in patients with colorectal cancer: Association with disease progression, Gastroenterology 110:375-382 (1996). 157. J. McCormack, and R. Denton, The role of intramitochondrial Ca2+ in the regulation of oxidative phosphorylation in mammalian tissues, Biochem Soc Trans 21:793-799 (1993). 158. D. Toomey, H. Redmond, and D. Bouchier-Hayes, Mechanisms mediating cancer cachexia, Cancer 76:2418-2426 (1995). 159. D. Gough, S. Heys, and O. Eremin, Cancer cachexiaa; pathophysiological mechanisms, Eur J Surg Oncol 22:192-196 (1996). 160. J. Argilés, and F. López-Soriano, Cancer cachexia. A key role for TNF? (Review), Intl J Oncol 10:565-572 (1997). 161. M. J. Tisdale, Wasting in cancer, J Nutr 129:243S-246S (1999). 162. L. Sánchez, A. Chirino, and P. Bjorkman, Crystal structure of human ZAG, a fatdepleting factor related to MHC molecules, Science 283:1914-1919 (1999). 163. S. Wigmore, P. Todorov, M. Barber, J. Ross, M. Tisdale, and K. Fearon, Characteristics of patients with pancreatic cancer expressing a novel cancer cachectic factor, Br J Surg 87:53-58 (2000).

74

Chapter 4

164. H. Smith, and M. Tisdale, Induction of apoptosis by a cachectic-factor in murine myotubes and inhibition by eicosapentaenoic acid., Apoptosis 8:161-169 (2003). 165. K. Fearon, M. von Meyenfeldt, A. Moses, et al., Effect of a protein and energy dense n-3 fatty acid enriched oral supplement on loss of weight and lean tissue in cancer cachexia: a randomised double blind trial., Gut 52:1479-1486 (2003). 166. L. Levin, and W. Gevers, Metabolic alterations in cancer. Part I: Carbohydrate metabolism, S Afr Med J 59:518-521 (1981A). 167. L. Levin, and W. Gevers, Metabolic alterations in cancer, Part II: Protein and fat metabolism, S Afr Med J 59:553-556 (1981B). 168. K. Bennegard, F. Lundgren, and K. Lundholm, Mechanisms of insulin resistance in cancer associated malnutrition., Clin Physiol 6:539-547 (1986). 169. J. Tayek, A review of cancer cachexia and abnormal glucose metabolism in humans with cancer, J Am Coll Nutr 4:445-456 (1992). 170. A. Rofe, C. Bourgeois, P. Coyle, A. Taylor, and E. Abdi, Altered insulin response to glucose in weight-losing cancer patients, Anticancer Res 14:647-650 (1994). 171. K. Smith, and M. Tisdale, Increased protein degradation and decreased protein synthesis in skeletal muscle during cancer cachexia, Br J Cancer 67:680-685 (1993). 172. K. Lundholm, S. Edstrom, I. Karlberg, L. Ekman, and T. Schersten, Glucose turnover, gluconeogenesis from glycerol, and estimation of net glucose cycling in cancer patients, Cancer 50:1142-1150 (1982). 173. G. Hotamisligil, Mechanisms of TNF-alpha-induced insulin resistance, Exp Clin Endocrinol Diabetes 107:119-125 (1999). 174. X. Chen, N. Iqbal, and G. Boden, The effects of free fatty acids on gluconeogenesis and glycogenolysis in normal subjects., J Clin Invest 103:365-372 (1999). 175. A. Dresner, D. Laurent, M. Marcucci, et al., Effects of free fatty acids on glucose transport and IRS-1-associated phosphatidylinositol 3-kinase activity., J Clin Invest 103:253-259 (1999). 176. M. E. Griffin, M. J. Marcucci, G. W. Cline, et al., Free fatty acid-induced insulin resistance is associated with activation of protein kinase C theta and alterations in the insulin signaling cascade, Diabetes 48:1270-4 (1999). 177. P. Arner, Insulin resistance in type 2 diabetes: role of fatty acids, Diabetes Metab Res Rev 18 Suppl 2:S5-9 (2002). 178. G. Boden, and G. I. Shulman, Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction, Eur J Clin Invest 32 Suppl 3:14-23 (2002). 179. T. K. Lam, H. Yoshii, C. A. Haber, et al., Free fatty acid-induced hepatic insulin resistance: a potential role for protein kinase C-delta, Am J Physiol Endocrinol Metab 283:E682-91 (2002). 180. C. L. Soltys, L. Buchholz, M. Gandhi, A. S. Clanachan, K. Walsh, and J. R. Dyck, Phosphorylation of cardiac protein kinase B is regulated by palmitate, Am J Physiol Heart Circ Physiol 283:H1056-64 (2002). 181. C. Yu, Y. Chen, G. W. Cline, et al., Mechanism by which fatty acids inhibit insulin activation of insulin receptor substrate-1 (IRS-1)-associated phosphatidylinositol 3kinase activity in muscle, J Biol Chem 277:50230-50236 (2002). 182. R. Eastman, R. Carson, D. Orloff, et al., Glucose utilization in a patient with hepatoma and hypoglycemia: Assessment by a positron emission tomography., J Clin Invest 89:1958-1963 (1992). 183. D. Le Roith, Insulin-like growth factors, New Engl J Med 336:663-640 (1997). 184. R. Behal, D. Buxton, J. Robertson, and M. Olson, Regulation of the pyruvate dehydrogenase multienzyme complex., Annu Rev Nutr 13:497-520 (1993).


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185. R. Scholz, M. Olson, A. Schwab, U. Schwabe, C. Noell, and W. Braun, The effect of fatty acids on the regulation of pyruvate dehydrogenase in perfused rat liver, Eur J Biochem 86:519-530 (1978). 186. K. Cusi, K. Maezono, A. Osman, et al., Insulin resistance differentially affects the PI 3-kinase- and MAP kinase-mediated signaling in human muscle., J Clin Invest 105:311-320 (2000). 187. Y. Kruszynska, D. Worrall, J. Ofrecio, J. Frias, G. Macaraeg, and J. Olefsky, Fatty acid-induced insulin resistance: decreased muscle PI3K activation but unchanged Akt phosphorylation, J Clin Endocrinol Metab 87:226-234 (2002). 188. M. Brauer, R. Inculet, G. Bhatnagar, G. Marsh, A. Driedger, and R. Thompson, Insulin protects against hepatic bioenergetic deterioration induced by cancer cachexia: an in vivo 31P magnetic resonance spectroscopy study., Cancer Res 54:6383-6386 (1994). 189. A. Tsuburaya, D. Blumberg, M. Burt, and M. Brennan, Energy depletion in the liver and in isolated hepatocytes of tumor-bearing animals, J Surg Res 59:421-427 (1995).

Chapter 5 Mitochondrial Function in Cell Growth and Death

5.1.

INTRODUCTION

The evidence considered thus far strongly suggests that the pivotal role of mitochondria in regulation of cell growth and cell death is characterized by contrasting changes in mitochondrial substrate oxidation and energetics. Thus, mitosis-associated increases in and oxidative phosphorylation are fueled by aerobic glycolysis or glutamine; under hypoxic conditions, ATP is generated primarily in cytosol by anaerobic glycolysis. In either case, fatty acid oxidation is suppressed in the “average” individual mitochondrial organelle, permitting activation of glycolysis and the numerous biosynthetic, translocational, signaling, and regulatory processes required in growth, including fatty acid and sterol biosynthesis. The evidence discussed in the following paragraphs suggests that similar conditions are fostered in the rodent hepatocyte proliferative response to xenobiotic peroxisome proliferators. In the evolution of programmed cell death (apoptosis), in contrast, the central role played by mitochondria leads to compromised energetics, including decreases in and [ATP]/[AMP], increased ROS, and activation of lytic enzymes (caspases). In addition, endogenous and xenobiotic agents of growth arrest (discussed in Chapter 6: Metabolic Effects of Antiproliferative Agents) increase mitochondrial fatty acid oxidation, absolutely and/or relative to ATP synthesis, precluding mitochondrial energetic and redox conditions that would support mitosis.

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5.2.

PEROXISOME PROLIFERATOR ACTIVATED RECEPTORS (PPAR) AND LIGANDS: MITOCHONDRIAL REDOX, MACROPHAGES, AND MITOSIS

5.2.1. is a member of the nuclear receptor superfamily 1-5 . Its activator/ligands are chemically heterogeneous, and include the fibrate class of hypolipidemic agents and phthalate ester plasticizers. The fibrates immediately inhibit mitochondrial CPT-I6,7 (Fig. 5.1), and activate in the hepatocyte and other cells in which it is expressed, heterodimerization with the retinoid X is required for subsequent transcriptional activation of genes the promoters of which contain a peroxisome proliferator response element (PPRE)8. Among these are genes that encode enzymes of mitochondrial and extramitochondrial fatty acid oxidation, including cytochrome P450 4A-mediated in the SER, mitochondrial and peroxisomal and liver fatty acid binding protein (L-FABP). Significantly, malic enzyme, a 9 key contributor to fatty acid biosynthesis, is also induced by . In primary hepatocytes, long-chain dicarboxylic acid products of are activators of upregulation of peroxisomal and L-FABP10,11.

In vivo, transcriptional activation begins within approximately 1 hr after administration of peroxisome proliferators to rodents12; increases in abundance of cognate mRNAs and in activity of enzymes of fatty acid oxidation can be demonstrated in vitro within 24 hours10,11. In parallel with these effects on fatty acid oxidation, increases in expression of cell cyclerelated proteins13 and accelerated hepatocyte mitosis also begin within 24 hr14-20, progressing from periportal zone I to pericentral zone III of the hepatic acinus17, i.e., a sequence similar to that observed during liver regeneration21. Notably, hepatocyte proliferation initiated by peroxisome proliferators is less regularly associated with increased expression of immediate early genes and proinflammatory cytokines than is that which Consistent with this, Kupffer cell follows 70% hepatectomy19. proinflammatory cytokine production plays a significant role in regenerative mitosis22-26, but appears to be less important in the mitotic response to peroxisome proliferators27 and/or involves alternative cytokine mediators, 28 e.g., interleukin-1 (IL-1) and/or IL-6 rather than . These differences in hepatocellular proliferative response to peroxisome proliferators and hepatectomy appear to reflect corresponding differences in FFA

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Figure 5.1. Metabolism of fatty acids in rodent hepatocyte. Both mitochondrial and extramitochondrial FA oxidation pathways are transcriptionally activated by Direct inhibition of carnitine palmitoyl transferase-I by malonyl CoA, cytokeratins 8 and 18, or peroxisome proliferators favors diversion of FA toward esterification and extramitochondrial oxidation. Active AMPK abrogates both malonyl CoA-dependent and malonyl CoAindependent (cytokeratin-mediated) inhibition of CPT-I, thereby promoting mitochondrial fatty acid oxidation. Activation of the transcriptional regulator may reflect cellular abundance of FA. Abbreviations — AMPK: adenosine monophosphate-activated protein kinase; CPT-I: carnitine palmitoyltransferase-I; FA: fatty acid; FA-CoA: fatty acid-coenzyme A thioester; PL: glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

mobilization (see below). The duration of the initial increase in hepatocyte mitosis depends on the particular peroxisome proliferator used, but in rodents sustained administration of all eventuates in hepatocellular carcinoma; this process, and the hepatocellular proliferative response that 29-31 follows partial hepatectomy, are also . Significantly, peroxisome proliferators not only induce mitosis but also may suppress

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Figure 5.2. Growth modulator regulation of rodent hepatocyte fatty acid oxidation. Pathway arrow thickness approximates relative activity. Top: Inhibition of CPT-I, and thus mitochondrial fatty acid oxidation and ketogenesis, by the peroxisome proliferator clofibrate. Although cellular mitochondrial fatty acid oxidation increases absolutely, the fatty acid burden is diminished relative to an increased mitochondrial oxidative capacity; extramitochondrial FA oxidation is relatively enhanced. Bottom: Augmentation of mitochondrial fatty acid oxidation and ketogenesis by butyrate, with relative suppression of extramitochondrial FA oxidation. Abbreviations — CPT-I: carnitine palmitoyltransferase-I; FA: fatty acid; peroxisome proliferator-activated

hepatocyte apoptosis, both spontaneous and induced by diverse agents including DNA damage, and Fas ligation32. 5,33,34 Long chain fatty acids are activator ligands of , while evidence that the peroxisome proliferators themselves are direct activators has been less conclusive 1-3 . Activation of by 8(S)hydroxyeicosatetraenoic acid33, 34 and WY 14,64333, 35 has been reported, whereas evidence pertaining to leukotriene B4 is conflicting33, 35. However, a leading interpretation is that, by inhibiting CPT-I6, 7, peroxisome proliferators induce extramitochondrial cellular fatty acid overload (Figs. 5.1 and 5.2), 1,2,36-38 which in turn activates . This concept is consistent with the observation that hepatocellular fatty acid overload in the absence of xenobiotic peroxisome proliferators, e.g., resulting from high fat diet, diabetes mellitus, and starvation, is also associated with activation of genes39-42. Thus, although certain eicosanoids and xenobiotic peroxisome proliferators theoretically could activate by


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dual mechanisms, i.e., directly as activator ligands and indirectly by perturbing cellular fatty acid metabolism, it is unclear for some of these agents that their cellular concentrations in vivo are adequate to permit direct activation. In contrast, cellular fatty acids themselves are clearly sufficient, and in most circumstances necessary, for activation10,11,43. In any case, recent evidence indicates that activation of transcriptional regulation by PPARs requires in addition their interaction with the respective FABPligand complexes44, 45, and that association with ligand prevents ubiquitination and proteosomal degradation46. Activities of and are subject to regulation by phosphorylation47-49 (see below, and Chapter 5.2.2.: How do peroxisome proliferators induce hepatocyte mitosis and, if fatty acid-driven, why does it not also result from fatty acid overload associated with high fat diet, diabetes mellitus, or starvation? An explanation for the mitogenic effect of the peroxisome proliferators, for its absence in other hepatocyte fatty acid overload states, and for the differences in immediate early gene and inflammatory cytokine expression between peroxisome proliferator- and hepatectomy-induced hepatocyte proliferation, may reside in the striking differences in mitochondrial fatty acid oxidation that characterize these circumstances. Inhibition of CPT-I by xenobiotic peroxisome proliferators effects a relatively extramitochondrial partitioning of cellular fatty acids, restricting fatty acid entry into an expanding mitochondrial compartment the oxidative capacity of which is also increasing, as occurs in proliferating cells in the absence of peroxisome proliferators50. As a result, although the rate of mitochondrial fatty acid oxidation by the cell as a whole may increase absolutely, it is diminished in relation to an augmented mitochondrial oxidative capacity and citric acid cycle activity51. Indeed, expression of itself is inhibited by under hypoxic conditions52 (also, see Chapter 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation). Accordingly, peroxisome proliferators decrease mitochondrial matrix reducing potential (reflected in the ratio of to [acetoacetate]), e.g., in human subjects under controlled experimental conditions53, in perfused livers prepared from peroxisome proliferator-pretreated rats54, 55 and in hepatocyte cultures exposed to these agents in vitro only56. This redox shift, by diminishing otherwise excessive intramitochondrial generation of ROS, fosters an insulin-like enhancement 57 of glycolysis, pyruvate dehydrogenase activity, lipogenesis, and . Moreover, the mitogenic effect of insulin includes a MAPK-dependent phosphorylation of PPARs, enhancing the transcriptional activity of 48 while inhibiting that of .

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Together, these favorable changes in intramitochondrial redox status reflect a decrease in fatty acid oxidation relative to mitochondrial oxidative capacity. This would provide at least a partial explanation for activator-induced decreases in cellular oxidative stress, signaling, and 58 proinflammatory cytokine production in aging mice , for decreases in endothelial cell expression of and vascular cell adhesion molecule-159, and for suppression of experimental atherosclerosis 60, 61. They may also account for the above noted observation that peroxisome proliferators induce a lesser hepatic and proinflammatory cytokine repsonse than does partial hepatectomy19; in the latter, greatly increased FFA flux subjects both hepatocytes and Kupffer cells to augmented fatty acid oxidation and oxidative stress (see Chapter 8: Metabolism and Gene Expression in Liver Regeneration). Furthermore, these direct effects of FFA-induced oxidative stress after partial hepatectomy may provide an explanation for the greater dependence of hepatocellular mitosis on macrophage (Kupffer cell) activation and production in that setting, as compared with mitosis induced by peroxisome proliferators 22, 24, 26, 27. Decreased intramitochondrial oxidative stress not only contributes to diminished activation, but may also increase expression62. The effects of PPAR activation on macrophage function and survival are complex. expression was not detected in Kupffer cells25, suggesting that peroxisome proliferator-induced changes in Kupffer cell fatty acid 22,24,26 metabolism and secretion are . In the arterial wall, in contrast, monocyte-derived macrophages express both 49,63,64 and . In these key celluar determinants of the evolution of the atherosclerotic lesion, mechanisms otherwise similar to those utilized in Kupffer cells appear to be involved in the generation of proinflammatory cytokines in response to lipid overload49,59,63-69 . Further evidence in support of the importance of a mitochondrial origin of ROS in macrophages is provided in dramatic fashion by the inverse relationship between ROS generation and expression of uncoupling proteins (UCP), which dissipate intramitochondrial ROS and oxidative stress (see Chapter 6.4.: Salicylates and other Nonsteroidal Anti-Inflammatory Drugs). Thus, deletion of UCP expression in macrophages augments both ROS generation and killing of parasites70. Moreover, deletion of the adipocyte FABP isoform aP2, normally expressed in macrophages, is protective against atherosclerosis71,72 (also, see Chapter 7.4.: Consequences of Intramitochondrial Oxidative Stress: Atherosclerosis and Beyond). is also expressed in murine T and B lymphocytes73, implying its possible role in immune regulation. In contrast to the compartmentalizing effects of peroxisome proliferators on intracellular fatty acid distribution and utilization, an absolute increase in cellular fatty acid load (e.g., resulting from elevated plasma [FFA] associated


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with high fat diet, obesity, insulin resistance, diabetes mellitus, or starvation), may be sufficient, per se, to drive accelerated mitochondrial fatty acid oxidation74. In addition to the absence of CPT-I inhibition by a xenobiotic peroxisome proliferator, this may reflect fatty acid override of physiological CPT-I control75-79, likely attributable at least in part to fatty acid-induced changes in mitochondrial membrane physical properties79-81. As a result, states of fatty acid excess may exert an inhibitory effect on proliferation of normal cells through the combined effects of: 1) sustained and excessive increases in mitochondrial matrix reducing potential, ROS generation (oxidative stress), and (in hepatocytes and astrocytes) ketogenesis; 2) fatty acid inhibition of glycolysis and of the key citric acid cycle-related enzymes pyruvate dehydrogenase and 2-oxo-glutarate dehydrogenase82, and inhibition of the latter by ROS83; 3) fatty acid inhibition of PI3K-mediated insulin signaling (and likely wg/Wnt-like signaling generally)74, 84-91; and 4) relatively weak uncoupling of oxidative 92-96 phosphorylation by underivatized fatty acids, compromising . These consequences of increased mitochondrial exposure to and oxidation of fatty acids are diametrically opposed to the conditions required for cell proliferation, and to the growth-promoting effects of peroxisome proliferators and insulin. Moreover, they closely resemble the changes induced by certain growth inhibitors as discussed below (see this Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer; Chapter 6: Metabolic Effects of Antiproliferative Agents; and, Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). In addition to increasing mitochondrial ROS generation, fatty acid excess may augment ROS-generating peroxisomal fatty acid It is clear that administration of fibrate hypolipidemic agents does not elicit in human subjects peroxisomal responses that are quantitatively similar to those in rodents97-99, despite the fact that is expressed in human liver. These differences may reflect lower abundance98,100 and/or potentially inactivating differences in the signaling pathway 97, 99. Nevertheless, human does participate in the regulation of gene expression in response to fibrates, in both liver101 and HepG2 cells102, and in clinical studies is clearly implicated in the induction of significant changes in mitochondrial fatty acid oxidation, redox potential, and ketogenesis as discussed above53. As noted, putative ligands may also act independently of e.g., by inhibiting CPT-I6,7 or inducing lymphocyte IL-4 expression103.

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5.2.2. In contrast to the proliferative effects of promotes cell differentiation and is generally antiproliferative, as demonstrated by its central role in adipocyte differentiation and the effects of activator/ligands on growth of cancer cell lines and experimental tumors in vivo. In vivo, the heterodimer appears to be activated as a transcriptional regulator by certain fatty acids and eicosanoids. In addition to its abundant expression in adipose tissue and its importance in adipocyte differentiation, is expressed in many other tissues, including macrophages and colonic epithelium, and it contributes to the pathogenesis of the endothelial lesion of atherosclerosis (see Chapter 7.4.1.: Atherosclerosis and Arterial Hypertension). Consistent with its suppression of macrophage activation and pro-inflammatory cytokine expression68, 104, and its perceived normal role as a mediator of cell differentiation, expression in Caco-2 cells is induced by the antiproliferative short chain 106 fatty acid butyrate105, and in monocytes is regulated by . Also consistent with its antiproliferative effect, and augment mitochondrial oxidation and, variably, gluconeogenesis and Forkhead, in diverse tissues107,108 (also, see Chapter 7.4.2.: Adverse Effects of Fatty Acid Oxidation-Induced ROS in Various Organs). Activator/ligands of supppressed transcriptional activation of COX109 2 and proinflammatory cytokines110. ligands also induced differentiation or decreased growth, in vitro or in vivo, of cells derived from human liposarcomas111, cancers of the breast and prostate112-114, and colon115, 116; despite these significant observations, a preliminary phase II trial of troglitazone treatment of breast cancer was negative117, growth arrest and/or apoptosis in part reflect impaired cell cycle regulation118,119 and/or inhibition of tumor angiogenesis120. Surprisingly, however, the thiazolidinedione activator/ligands enhanced rather than suppressed the formation of colon polyps121 and tumors122 in mice, which bear an Apc mutation predisposing to tumorigenesis in this murine model of familial adenomatous polyposis. In the latter study, levels were increased in mouse colon by activation while cyclooxygenase-2 (COX2) levels were unaffected. The opposite responses reported between these two studies and those cited in the previous paragraph possibly reflect unique characteristics of the mouse model, such as tumor-associated phosphorylation-dependent differences in activity48,123,124, or relative insensitivity to because of pre-existing Apc pathway defects125. In two studies in which the human colon cancer cell line HT 29 was used, expression of was 116 122 increased , and activation induced an increase in . The

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Figure 5.3. Apoptosis: interaction of mitochondria and plasma membrane receptors. Mitochondrial- and receptor-initiated pathways converge at caspase 3. Yellow lines indicate membrane receptor signaling-initiated effects on mitochondria. Red lines indicate primary mitochondrial events and their effects on Fas and/or the membrane receptor-activated caspase cascade; the dashed line indicates an hypothesized mitochondrial mediation of at least some instances of Fas ligand-independent Fas activation (with or without Fas trimerization). AIF may exert pro-apoptotic effects in both cytoplasm and nucleus. Abbreviations -- AIF: apoptosis-inducing factor; APAF-1: apoptosis protease-activating factor-1; Bcl-2 family members: Bcl-2 (anti-apoptotic), Bid (pro-apoptotic); Fas, FasR: Fas receptor (CD95); ROS: reactive oxygen species; TNF: tumor necrosis factor; TNFR-1: tumor necrosis factor receptor-1; UVB: ultraviolet B light Reproduced with permission 148 .


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effect of thiazolidinediones on HT 29 cell growth was not described in either report, but subsequently were shown to induce apoptosis in this cell line126. Recent findings provide insight and raise new questions concerning these antiproliferative effects. Thus, troglitazone, and the 12,l4 natural ligand -prostaglandin may produce similar effects independent of itself110,127-131. The mechanism is not known. However, the role of as an activator ligand of in vivo has been questioned132. In addition, troglitazone posses a quinone-like function which could contribute to its demonstrated ROS scavenging activity133 or, under different conditions, to an apoptotic/antiproliferative effect analogous to that of menadione134.

5.2.3. A third PPAR, i.e., (synonymously an Apcsuppressible, component in wg/Wnt-like signaling, has been implicated in the regulation of lipid metabolism and cell proliferation135-138, and as a target for NSAID-induced growth inhibition135,136,138 .(see Chapter 6.4.: Salicylates and Other Nonsteroidal AntiInflammatory Drugs). Its expression, and that of COX-2, are increased in endometrial carcinoma139. is important in terminal adipocyte differentiation as an inducer of expression140; it also regulates 141 macrophage lipid accumulation , and modulates and 142 activity . Like is also expressed in colonocytes in which its function, while unknown136, appears to be unrelated to cancer143.

5.3.

MITOCHONDRIAL FUNCTION IN APOPTOSIS

In addition to the importance of and ATP generation in cell proliferation, mitochondria are also involved at the opposite extreme of the growth spectrum, i.e., apoptosis, or programmed cell death144-148. Understanding of this complex process is evolving rapidly; it may be initiated by several different events and may proceed via at least two major pathways; it often involves shifts in homeostasis149. Following is a brief discussion of necessarily selected evidence concerning the relationship of mitochondrial function to the overall process of apoptosis (Fig. 5.3 - facing this page). The major cell components involved in apoptosis include: 1) members of the TNF cell surface receptor family, especially TNF receptor 1 (TNFR1) and Fas; 2) mitochondria; 3) pro- and anti-apoptotic members of the Bcl-2 protein family, the role of which is understandable to a large extent in terms of their mitochondrial effects; and 4) a group of cysteine proteases

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(“caspases”) which cleave substrates at specific aspartate residues. The death program begins either with a disturbance of mitochondrial function or with activation (usually involving trimerization) of TNFR1, Fas, or other TNFR family members by their respective ligands, e.g., and Fas ligand (FasL). As the apoptotic process evolves in a given cell, both mitochondria and surface receptors usually participate. Importantly, alternative TNFR1 signaling pathways150 may activate PI3K/Akt/PKB- and cell survival mechanisms151-154. As PI3K/Akt/PKB signaling is subject to inhibition by fatty acids74, 84-91, 155-157, selection between these opposing TNFR1 signaling alternatives may reflect at least in part the status of cellular fatty acid metabolism (see Chapter 3.3.2.: Wingless/WntLike Signaling: Convergence of Antecedents and the Unpredictable; and, Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin). Fas signaling may also initiate survival signaling under certain conditions158. When apoptosis is initiated at the cell surface, mitochondria may become involved through activation (truncation) of the pro-apoptotic Bcl-2 family member Bid by caspase 8159, cleavage of the anti-apoptotic Bcl-2 (or Bcl-xL, presumably) to a pro-apoptotic form by caspase 3160 (Fig. 5.3 - facing page 85), or via activation of NO synthase and thus NO-mediated inhibition of mitochondrial respiration161. When apoptosis is initated at the mitochondria, it frequently reflects intramitochondrial generation of oxidative stress, e.g., resulting from disruption of the electron transport chain by chemical or physical agents146,162 or ischemia/reperfusion injury. Depending on its severity, intramitochondrial oxidative stress may initiate an extremely rapid apoptotic sequence163, or may be associated with growth arrest, e.g., as occurs prior to the onset of mitosis that follows partial hepatectomy (see Chapter 8: Metabolism and Gene Expression in Liver Regeneration). Irrespective of whether mitochondrial participation in apoptosis is primary, or secondary to TNFR1/Fas activation, it frequently is characterized by conversion of the VDAC complex (Fig. 5.4) to the permeability transition pore, with collapse of the latter often preceded by early and transient 164 hyperpolarization . Formation of the permeability transition pore, an early event in dexamethasone-induced lymphocyte or thymocyte apoptosis146, 165, 166, is usually associated with decreased intramitochondrial thiols167 and increased oxidative stress. These changes are generally linked to the release from mitochondrial intermembrane space of procaspases 2, 3, 8, and 9168,169, and caspaseactivating proteins including the second mitochondrial activator of caspases (Smac/Diablo)170, and cytochrome c171; intramitochondrial localization of procaspases has been questioned recently172. Procaspase 2 is widely distributed within the cell and, upon activation, may promote release of mitochondrial intermembrane proteins173. Apoptosis-inducing factor (AIF)


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may normally function in mitochondrial redox regulation, via incompletely understood mechanisms174; upon release during apoptosis, it exerts its proapoptotic effects at both nuclear and cytoplasmic locations175. In cytosol cytochrome c combines with Apaf-1 (apoptotic protease-activating factor-1, released from the mitochondrial membrane), dATP or ATP, and procaspase 9. The resulting complex (the “apoptosome”) cleaves and activates procaspase 9176. Antiapoptotic effects are exerted by cellular inhibitors of apoptosis (cIAP 1, 2), which directly inhibit caspases and act as ubiquitin ligases for Smac/Diablo177.

Figure 5.4. Mitochondrial voltage-dependent anion channel (VDAC) components and relationship to electron transport chain; schematic representation. VDAC complex-mediated juxtaposition of hexokinase II and the adenine nucleotide translocase in proliferating cells optimizes energetics of glucose phosphorylation to glucose-6-phosphate and entry into the glycolytic pathway, leading to pyruvate formation. Detailed structural relationships remain incompletely defined (see Chapter 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation). Increased mitochondrial fatty acid oxidation inhibits glucose uptake, hexokinase, and pyruvate dehydrogenase, and predisposes to electron “leakage” from the electron transport chain; the latter results in one-electron reduction of oxygen and formation of and derivative ROS. Excess cellular fatty acids may disrupt hexokinase II, VDAC, and the contact sites. Unlike anti-apoptotic Bcl-2 family members, pro-apoptotic members may associate with, and perturb, the mitochondrial membrane after activation of the apoptotic cascade. Abbreviations — CYT C: cytochrome c; inner mitochondrial membrane potential; superoxide anion; UBIQ: ubiquinone; ROS: reactive oxygen species; VDAC: voltage-dependent anion channel. With permission (see Fig. 3.4.)

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5.3.1. The Bcl-2 Protein Family The importance of mitochondria and of intramitochondrial oxidative stress in apoptosis and in growth regulation more broadly is underscored by accumulating evidence concerning the Bcl-2 protein family. Antiapoptotic (pro-survival) members of the Bcl-2 family (Bcl-2 and Bcl-xL) inhibit 178-180 mitochondrial release of cytochrome c and AIF and disruption of .

Figure 5.5. Wingless/Wnt-like signaling: cell growth and suppression of cell death. Activation of PI3K-Akt/PKB via diverse receptor and non-receptor tyrosine kinases (including EGF, HGF, insulin, IGF-1, PDGF, and receptors) and/or ROS leads to Akt/PKB phosphorylation, activation of SREBPs, MDM2, and telomerase, and inactivation of the apoptosis-promoting and/or growth inhibiting proteins Bad, caspase 9, Forkhead, p53, and (not shown) Inactivation of abrogates its inhibition of proteins important in storage and utilization of glucose (glycogen synthase, pyruvate dehydrogenase), cell cycle progression (cyclin D1), and gene transcription The resulting increase in c-myc expression leads, among its other effects, to transcriptional activation of the gene encoding the cyclin-dependent kinase cdk4. Abbreviations — COX 2: cyclo-oxygenase 2 (prostaglandin G/H synthase 2); EGF: epidermal growth factor; HCC: hepatocellular carcinoma; HGF: hepatocyte growth factor; IGF-1: insulin-like growth factor-1; PDGF: platelet-derived growth factor; PDH: pyruvate dehydrogenase; SREBPs: sterol regulatory element binding proteins; tumor necrosis


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they may also protect against apoptosis in growth-arrested IL-3-deprived cells by inducing a “metabolic arrest” associated with decreased glycolysis l81,182 and . The protective effect of anti-apoptotic Bcl-2 family proteins may be augmented by an inactivating phosphorylation of the pro-apoptotic protein Bad. As noted earlier, this and several other survival-enhancing (apoptosis-suppressing) reactions154,183-186, including activation of mitochondrial hexokinase182, 187, are mediated by the serine-threonine protein kinase Akt/PKB (see Fig. 5.5, and Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). Akt/PKB is activated downstream of PI3K, an effector of the signaling cascades initiated by insulin/IRS-I, HGF/c-Met, interferon and other growth factors188-190, 154 as well as by (also, see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin). Bcl-xL prevents early changes in mitochondrial membrane potential, including excessive increases that otherwise may lead to mitochondrial swelling 191 , and the decreases associated with ROS generation192. The activity of Bcl-2 may be modified positively or negatively by several implicated kinases193, may be Bax heterodimerization-dependent or -independent194, and may protect against cell death despite cytochrome c entry into cytosol, whether released from mitochondria by Bax195 or injected directly196,197 . At least in part, the antiapoptotic effect of Bcl-2 appears to reflect its sequestration of the Bidmediated active conformer of Bak and possibly of Bax198. Thus, Bcl-2 and Bcl-xL represent pro-survival and anti-oxidant factors in the face of diverse pro-apoptotic stimuli199, actions which can be understood to a large extent in terms of their direct and/or indirect effects on mitochondrial integrity. Although the mechanism remains undefined200, release of apoptosisrelated proteins from the intermembrane space is believed to occur as a consequence of the mitochondrial permeability transition or outer membrane rupture. It may also occur independently of these events146,201,202, via a novel large pore formed by hetero-oligomerization between truncated Bid and Bak203, as a or -independent effect of Bax204, or an interaction between Bax and VDAC205. The latter would be consistent with other evidence that VDAC interacts with Bcl-2 family members206,207, that mitochondrial contact sites are involved208, and that binding of hexokinase II to VDAC inhibits Bax-induced release of cytochrome c209,210 . In addition, cytochrome c release requires disruption of its interaction with cardiolipin211 and remodeling of mitochondrial cristae212, potentially perturbing other electron transport chain components213. In any case, apoptosis appears to require either Bax or Bak214. This reflects in part their regulation of release from the endoplasmic reticulum215, as well as their interaction with cardiolipin to generate large pores in the outer mitochondrial membrane216.

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In cytosol, cytochrome c activation of the apoptosome may be inhibited by the small stress (“heat shock”) protein Hsp27217,218. In a time-dependent manner219, transfer of cytochrome c to cytosol also compromises electron transport chain function beyond complex III, thereby promoting the intramitochondrial generation of ROS, depletion of thiols167, and oxidative stress220. Cytosolic cytochrome c may re-enter the intermembrane space and transiently support electron transport chain function 219 . Collectively, however, the net effect of these events is to convert the mitochondrial power plant to an engine of cell destruction.

5.3.2. wg/Wnt-Like Signaling: Convergence of Mitochondria, p53, and Telomerase Important interactions of Bcl-2 family members and mitochondria with the tumor suppressor protein p53 and with telomerase have been identified221. p53’s well established role as an antiproliferative transcriptional activator222 has been shown to include increased expression of and Bax. In addition, however, p53 directly impacts the signaling cascades that link intermediary metabolism to growth and survival signaling. Thus, p53, in its mutually inhibitory regulatory interaction with Akt/PKB223,224, increases the expression of PTEN, the inhibitor of wg/Wntlike PI3K/Akt/PKB signaling225. p53 also suppresses expression of SREBP1c226 and thereby the lipogenic response required in proliferating cells (see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable; and Chapter 4.4.: Fatty Acids, Modulation of Cell Growth, and the AMP-Activated Protein Kinase). Moreover, p53 activates the expression of two related pro-apoptotic BH3only227 proteins (p53-upregulated modulator of apoptosis — and which bind to Bcl-2 and Bcl-xL and rapidly induce apoptosis228,229. In addition, a fraction of cellular p53 itself rapidly localizes to the mitochondria under conditions which predispose to apoptosis223,230-233 . This activates initial events in apoptosis, including formation of a complex between p53 and antiapoptotic Bcl-2 family members Bcl-2 and Bcl-xL (which are also subject to formation of complexes with p53-induced PUMAs), leading to outer membrane permeabilization234, altered cytochrome c release, and caspase activation. Overexpression of anti-apoptotic Bcl-2 family members abrogates apoptosis and p53’s mitochondrial localization, but not its induction of cell cycle arrest231. p53 may also interact physically and/or functionally with NADH quinone oxioreductase235 and may directly activate 236 .


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Participation of p53 in apoptosis also appears to involve a not fully characterized functional and physical interaction with telomerase237, 238. Thus, impaired telomerase activity and/or excessively shortened telomeres may induce growth arrest or apoptosis. The latter, which may be p53dependent or p53-independent, is prevented by Bcl-2239, suggesting mitochondrial involvement. Telomerase also inhibits apoptosis, reflecting its phosphorylation-dependent activation by wg/Wnt-like signaling via PI3K and Akt/PKB240 (also, see Fig. 5.5). Accordingly, the anti-apoptotic action of telomerase, being Akt/PKB-dependent, represents one of several examples of the mutually opposing actions between pro-apoptotic forces (including p53) and those of wg/Wnt-like survival signaling, as discussed above and in the following paragraphs. Telomerase and p53 also may interact in carcinogenesis241, 242. Several lines of evidence further emphasize the importance of mitochondrial metabolism and energetics in the control of cell survival and death. Thus, Bcl-2 regulates inner mitochondrial membrane proton flux and both in protecting against apoptosis243 and under non-stress conditions244, suggesting a more general role in the regulation of mitochondrial function. In addition, resistance of various human and murine leukemia cell lines to dexamethasone-induced apoptosis was directly correlated with cellular ATP levels and Bcl-2 expression245. Bcl-2 expression also protected against cytotoxicity in L-929 cells246 and hypoxia-induced apoptosis in cell culture and solid tumor cells in vivo178,247. These actions are consistent with Bcl-2’s salutary metabolic and mitochondrial effects, and support TNFR1 survival signaling through rather than through the death-inducing DISC (see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin). Conversely, pro-apoptotic Bax and Bak mediate the mitochondrial apoptotic response to hypoxia248. Notably, the ability of Bcl-2 to protect against xenobiotic respiratory chain inhibitors was variable depending on cell type and conditions178,249. Glutamine as sole substrate maintained L-929 cell ATP concentrations equal to those obtained with glucose, but unlike glucose predisposed these cells to apoptosis250. Possibly, this reflects: a) less stringent control of glutamine-supported electron transport chain flux than is effected by PDH-mediated glucose/pyruvate oxidation, thereby resembling the consequences of excess fatty acid oxidation in predisposing to increased ROS generation (see Chapter 6.2.: Butyrate; and Chapter 7.2.: Adverse Effects of Fatty Acids on Mitochondrial Function); b) inhibition of mitochondrial glutathione transport by glutamine-derived glutamate251; or c) both of these actions. Evidence of a physical interaction of CPT-I with Bcl-2 but not Bax252, and the localization of both CPT-I81 and Bcl-2 to the contact sites between mitochondrial inner and outer membranes,

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suggests the possibility of a direct relationship of the Bcl-2 family to mitochondrial fatty acid metabolism. Consistent with diverse evidence in non-hepatic cells that a disturbance in mitochondrial energetics predisposes to apoptosis, hepatocytes located in the relatively oxygen-poor pericentral zone 3 of the hepatic acinus253,254, or maintained in culture under zone 3equivalent conditions255, were more susceptible to apoptosis than those in periportal (zone 1) or equivalent. In hepatocytes and certain immunocytes mitochondria are obligatory downstream mediators of the apoptotic cascade initiated by Fas256. The role of ceramide in apoptosis has been controversial257-260. Available evidence indicates that it is formed early in the process, may be a precursor of mitochondrial ganglioside GD3261,262, may increase outer membrane permeability to proteins263, and that it directly or indirectly impairs electron transport chain function at complex III264-267. Ceramide also inhibits Akt/PKB-mediated wg/Wnt-like signaling268. These interactions thus reflect antagonism between opposing signal transduction pathways for growth arrest or death (ceramide) and for cell survival or proliferation (wg/Wnt-like) (see above, this section, and Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). In addition to ceramide, and/or mediated in part by it, apoptosis associated with electron transport chain inhibition at complex III may also be induced in L929 269 fibrosarcoma cells exposed to and in lymphoblastoid cells exposed 249 to classical xenobiotic inhibitors . On the basis of these considerations, integration of intermediary metabolism and signal transduction can be taken to higher levels of complexity and of importance to regulation and survival for the cell and the organism (also, see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). Accordingly, wg/Wntlike PI3K/Akt/PKB signaling, activated by any of several polypeptide growth factors, as well as by inhibition of in turn activates or promotes several pro-survival and/or proliferative influences. Thus, it: 1) increases the activity of several glycolytic pathway components, among them mitochondria-associated hexokinase II, the effects of which include stabilization of the VDAC complex and suppression of Bax-induced cytochrome c release; 2) suppresses mitochondrial fatty acid oxidation, through its activation of both SREBP-linked lipogenesis (and thereby malonyl CoA-induced inhibition of CPT-I), as well as of LXR-mediated suppression of fatty acid oxidation pathways; 3) activates telomerase, and thereby its anti-apoptotic effect; 4) inactivates thereby promoting glycolysis, transcriptional regulation and cell cycle progression, and abrogation of cell-cell adherens junctions; 5) modulates cellular distribution and regulatory function of the Cip/Kip cell


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cycle inhibitors; 6) inactivates the cytosolic inhibitor of thereby promoting the latter’s transcriptional activation of genes important in cell survival and redox regulation, including pro-survival Bcl-2 family members; 7) stabilizes thereby activating the pro-survival and proliferative response classically associated with hypoxia; and 8) directly activates VEGF expression while inactivating antiproliferative Forkhead transcription factors. In addition, PI3K/Akt/PKB’s mutually opposing interaction with p53 contributes to suppression of the latter’s antiproliferative and apoptosisinducing effects. These antiproliferative and antisurvival effects of p53 include:1) increased expression of PTEN (lipid phosphatase inhibitor of PI3K/Akt/PKB signaling), of the cell cycle inhibitor and of the pro-apoptotic Bcl-2 family members Bax and and 2) p53 interaction with Smads in mediation of receptor signaling; and 3) p53 localization to mitochondria where it participates in formation of apoptosisinducing complexes with anti-apoptotic (pro-survival) Bcl-2 family members Bcl-2 and Bcl-xL. PI3K/Akt/PKB also interacts with other determinants of survival and growth: 1) it is stabilized by Hsp 27 and Hsp 90; 2) it is inhibited by ceramide; and 3) it inactivates several pro-apoptotic factors, including the Bcl-2 family member Bad, formation of the FAS-associated DISC/procaspase 8 complex, and (acting in conceit with Bcl-2/Bcl-xL) Baxinduced mitochondrial release of cytochrome c and AIF. Thus, perhaps uniquely among growth factor signaling pathways, wg/Wnt-like PI3K/Akt/PKB signal transduction resides at — and effects — the coordinated convergence at which intermediary metabolism, cell-cell interaction, cell cycle progression, and transcriptional regulation are marshalled in a pro-survival, growth-promoting milieu in support of closely regulated cell cycle progression and cell proliferation.

5.3.3. Mitochondria and FasL-Independent Activation of Fas In addition to the well-established interactions between the mitochondria and surface membrane death receptors, it is now clear that Fas may be activated in the absence of Fas ligand. In many such instances, moreover, convincing evidence suggests that this occurs subsequent to an antecedent, and possibly initiating, disturbance of mitochondrial function148, e.g., caused by ultraviolet B (UVB) light270-273, toxic (non-polar) bile acids274,275 menadione134, NSAIDs276, and cancer chemotherapeutic agents277, and that it may be an obligate mediator of butyrate-induced apoptosis278. UVB also altered signaling279 and cell cycle regulation280, several hours after the mitochondrial changes.

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The concept that perturbed mitochondrial function may have broader significance and a more central role in apoptosis than has been appreciated is supported by evidence that toxic (non-polar) bile acid-induced apoptosis is associated with increased plasma membrane expression of Fas and TRAILreceptor 2281, that ethanol-induced mitochondrial dysfunction is associated with increased expression of Fas282, and that the non-toxic bile acid ursodeoxycholic acid prevents the mitochondrial changes, and apoptosis, 283,284 caused by diverse agents including toxic bile acids and . This 285 protective effect, and that afforded by IGF-1 , appear to be mediated by PI3K/Akt/PKB285,286 and/or mitochondrial GSH283. They resemble in scope, if not in specific mechanism, the broad antiapoptotic effects of the peroxisome proliferator nafenopin32 (see Chapter 5.2.1.: While the mechanism of FasL-independent Fas activation (and its plasma membrane expression) is unknown, the adverse mitochondrial effects of those chemical and physical agents that elicit this phenomenon would in common include generation of ROS and/or partial depletion of ATP, either of which is sufficient to induce Fas expression287-289, and/or Fas-mediated apoptosis288,289. Moreover, the activation of or its subunits, required under certain conditions for Fas expression290,291 , would be an expected consequence of mitochondrial oxidative stress. The mitochondrial location of procaspase 8169, an implied mitochondrial role in ceramide generation292, and the above noted evidence of early Fas-activated NO inhibition of mitochondrial respiration161, may also contribute to the potentially bidirectional interaction between Fas and mitochondrial function. In any case, it appears likely that in a diversity of settings, mitochondrial initiation of apoptosis could lead secondarily to recruitment of Fas as an active participant in the death program, albeit in these instances “downstream” of mitochondia. This proposed activation sequence is consistent with increased hepatocellular apoptosis and Fas expression (plausibly induced by fatty acid oxidation — see Chapter 7.4.2.: Adverse Effects of Fatty Acid OxidationInduced ROS in Various Organs) in human nonalcoholic steatohepatitis293, and is directly supported by non-polar bile acid-induced, FADDFurthermore, human independent, activation of caspase 8294, immunodeficiency virus-1 (HIV-1) infection may induce mitochondrial membrane permeabilization-mediated T lymphocyte death, independent of both caspase activity and death receptor signaling295. Moreover, as discussed above (see Chapter 4.4.: Fatty Acids, Modulation of Cell Growth, and the AMP-Activated Protein Kinase), a decrease in [ATP] induced by these adverse mitochondrial effects would increase [AMP] and [AMP]/[ATP] through the action of adenylate kinase. This in turn would activate AMPK, thereby increasing CPT-I activity and mitochondrial fatty acid oxidation, and further contribute to the generation


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of intramitochondrial oxidative stress. Augmentation of CPT-I activity and mitochondrial fatty acid oxidation would also result from caspase-3mediated disruption of intermediate filaments296, reflecting the importance of cytokeratins 8 and 18 in CPT-I regulation297,298 and cell survival299, e.g., in protecting against hepatotoxic300 and and Fas-induced301, 302 cell death. 303 Caspase 3 also disrupts wg/Wnt signaling through cleavage of . Mitochondrial and cell membrane death receptor signaling converge at caspase 3 (Fig. 5.3 - facing page 85), the substrates of which include several of the aforementioned proteins that are essential to cell survival. Thus, this pivotal lytic agent of death links disruption of survival signaling, mitochondrial integrity, and metabolism. Caspase 3 targets include 303 303 Akt/PKB304, , Bcl-2160, intermediate filaments296, , and 306 . Depending on its residual activity, inactivation of could disable regulation of G1 cell cycle progression, thereby promoting cell cycle-associated death (see this Chapter 5.3.4. Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer) or impairing D cyclin activation307,308. In addition to these early events in the apoptotic cascade, extramitochondrial changes ensue, including phosphatidyl serine display on the plasma membrane outer leaflet triggered by the fall in cellular [ATP]309, chromatin condensation, DNA fragmentation, and increased plasma membrane permeability. As chromatin condensation usually occurs after collapse, it appears likely that the ATP required for energy-consuming apoptosis-associated reactions is provided by anaerobic glycolysis and by any residual functioning mitochondria310-312. More extensive depletion of ATP311,312 or failure of caspase activation313 results in necrosis rather than apoptosis.

5.3.4. Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer In the aggregate, available evidence supports a central role for mitochondria in apoptosis in many cell types including hepatocytes. The critical impact of these organelles reflects disruption of electron transport, decreased intramitochondrial thiols167, collapse of possible release of procaspases and caspase activators, and failure of ATP generation314,315. As a corollary, these energetic considerations may clarify the unexpected association of apoptosis with cell cycle progression. That is, that apoptosis may be more likely to occur under conditions that favor cell cycle progression, whereas conditions that inhibit cell cycle progression may be This association is evident even in terminally protective306,316-327. differentiated neurons328,329, in which cell cycle progression does not lead to

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mitosis (see Chapter 15.4.2.: Wingless/Wnt-Like Signal Transduction and Effects of Lithium). These seemingly paradoxical relationships can be linked to identifiable signaling cascades330. Viewed in an energetic context, however, they may at least in part reflect the incompatibility of cell cycle-associated increases in ATP demand50, 331-334 with compromised mitochondrial ATP generating capacity. This incompatibility becomes unacceptable as the vulnerable cell is driven irreversibly beyond the G1 restriction point and committed to S phase entry. In S phase, already high expenditure of energy (ATP) increases, yet growth arrest is not possible before the G2 checkpoint. The fall in [ATP] that would be inevitable in an energetically compromised cell under such conditions would lead to an adenylate kinase-mediated increase in [AMP] and, therefore, to AMPK activation335. As discussed earlier (see Chapter 4: Fatty Acids, Modulation of Cell Growth, and the AMPKActivated Protein Kinase), AMPK among its other effects inhibits fatty acid synthesis and increases mitochondrial fatty acid oxidation; the latter predisposes to ROS generation and oxidative stress. While fatty acid oxidation also tends to suppress glycolysis, AMPK may simultaneously activate glycolysis and mitochondrial fatty acid oxidation in cardiac and skeletal muscle336. Under certain conditions, c-myc may also contribute to cell death by inhibiting activation337, or by suppressing p53-induced expression, thus furthering inappropriate cell cycle progression338. In the case of the retinoblastoma (Rb)-E2F pathway, inactivating phosphorylation of the critical Ser567 residue of Rb drives cell cycle progression; thus, it is directly related to cell cycle-associated apoptosis, and inversely related to cell cycle arrest and survival339. Such deregulation of Rb-E2F may also lead to increased expression of procaspase 7340. At this point it may be appropriate to sound a cautionary note concerning the interpretation of AMPK effects as studied with the cell permeant compound 5-aminoimidazole-4-carboxamide ribonucleoside (AICAR), a precursor of the AMP analog ZMP and activator of AMPK335. Extremely important information concerning the identity of AMPK substrates has been obtained through the use of this agent, which has thus proved to be a valuable investigative tool. In its experimental use, however, it may activate AMPK under conditions in which there has been no antecedent perturbation of cell energetics; that is, cellular [ATP] and [AMP] may be relatively normal, at least initially. Physiological activation of AMPK, in contrast, by definition signals the fact that cellular energetics are already compromised: [ATP] is low and/or [AMP] or [AMP]/[ATP] high. In some experiments, this distinction may not be of great consequence. However, in those studies in which experimental findings are critically related to, or influenced by, cellular or mitochondrial energy balance, results could be misleading. For


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example, effects of physiological activation of AMPK by an unfavorable change in cellular adenine nucleotide balance under conditions in which ATP demand is increasing, e.g., as the result of stress or cell cycle progression341,342, may differ from those that result from AMPK activation by AICAR under conditions in which [ATP] and ATP consumption are relatively normal and stable. In the former circumstance, programmed cell death would be far more likely to occur than in the latter. As a corollary, introduction of an antiproliferative agent (thereby preventing further deterioration of adenine nucleotide balance) would more likely exert a positive effect on cell survival in the face of high ATP demand than in the latter case, in which ATP demand is relatively stable. Such a caveat is also supported by recent evidence that, with or without AICAR, AMPK activation may be modulated independent of prior disturbance in adenine nucleotide balance (presumably including that introduced by the AICARderived AMP analog ZMP) by insulin343, leptin344, 345, and other factors346. Moreover, AICAR acting independently of AMPK may influence adipocyte glucose uptake347 and induce apoptosis348, 349. These considerations may be especially relevant to those instances in which AMPK effects may appear to be contradictory, e.g., simultaneous activation of glycolysis and fatty acid oxidation330-352 despite the generally reciprocal catabolic utilization of these substrates353,354. As noted above, however, concurrent activation of these processes by AMPK has been observed in cardiac and/or skeletal muscle336 and as a consequence of stimulation by adiponectin globular domain355. Induction of apoptosis caused by forced cell cycle progression of metabolically marginal cells would be expected to vary with in vitro conditions, and among cells in any given experiment. Accordingly, heterogeneity with respect to likelihood of successful cell cycle completion may account for conflicting reports of apoptosis in experimental hepatitis B (HBV) and C (HCV) infection, both of which promote cell cycle progression, largely attributable to effects of HBV X protein (HBx) or HCV core protein148,356-366. Among these effects is suppression of the Cip/Kip cell cycle inhibitors and/or by HCV core protein and/or HBx367370 . Thus, in a process of natural selection in microcosm, virus-infected cells, in which substrate and energy normally required for cell cycle support are diverted toward the superimposed demands of virus replication, may be more likely to become ATP-depleted and undergo (non-immune-mediated) programmed death. In contrast, replicatively and metabolically more competent cells, e.g., those with inactivating mutations of Apc, p53, or PTEN, would be more likely to survive; in vivo, comparable events may contribute to evolving carcinogenesis. The linkage between apoptosis and the cell cycle would also provide a rational basis for: 1) the rarity in human cancer (thus, implied cell survival

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value of the wild type gene) of inactivating mutations of the Cip/Kip family of cyclin E/cdk2 G1/S checkpoint inhibitors307; 2) Akt/PKB’s stabilization of 371 the Cip/Kip family member , and its phosphorylation-induced 371 372,373 cytoplasmic localization of both and ; and 3) in 304 306 apoptosis, caspase 3-mediated cleavage of both Akt/PKB and . Seemingly paradoxical Bcl-2 family effects on tumorigenesis, i.e., suppression by Bcl-2374 and acceleration by Bax375, could thus reflect a decrease or increase, respectively, in proliferative drive, in part compensatory in response to loss of functioning hepatocytes (see Chapter 8: Metabolism and Gene Expression in Liver Regeneration). Finally, the cell cycle-apoptosis linkage may provide an opportunity for a novel approach to cancer treatment based on metabolic manipulation. In such an approach, cells in the tumor initially would be brought to cell cycle arrest, then induced to near synchronous cell cycle re-entry, thus incurring obligatory increases in ATP demand. It is hypothesized that, under such conditions, pharmacological disruption of mitochondrial energetics and/or glycolysis soon after the G1/S transition would lead inexorably to progressive ATP depletion and cell death. In contrast, non-transfomed cells, being subject to usual growth regulatory constraints, would be less susceptible to, and less likely to incur the consequences of, such artificially-induced cell cycle progression.

5.4. REFERENCES 1. S. Green, PPAR: a mediator of peroxisome proliferator action, Mutat Res 333:101-109 (1995). 2. K. Schoonjans, B. Staels, and J. Auwerx, Role of the peroxisome proliferator-activated receptor (PPAR) in mediating the effects of fibrates and fatty acids on gene expression, J Lipid Res 37:907-925 (1996). 3. T. Willson, and W. Wahli, Peroxisome proliferator-activated receptor agonists, Curr Opin Chem Biol 1:235-241 (1997). 4. F. Gonzalez, J. Peters, and R. Cattley, Mechanism of action of the nongenotoxic peroxisome proliferators: role of the peroxisome proliferator-activator receptor alpha, J Natl Cancer Inst 90:1702-1709 (1998). 5. B. Desvergne, and W. Wahli, proliferator-activated, receptors, nuclear control of metabolism., Endocrine Rev 20:649-688 (1999). 6. P. Eacho, and P. Foxworthy, Inhibition of hepatic fatty acid oxidation by bezafibrate and bezafibroyl CoA., Biochem Biophys Res Commun 157:1148-1153 (1988). 7. P. Foxworthy, and P. Eacho, Inhibition of hepatic fatty acid oxidation at carnitine palmitoyltransferase I by the peroxisome proliferator 2-hydroxy-3-propyl-4-[6-(tetrazol-5yl) hexyloxy]acetophenone., Biochem J 252:409-414 (1988). 8. S. Lee, T. Pineau, J. Drago, et al., Targeted disruption of the alpha isoform of the peroxisome proliferator-activated receptor gene in mice results in abolishment of the pleiotropic effects of peroxisome proliferators, Molec Cell Biol 15:3012-3022 (1995).


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9. H. Castelein, T. Gulick, P. Declercq, G. Mannaerts, D. Moore, and M. Baes, The peroxisome proliferator activated receptor regulates malic enzyme gene expression., J Biol Chem 269:26754-26758 (1994). 10. R. Kaikaus, W. Chan, N. Lysenko, R. Ray, P. Ortiz de Montellano, and N. Bass, Induction of peroxisomal fatty acid beta-oxidation and liver fatty acid binding protein by peroxisome proliferators: Mediation via the cytochrome P-450 4A1 omega-hydroxylase pathway, J Biol Chem 268:9592-9603 (1993A). 11. R. Kaikaus, Z. Sui, N. Lysenko, et al., Regulation of pathways of extramitochondrial fatty acid oxidation and liver fatty acid-binding protein by long-chain monocarboxylic fatty acids in hepatocytes: effect of inhibition of carnitine palmitoyltransferase I, J Biol Chem 268:26866-26871 (1993B). 12. J. Hardwick, B. Song, E. Huberman, and F. Gonzalez, Isolation, complementary DNA sequence, and regulation of rat hepatic lauric acid omega-hydroxylase (cytochrome P-450 LAw): Identification of a new cytochrome P-450 gene family, J Biol Chem 262:801-810 (1987). 13. J. Rininger, T. Goldsworthy, and J. Babish, Time course comparison of cell-cycle protein expression following partial hepatectomy and WY14,643-induced hepatic cell proliferation in F344 rats, Carcinogenesis 18:935-941 (1997). 14. J. Reddy, M. Rao, D. Azarnoff, and S. Sell, Mitogenic and carcinogenic effect of hypolipidemic peroxisome proliferators [4-chloro-6(2,3-xylidino)-2-pyrimidylthio]acetic acid (Wy-14,643) in rat and mouse liver, Cancer Res 39:152-161 (1979). 15. P. Eacho, R. Lanier, and C. Brodhecker, Hepatocellular DNA synthesis in rats given peroxisome proliferating agents: comparison of WY-14,643 to clofibric acid, nafenopin and LY171883., Carcinogenesis 12:1557-1561 (1991). 16. D. Marsman, T. Goldsworthy, and J. Popp, Contrasting hepatocytic peroxisome proliferation, lipofuscin accumulation and cell turnover for the hepatocarcinogens Wy14,643 and clofibric acid, Carcinogenesis 13:1011-1017 (1992). 17. N. Barrass, R. Price, B. Lake, and T. Orton, Comparison of the acute and chronic mitogenic effects of the peroxisome proliferators methylclofenapate and clofibiric acid in rat liver., Carcinogenesis 14:1451-1456 (1993). 18. T. Ohmura, G. Ledda-Columbano, R. Piga, et al., Hepatocyte proliferation induced by a single dose of a peroxisome proliferator, Am J Pathol 148:815-824 (1996). 19. M. Menegazzi, P. Carcereri-De, A, H. Suzuki, et al., Liver cell proliferation induced by nafenopin and cyproterone acetate is not associated with increases in activation of transcription factors NF-kappaB and AP-1 or with expression of tumor necrosis factor alpha, Hepatology 25:585-592 (1997). 20. T. Ohmura, S. Katyal, J. Locker, G. Ledda-Columbano, and H. Shinozuka, Induction of cellular DNA synthesis in the pancreas and kidneys of rats by peroxisome proliferators, 9cis retinoic acid, and 3,3’,5-triiodo-L-thyronine, Cancer Res 57:795-798 (1997). 21. H. Rabes, R. Wirsching, H.-V. Tuczek, and G. Iseler, Analysis of cell cycle compartments of hepatocytes after partial hepatectomy, Cell Tissue Kinet 9:517-532 (1976). 22. I. Rusyn, H. Tsukamoto, and R. Thurman, WY-14 643 rapidly activates nuclear factor kappaB in Kupffer cells before hepatocytes, Carcinogenesis 19:1217-1222 (1998). 23. M. Rose, C. Rivera, B. Bradford, et al., Kupffer cell oxidant production is central to the mechanism of peroxisome proliferators, Carcinogenesis 20:27-33 (1999). 24. I. Rusyn, C. Bradham, L. Cohn, et al., Corn oil rapidly activates nuclear factor-kappaB in hepatic Kupffer cells by oxidant-dependent mechanisms, Carcinogenesis 20:2095-2100 (1999).

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25. J. Peters, I. Rusyn, M. Rose, F. Gonzalez, and R. Thurman, Peroxisome proliferatoractivated receptor alpha is restricted to hepatic parenchymal cells not Kupffer cells: implications for the mechanism of action of peroxisome proliferators in hepatocarcinogenesis, Carcinogenesis 21:823-826 (2000A). 26. W. Parzefall, W. Berger, E. Kainzbauer, O. Teufelhofer, R. Schulte-Hermann, and R. Thurman, Peroxisome proliferators do not increase DNA synthesis in purified rat hepatocytes, Carcinogenesis 22:519-523 (2001). 27. J. Lawrence, G. Wollenberg, and J. DeLuca, Tumor necrosis factor alpha is not required for WY14,643-induced cell proliferation, Carcinogenesis 22:381-386 (2001). 28. S. Anderson, C. Dunn, R. Cattley, and J. Corton, Hepatocellular proliferation in response to a peroxisome proliferator does not require TNFalpha signaling., Carcinogenesis. 22:1843-1851 (2001). 29. J. Peters, R. Cattley, and F. Gonzalez, Role of PPAR alpha in the mechanism of action of the nongenotoxic carcinogen and peroxisome proliferator Wy-14,643, Carcinogenesis 18:2029-2033 (1997). 30. S. P. Anderson, L. Yoon, E. B. Richard, C. S. Dunn, R. C. Cattley, and J. C. Corton, Delayed liver regeneration in peroxisome proliferator-activated receptor-alpha-null mice, Hepatology 36:544-54 (2002). 31. F. J. Gonzalez, The peroxisome proliferator-activated receptor alpha (PPARalpha): role in hepatocarcinogenesis, Mol Cell Endocrinol 193:71-9 (2002). 32. J. Gill, N. James, R. Roberts, and C. Dive, The non-genotoxic hepatocarcinogen nafenopin suppresses rodent hepatocyte apoptosis induced by TGFbeta1, DNA damage and Fas, Carcinogenesis 19:299-304 (1998). 33. B. Forman, J. Chen, and R. Evans, Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-activated receptors alpha and delta., Proc Natl Acad Sci USA 94:4312-4317 (1997). 34. S. Kliewer, S. Sundseth, S. Jones, et al., Fatty acids and eicosanoids regulate gene expression through direct interactions with peroxisome proliferator-activated receptors alpha and gamma, Proc Natl Acad Sci USA 94:4318-4323 (1997). 35. P. Devchand, H. Keller, J. Peters, M. Vazquez, F. Gonzalez, and W. Wahli, The PPARalpha-leukotriene B4 pathway to inflammation control., Nature 384:39-43 (1996). 36. E. Lock, A. Mitchell, and C. Elcombe, Biochemical mechanisms of induction of hepatic peroxisome proliferation, Annu Rev Pharmacol Toxicol 29:145-163 (1989). 37. B. Lake, Mechanisms of hepatocarcinogenicity of peroxisome-proliferating drugs and chemicals, Annu Rev Pharmacol Toxicol 35:483-507 (1995). 38. C. Soontjens, J. Rafter, and J.-Å. Gustafsson, Ligands for orphan receptors?, J Endocrinol 150:S241-S257 (1996). 39. C. Neat, M. Thomassen, and H. Osmundsen, Induction of peroxisomal beta-oxidation in rat liver by high-fat diets, Biochem J 186:369-371 (1980). 40. P. Mortensen, Dicarboxylic acids and the lipid metabolism, Danish Med Bull 31:121-145 (1984). 41. H. Osmundsen, J. Bremer, and J. Pedersen, Metabolic aspects of persoxisomal betaoxidation, Biochim Biophys Acta 1085:141-158 (1991). 42. D. Kroetz, P. Yook, P. Costet, P. Bianchi, and T. Pineau, Peroxisome proliferatoractivated receptor alpha controls the hepatic CYP4A induction adaptive response to starvation and diabetes, J Biol Chem 273:31581-31589 (1998). 43. N. Bass, R. Appel, E. Goetzl, A. Dannenberg, and A. Nanji, Peroxisome proliferator activated receptor-alpha-mediated gene expression and adaptation to fatty acid overload in alcoholic liver disease., Alcoholism Clin Exp Res 22:749-750 (1998).


101

44. C. Wolfrum, C. Borrmann, T. Börchers, and F. Spener, Fatty acids and hypolipidemic drugs regulate peroxisome proliferator-activated receptors a- and g-mediated gene expression via liver fatty acid binding protein: A signaling path to the nucleus, Proc Natl Acad Sci USA 98:2323-2328 (2001). 45. N. S. Tan, N. S. Shaw, N. Vinckenbosch, et al., Selective cooperation between fatty acid binding proteins and peroxisome proliferator-activated receptors in regulating transcription, Mol Cell Biol 22:5114-27 (2002). 46. C. Blanquart, O. Barbier, J. C. Fruchart, B. Staels, and C. Glineur, Peroxisome proliferator-activated receptor alpha (PPARalpha) turnover by the ubiquitin-proteasome system controls the ligand-induced expression level of its target genes, J Biol Chem 277:37254-9 (2002). 47. A. Shalev, C. Siegrist-Kaiser, P. Yen, et al., The peroxisome proliferator-activated receptor alpha is a phosphoprotein: regulation by insulin, Endocrinology 137:4499-4502 (1996). 48. C. Juge-Aubry, E, Hammar, C. Siegrist-Kaiser, et al., Regulation of the transcriptional activity of the peroxisome proliferator-activated receptor alpha by phosphorylation of a ligand-independent trans-activating domain, J Biol Chem 274:10505-10510 (1999). 49. B. Neve, J.-C. Fruchart, and B. Staels, Role of the peroxisome proliferator-activated receptors (PPAR) in atherosclerosis, Biochem Pharmacol 60:1245-1250 (2000). 50. C. Van den Bogert, P. Muus, C. Haanen, A. Pennings, T. Melis, and A. Kroon, Mitochondrial biogenesis and mitochondrial activity during the progression of the cell cycle of human leukemic cells, Exp Cell Res 178:143-153 (1988). 51. C. Prager, H. Schön, M. Nikfardjam, et al., Clofibrate elevates enzyme activities of the tricarboxylic acid cycle in rat liver, J Lipid Res 34:359-364 (1993). 52. S. Narravula, and S. Colgan, Hypoxia-inducible factor 1-mediated inhibition of peroxisome proliferator-activated receptor alpha expression during hypoxia, J Immunol 166:7543-7548 (2001). 53. B. Wolfe, J. Kane, R. Havel, and H. Brewster, Mechanism of the hypolipemic effect of clofibrate in postabsorptive man, J Clin Invest 52:2146-2159 (1973). 54. M. Laker, and P. Mayes, The immediate and long term effects of clofibrate on the metabolism of the perfused rat liver, Biochem Pharmacol 28:2813-2827 (1979). 55. K. Yamamoto, N. Fukuda, L. Zhang, and T. Sakai, Altered hepatic metabolism of fatty acids in rats fed a hypolipidaemic drug, fenofibrate, Pharmacol Res 33:337-342 (1996). 56. R. Ockner, N. Lysenko, N. Wu, and N. Bass, Hepatocyte growth inhibitors modulate mitochondrial and extramitochondrial fatty acid oxidation, Hepatology 24:253A (Abstract) (1996). 57. L. Gustafson, F. Kuipers, C. Wiegman, H. Sauerwein, J. Romijn, and A. Meijer, Clofibrate improves glucose tolerance in fat-fed rats but decreases hepatic glucose consumption capacity, J Hepatol 37:425-431 (2002). 58. M. Poynter, and R. Daynes, Peroxisome proliferator-activated receptor alpha activation modulates cellular redox status, represses nuclear factor-kappaB signaling, and reduces inflammatory cytokine production in aging, J Biol Chem 273:32833-32841 (1998). 59. N. Marx, G. Sukhova, T. Collins, P. Libby, and J. Plutzky, PPARalpha activators inhibit cytokine-induced vascular cell adhesion molecule-1 expression in human endothelial cells, Circulation 99:3125-3131 (1999). 60. H. Duez, Y. S. Chao, M. Hernandez, et al., Reduction of atherosclerosis by the peroxisome proliferator-activated receptor alpha agonist fenofibrate in mice, J Biol Chem 277:48051-7 (2002).

102

Chapter 5

61. G. A. Francis, J. S. Annicotte, and J. Auwerx, PPAR-alpha effects on the heart and other vascular tissues, Am J Physiol Heart Circ Physiol 285:H1-9 (2003). 62. P. Delerive, P. Gervois, J.-C. Fruchart, and B. Staels, Induction of IkBa expression as a mechanism contributing to the anti-inflammatory activities of peroxisome proliferatoractivated receptor-alpha activators., J Biol Chem 275:36703-36707 (2000B). 63. G. Chinetti, S. Griglio, M. Antonucci, et at.. Activation of proliferator-activated receptors alpha and gamma induces apoptosis of human monocyte-derived macropages., J Biol Chem 273:25573-25580 (1998). 64. P. Delerive, C. Furman, E. Teissier, J.-C. Fruchart, P. Duriez, and B. Staels, Oxidized phosopholipids activate PPARa in a phospholipase A2-dependent manner., FEBS Letters 471:34-38 (2000A). 65. P. Delerive, B. De, K, S. Besnard, et al., Peroxisome proliferated-activated receptor alpha negatively regulates the vascular inflammatory gene response by negative cross-talk with transcription factors NF-kappaB and AP-1., J Biol Chem 274:32048-32054 (1999). 66. R. De Caterina, J. Liao, and P. Libby, Fatty acid modulation of endothelial activation., Am J Clin Nutr 71:213S-23S (2000). 67. B. Hennig, P. Meerarani, P. Ramadass, B. Watkins, and M. Toborek, Fatty acid-mediated activation of vascular endothelial cells, Metab Clin Exper 49:1006-1013 (2000). 68. C. Glass, Potential roles of the peroxisome proliferator-activated receptor-gamma in macrophage biology and atherosclerosis, J Endocrinol 169:461-464 (2001). 69. P. Libby, P. Ridker, and A. Maseri, Inflammation and atherosclerosis, Circulation 105:1135-1143 (2002). 70. D. Arsenijevic, H. Onuma, C. Pecqueur, et al., Disruption of the uncoupling protein-2 gene in mice reveals a role in immunity and reactive oxygen species production, Nat Genet 26:435-9 (2000). 71. L. Makowski, J. Boord, K. Maeda, et al., Lack of macrophage fatty-acid-binding protein aP2 protects mice deficient in apolipoprotein E against atherosclerosis, Nature Med 7:699705 (2001). 72. M. Perrella, A. Pellacani, M. Layne, et al., Absence of adipocyte fatty acid binding protein prevents the development of accelerated atherosclerosis in hypercholesterolemic mice, FASEB J 15:1774-1776 (2001). 73. D. C. Jones, X. Ding, and R. A. Daynes, Nuclear receptor peroxisome proliferatoractivated receptor alpha (PPARalpha) is expressed in resting murine lymphocytes. The PPARalpha in T and B lymphocytes is both transactivation and transrepression competent, J Biol Chem 277:6838-45 (2002). 74. G. Boden, and G. I. Shulman, Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction, Eur J Clin Invest 32 Suppl 3:14-23 (2002). 75. S. Mills, D. Foster, and J. Mcgarry, Interaction of malonyl-CoA and related compounds with mitochondria from different rat tissues: Relationship between ligand binding and inhibition of carnitine palmitoyltransferase I, Biochem J 214:83-91 (1983). 76. L. Drynan, P. Quant, and V. Zammit, Flux control exerted by mitochondrial outer membrane carnitine palmitoyltransferase over beta-oxidation, ketogenesis and tricarboxylic acid cycle activity in hepatocytes isolated from rats in different metabolic states., Biochem J 317:791-795 (1996A). 77. L. Drynan, P. Quant, and V. Zammit, The role of changes in the sensitivity of hepatic mitochondrial overt carnitine palmitoyltransferase in determining the onset of the ketosis of starvation in the rat., Biochem J 318:767-770 (1996B).


103

78. J. Sleboda, K. Risan, O. Spydevold, and J. Bremer, Short-term regulation of carnitine palmitoyltransferase I in cultured rat hepatocytes: spontaneous inactivation and reactivation by fatty acids, Biochim Biophys Acta 1436:541-549 (1999). 79. J. McGarry, and N. Brown, Reconstitution of purified, active and malonyl-CoA-sensitive rat liver carnitine palmitoyltransferase I: relationship between membrane environment and malonyl-CoA sensitivity, Biochem J 349:179-187 (2000). 80. F. Fraser, C. Corstorphine, and V. Zammit, Topology of carnitine palmitoyltransferase I in the mitochondrial outer membrane., Biochem J 323:711-718 (1997). 81. F. Fraser, R. Padovese, and V. Zammit, Distinct kinetics of carnitine palmitoyltransferase I in contact sites and outer membranes of rat liver mitochondria., J Biol Chem 276:2018220185(2001). 82. E. Lenartowicz, and M. Olson, The inhibition of alpha-ketoglutarate oxidation by fatty acids in rat liver mitochondria, J Biol Chem 253:5990-5996 (1978). 83. A. Nulton-Persson, and L. Szweda, Modulation of mitochondrial function by hydrogen peroxide, J Biol Chem 276:23357-61 (2001). 84. X. Chen, N. Iqbal, and G. Boden, The effects of free fatty acids on gluconeogenesis and glycogenolysis in normal subjects., J Clin Invest 103:365-372 (1999). 85. A. Dresner, D. Laurent, M. Marcucci, et al., Effects of free fatty acids on glucose transport and IRS-1-associated phosphatidylinositol 3-kinase activity., J Clin Invest 103:253-259 (1999). 86. K. Cusi, K. Maezono, A. Osman, et al., Insulin resistance differentially affects the PI 3kinase- and MAP kinase-mediated signaling in human muscle., J Clin Invest 105:311-320 (2000). 87. H. Steinberg, G. Paradisi, G. Hook, K. Crowder, J. Cronin, and A. Baron, Free fatty acid elevation impairs insulin-mediated vasodilation and nitric oxide production, Diabetes 49:1231-1238(2000). 88. P. Arner, Insulin resistance in type 2 diabetes: role of fatty acids, Diabetes Metab Res Rev 18 Suppl 2:S5-9 (2002). 89. Y. Kruszynska, D. Worrall, J. Ofrecio, J. Frias, G. Macaraeg, and J. Olefsky, Fatty acidinduced insulin resistance: decreased muscle PI3K activation but unchanged Akt phosphorylation, J Clin Endocrinol Metab 87:226-234 (2002). 90. J. McGarry, Banting lecture, 2001: dysregulation of fatty acid metabolism in the etiology of type 2 diabetes, Diabetes 51:7-18 (2002). 91. C. Yu, Y. Chen, G. W. Cline, et al., Mechanism by which fatty acids inhibit insulin activation of insulin receptor substrate-1 (IRS-1)-associated phosphatidylinositol 3-kinase activity in muscle, J Biol Chem 277:50230-50236 (2002). 92. V. Skulachev, Fatty acid circuit as a physiological mechanism of uncoupling of oxidative phosphorylation, FEBS Lett 294:158-162 (1991). 93. L. Wojtczak, and P. Schönfeld, Effect of fatty acids on energy coupling processes in mitochondria, Biochim Biophys Acta 1183:41-57 (1993). 94. P. Schönfeld, M. Wiêckowski, and L. Wojtczak, Thyroid hormone-induced expression of the ADP/ATP carrier and its effect on fatty acid-induced uncoupling of oxidative phosphorylation, FEBS Lett 416:19-22 (1997A). 95. P. Schönfeld, and R. Bohnensack, Fatty acid-promoted mitochondrial permeability transition by membrane depolarization and binding to the ADP/ATP carrier, FEBS Lett 420:167-170(1997B). 96. L. Wojtczak, and M. Wieckowski, The mechanisms of fatty acid-induced proton permeability of the inner mitochondrial membrane, J Bioenerg Biomembranes 31:447-455 (1999A).

104

Chapter 5

97. R. Hertz, and J. Bar-Tana, Peroxisome proliferator-activated receptor (PPAR) alpha activation and its consequences in humans, Toxicol Lett 102-103:85-90 (1998). 98. C. Palmer, M. Hsu, K. Griffin, J. Raucy, and E. Johnson, Peroxisome proliferator activated receptor-alpha expression in human liver, Molec Pharmacol 53:14-22 (1998). 99. N. Woodyatt, K. Lambe, K. Myers, J. Tugwood, and R. Roberts, The peroxisome proliferator (PP) response element upstream of the human acyl CoA oxidase gene is inactive among a sample human population: significance for species differences in response to PPs, Carcinogenesis 20:369-372 (1999). 100. D. Auboeuf, J. Rieusset, L. Fajas, et al., Tissue distribution and quantification of the expression of mRNAs of peroxisome proliferator-activated receptors and liver X receptoralpha in humans: no alteration in adipose tissue of obese and NIDDM patients., Diabetes 46:1319-1327 (1997). 101. B. Staels, J. Dallongeville, J. Auwerx, K. Schoonjans, and J. Fruchart, Mechanism of action of fibrates on lipid and lipoprotein metabolism, Circulation 98:2088-2093 (1998). 102. C. Wolfrum, C. Buhlmann, B. Rolf, T. Börchers, and F. Spener, Variation of liver-type fatty acid binding protein content in the human hepatoma cell line HepG2 by peroxisome proliferators and antisense RNA affects the rate of fatty acid uptake, Biochim Biophys Acta 1437:194-201 (1999). 103. R. Cunard, M. Ricote, D. DiCampli, et al., Regulation of cytokine expression by ligands of peroxisome proliferator activated receptors., J Immunol 168:2795-2802 (2002). 104. M. Ricote, A. Li, T. Willson, C. Kelly, and C. Glass, The peroxisome proliferatoractivated receptor-gamma is a negative regulator of macrophage activation, Nature 391:79-82 (1998). 105. A. Wächtershäuser, S. Loitsch, and J. Stein, PPAR-gamma is selectively upregulated in Caco-2 cells by butyrate, Biochem Biophys Res Comm 272:380-385 (2000). 106. U. Kintscher, S. Wakino, D. Bruemmer, et al., TGF-beta(1) induces peroxisome proliferator-activated receptor gamma1 and gamma2 expression in human THP-1 monocytes, Biochem Biophys Res Commun 297:794-9 (2002). 107. J. Lin, P. T. Tarr, R. Yang, et al., PGC-1beta in the regulation of hepatic glucose and energy metabolism, J Biol Chem 278:30843-8 (2003). 108. J. C. Yoon, G. Xu, J. T. Deeney, et al., Suppression of beta cell energy metabolism and insulin release by PGC-1alpha, Dev Cell 5:73-83 (2003). 109. K. Subbaramaiah, D. Lin, J. Hart, and A. Dannenberg, Peroxisome proliferator-activated receptor gamma ligands suppress the transcriptional activation of cyclooxygenase-2, J Biol Chem 276:12440-12448 (2001). 110. B. Hinz, K. Brune, and A. Pahl, 15-Deoxy-delta(12,14)-prostaglandin J2 inhibits the expression of proinflammatory genes in human blood monocytes via a PPAR-gammaindependent mechanism, Biochem Biophys Res Commun 302:415-20 (2003). 111. P. Tontonoz, S. Singer, B. Forman, et al., Terminal differentiation of human liposarcoma cells induced by ligands for peroxisome proliferator-activated receptor gamma and the retinoid X receptor, Proc Natl Acad Sci USA 94:237-241 (1997). 112. T. Kubota, K. Koshizuka, E. Williamson, et al., Ligand for peroxisome proliferatoractivated receptor gamma (troglitazone) has potent antitumor effect against human prostate cancer both in vitro and in vivo, Cancer Res 58:3344-3352 (1998). 113. E. Mueller, P. Sarraf, P. Tontonoz, et al., Terminal differentiation of human breast cancer through PPAR gamma, Mol Cell 1:465-470 (1998). 114. E. Mueller, M. Smith, P. Sarraf, et al., Effects of ligand activation of peroxisome proliferator-activated receptor gamma in human prostate cancer, Proc Natl Acad Sci USA 97:10990-10995 (2000).


105

115. J. Brockman, R. Gupta, and R. Dubois, Activation of PPARgamma leads to inhibition of anchorage-independent growth of human colorectal cancer cells., Gastroenterology 115:1049-1055 (1998). 116. P. Sarraf, E. Mueller, D. Jones, et al., Differentiation and reversal of malignant changes in colon cancer through PPARgamma, Nature Med 4:1046-1052 (1998). 117. H. J. Burstein, G. D. Demetri, E. Mueller, P. Sarraf, B. M. Spiegelman, and E. P. Winer, Use of the peroxisome proliferator-activated receptor (PPAR) gamma ligand troglitazone as treatment for refractory breast cancer: a phase II study, Breast Cancer Res Treat 79:391-7 (2003). 118. S. Wakino, U. Kintscher, Z. Liu, et al., Peroxisome proliferator-activated receptor gamma ligands inhibit retinoblastoma phosphorylation and G1->S transition in vasculare smooth muscle cells, J Biol Chem 275:22435-22441 (2000). 119. C. Clay, G. Atsumi, K. High, and F. Chilton, Early de novo gene expression is required for 15-deoxy-delta-12,14-prostaglandin J2-induced apoptosis in breast cancer cells., J Biol Chem 276:47131-47135 (2001). 120. D. Panigrahy, S. Singer, L. Q. Shen, et al., PPARgamma ligands inhibit primary tumor growth and metastasis by inhibiting angiogenesis, J Clin Invest 110:923-32 (2002). 121. E. Saez, P. Tontonoz, M. Nelson, et al., Activators of the nuclear receptor PPARgamma enhance colon polyp formation, Nature Med 4:1058-1061 (1998). 122. A.-M. Lefebvre, I. Chen, P. Desreumaux, et al., Activation of the peroxisome proliferator-activated receptor gamma promotes the development of colon tumors in C57BL/6J-APC Min/+ mice, Nature Med 4:1053-1057 (1998). 123. E. Hu, J. Kim, P. Sarraf, and B. Spiegelman, Inhibition of adipogenesis through MAP kinase-mediated phosphorylation of PPAR gamma, Science 274:2100-2103 (1996). 124. M. Adams, M. Reginato, D. Shao, M. Lazar, and V. Chatterjee, Transcriptional activation by peroxisome proliferator-activated receptor gamma is inhibited by phosphorylation at a consensus mitogen-activated protein kinase site., J Biol Chem 272:5128-5132 (1997). 125. G. D. Girnun, W. M. Smith, S. Drori, et al., APC-dependent suppression of colon carcinogenesis by PPARgamma, Proc Natl Acad Sci U S A 99:13771-6 (2002). 126. W. Yang, and H. Frucht, Activation of the PPAR pathway induces apoptosis and COX-2 inhibition in HT-29 human colon cancer cells, Carcinogenesis 22:1379-1383 (2001). 127. M. Wang, S. Wise, T. Leff, and T. Su, Troglitazone, an antidiabetic agent, inhibits cholesterol biosynthesis through a mechanism independent of peroxisome proliferatoractivated receptor-gamma, Diabetes 48:254-260 (1999). 128. Y. Hattori, K. Akimoto, and K. Kasai, The effects of thiazolidinediones on vascular smooth muscle cell activation by angiotensin II, Biochem Biophys Res Commun 273:1144-1149 (2000). 129. A. Chawla, Y. Barak, L. Nagy, D. Liao, P. Tontonoz, and R. Evans, PPAR-gamma dependent and independent effects on macrophage-gene expression in lipid metabolism and inflammation., Nat Med 7:48-52 (2001). 130. S. Palakurthi, H. Aktas, L. Grubissich, R. Mortensen, and J. Halperin, Anticancer effects of thiazolidinediones are independent of peroxisome proliferator-activated receptor gamma and mediated by inhibition of translation initiation, Cancer Res 61:6213-6218 (2001). 131. A. Abe, Y. Kiriyama, M. Hirano, et al., Troglitazone suppresses cell growth of KU812 cells independently of PPARgamma., Eur J Pharmacol 436:7-13 (2002).

106

Chapter 5

132. L. Bell-Parikh, T. Ide, J. Lawson, P. McNamara, M. Reilly, and G. FitzGerald, Biosynthesis of 15-deoxy-delta-12,14-PGJ2 andd the ligation of PPAR-gamma., J Clin Invest 112:945-955 (2003). 133. I. Inoue, S. Katayama, K. Takahashi, et al., Troglitazone has a scavenging effect on reactive oxygen species, Biochem Biophys Res Commun 235:113-116 (1997). 134. R. Caricchio, D. Kovalenko, W. Kaufmann, and P. Cohen, Apoptosis provoked by the oxidative stress inducer menadione (Vitamin K3) is mediated by the Fas/Fas ligand system., Clin Immunol 93:65-74 (1999). 135. T. He, T. Chan, B. Vogelstein, and K. Kinzler, PPARdelta is an APC-regulated target of nonsteroidal anti-inflammatory drugs, Cell 99:335-345 (1999). 136. R. Gupta, J. Tan, W. Krause, et al., Prostacyclin-mediated activation of peroxisome proliferator-activated receptor in colorectal cancer, Proc Natl Acad Sci USA 97:1327513280(2000). 137. C. Jehl-Pietri, C. Bastie, I. Gillot, S. Luquet, and P. Grimaldi, Peroxisome-proliferatoractivated receptor delta mediates the effects of long-chain fatty acids on post-confluent cell proliferation, Biochem J 350:93-98 (2000). 138. J. Peters, S. Lee, W. Li, et al., Growth, adipose, brain, and skin alterations resulting from targeted disruption of the mouse peroxisome proliferator-activated receptor beta(delta), Molec Cell Biol 20:5119-5128 (2000B). 139. B. Tong, J. Tan, L. Tajeda, et al., Heightened expression of cyclooxygenase-2 and peroxiome proliferator-activated receptor-delta in human endometrial adenocarcinoma, Neoplasia 2:483-490 (2000). 140. C. Bastie, S. Luquet, D. Holst, C. Jehl-Pietri, and P. Grimaldi, Alterations of peroxisome proliferator-activated receptor delta activity affect fatty acid-controlled adipose differentiation., J Biol Chem 275:38768-38773 (2000). 141. H. Vosper, L. Patel, T. Graham, et al., The peroxisome proliferator-activated receptor delta promotes lipid accumulation in human macrophages, J Biol Chem 276:44258-44265 (2001). 142. Y. Shi, M. Hon, and R. Evans, The peroxisome proliferator-activated receptor delta, an integrator of transcriptional repression and nuclear receptor signaling, Proc Natl Acad Sci USA 99:2613-2618 (2002). 143. Y. Barak, D. Liao, W. He, et al., Effects of peroxisome proliferator-activated receptor delta on placentation, adiposity, and colorectal cancer., Proc Natl Acad Sci USA 99:303308 (2002). 144. R. Schulte-Hermann, W. Bursch, and B. Grasl-Kraupp, Active cell death (apoptosis) in liver biology and disease, Prog Liver Dis 13:1 -36 (1995). 145. N. Zamzami, and G. Kroemer, The mitochondrion in apoptosis: how Pandora’s box opens, Nature Rev Molec Cell Biol 2:67-71 (2001). 146. E. Bossy-Wetzel, and D. Green, Apoptosis: checkpoint at the mitochondrial frontier., Mutation Res 434:243-251 (1999). 147. M.-T. Heemels, and Ed, Nature Insight: Apoptosis, Nature 407:769-816 (2000). 148. R. Ockner, Apoptosis and liver diseases: Recent concepts of mechanisms and significance, J Gastroenterol Hepatol 16:248-260 (2001). 149. S. Orrenius, B. Zhivotovsky, and P. Nicotera, Regulation of cell death: the calciumapoptosis link, Nat Rev Mol Cell Biol 4:552-65 (2003). 150. O. Micheau, and J. Tschopp, Induction of TNF receptor I-mediated apoptosis via two sequential signaling complexes, Cell 114:181-90 (2003). 151. A. Beg, and D. Baltimore, An essential role for NF-kappaB in preventing TNF-alphainduced cell death., Science 274:782-784 (1996).


107

152. D. Van Antwerp, S. Martin, T. Kafri, D. Green, and I. Verma, Suppression of TNFalpha-induced apoptosis by NF-kappaB, Science 274:787-789 (1996). 153. C.-Y. Wang, M. Mayo, and J. Baldwin, AS TNF- and cancer therapy-induced apoptosis: potentiation by inhibition of NF-kappaB, Science 274:784-787 (1996). 154. O. Ozes, L. Mayo, J. Gustin, S. Pfeffer, L. Pfeffer, and D. Donner, NF-kappaB activation by tumour necrosis factor requires the Akt serine-threonine kinase, Nature 401:82-85 (1999). 155. M. E. Griffin, M. J. Marcucci, G. W. Cline, et al., Free fatty acid-induced insulin resistance is associated with activation of protein kinase C theta and alterations in the insulin signaling cascade, Diabetes 48:1270-4 (1999). 156. T. K. Lam, H. Yoshii, C. A. Haber, et al., Free fatty acid-induced hepatic insulin resistance: a potential role for protein kinase C-delta, Am J Physiol Endocrinol Metab 283:E682-91 (2002). 157. C. L. Soltys, L. Buchholz, M. Gandhi, A. S. Clanachan, K. Walsh, and J. R. Dyck, Phosphorylation of cardiac protein kinase B is regulated by palmitate, Am J Physiol Heart Circ Physiol 283:H1056-64 (2002). 158. R. Budd, Death receptors couple to both cell proliferation and apoptosis., J Clin Invest 109:437-441 (2002). 159. A. Gross, X. Yin, K. Wang, et al., Caspase cleaved BID targets mitochondria and is required for cytochrome c release, while BCL-XL prevents this release but not tumor necrosis factor-R1/Fas death, J Biol Chem 274:1156-63 (1999). 160. D. Kirsch, A. Doseff, B. Chau, et al., Caspase-3-dependent cleavage of Bcl-2 promotes release of cytochrome c, J Biol Chem 274:21155-21161 (1999). 161. B. Beltran, M. Quintero, E. Garcia-Zaragoza, E. O’Connor, J. V. Esplugues, and S. Moncada, Inhibition of mitochondrial respiration by endogenous nitric oxide: a critical step in Fas signaling, Proc Natl Acad Sci U S A 99:8892-7 (2002). 162. D. Green, and G. Kroemer, The central executioners of apoptosis: caspases or mitochondria?, Trends Cell Biol 8:267-271 (1998). 163. W. Ding, H. Shen, and C. Ong, Critical role of reactive oxygen species and mitochondrial permability transition in microcystin-induced rapid apoptosis in rat hepatocytes., Hepatol 32:547-555 (2000). 164. A. Khaled, D. Reynolds, H. Young, C. Thompson, K. Muegge, and S. Durum, Interleukin-3 withdrawal induces an early increase in mitochondrial membrane potential unrelated to the Bcl-2 family Roles of intracellular pH ADP transport and F(0)F(1)ATPase, J Biol Chem 276:6453-6462 (2001). 165. P. Bernardi, The permeability transition pore. Control points of a cyclosporin A-sensitive mitochondrial channel involved in cell death., Biochim Biophys Acta 1996:1275 (1996). 166. D. Metivier, B. Dallaporta, N. Zamzami, et al., Cytofluorometric detection of mitochondrial alterations in early CD95/Fas/APO-1-triggered apoptosis of Jurkat T lymphoma cells: Comparison of seven mitochondrion-specific fluorochromes, Immunol Lett 61:157-163 (1998). 167. Marchetti, P, D. D, et al., Redox regulation of apoptosis: impact of thiol oxidation status on mitochondrial function, Eur J Immunol 27:289-296 (1997). 168. S. Susin, H. Lorenzo, N. Zamzami, et al., Mitochondrial release of caspase-2 and -9 during the apoptotic process, J Exp Med 189:381-393 (1999). 169. Z. Qin, Y. Wang, K. Kikly, et al., Pro-caspase-8 is predominantly localized in mitochondria and released into cytoplasm upon apoptotic stimulation, J Biol Chem 276:8079-8086 (2001).

108

Chapter 5

170. S. Srinivasula, P. Datta, X.-J. Fan, T. Fernandes-Alnemri, Z. Huang, and E. Alnemri, Molecular determinants of the caspase-promoting activity of Smac/DIABLO and its role in the death receptor pathway, J Biol Chem 275:36152-36157 (2000). 171. X. Liu, C. Kim, J. Yang, R. Jemmerson, and X. Wang, Induction of apoptotic program in cell-free extract: requirement for dATP and cytochrome c, Cell 86:147-157 (1996). 172. G. van Loo, X. Saelens, F. Matthijssens, et al., Caspases are not localized in mitochondria during life or death, Cell Death Differ 9:1207-11 (2002). 173. J. D. Robertson, M. Enoksson, M. Suomela, B. Zhivotovsky, and S. Orrenius, Caspase-2 acts upstream of mitochondria to promote cytochrome c release during etoposide-induced apoptosis, J Biol Chem 277:29803-9 (2002). 174. J. A. Klein, C. M. Longo-Guess, M. P. Rossmann, et al., The harlequin mouse mutation downregulates apoptosis-inducing factor, Nature 419:367-74 (2002). 175. C. Cande, I. Cohen, E. Daugas, et al., Apoptosis-inducing factor (AIF): a novel caspaseindependent death effector released from mitochondria, Biochimie 84:215-22 (2002). 176. G. Pan, E. Humke, and V. Dixit, Activation of caspases triggered by cytochrome c in vitro, FEBS Letts 426:151-154 (1998). 177. S. Hu, and X. Yang, Cellular inhibitor of apoptosis 1 and 2 are ubiquitin ligases for the apoptosis inducer Smac/DIABLO, J Biol Chem 278:10055-60 (2003). 178. S. Shimizu, Y. Eguchi, W. Kamiike, et al., Bcl-2 blocks loss of mitochondrial membrane potential while ICE inhibitors act at a different step during inhibition of death induced by respiratory chain inhibitors, Oncogene 13:21-29 (1996). 179. I. Marzo, C. Brenner, N. Zamzami, et al., The permeability transition pore complex: a target for apoptosis regulation by caspases and bcl-2-related proteins, J Exper Med 187:1261-1271 (1998). 180. N. Zamzami, C. Brenner, I. Marzo, S. Susin, and G. Kroemer, Subcellular and submitochondrial mode of action of Bcl-2-like oncoproteins, Oncogene 16:2265-2282 (1998). 181. J. Garland, and A. Halestrap, Energy metabolism during apoptosis: Bcl-2 promotes survival in hematopoietic cells induced to apoptose by growth factor withdrawal by stabilizing a form of metabolic arrest., J Biol Chem 272:4680-4688 (1997). 182. D. Plas, S. Talapatra, A. Edinger, J. Rathmell, and C. Thompson, Akt and Bcl-xL promote growth factor-independent survival through distinct effects on mitochondrial physiology, J Biol Chem 276:12041-12048 (2001). 183. T. Kelley, M. Graham, A. Doseff, et al., Macrophage colony-stimulating factor promotes cell survival through Akt/protein kinase B, J Biol Chem 274:26393-26398 (1999). 184. I. Fabregat, B. Herrera, M. Fernández, et al., Epidermal growth factor impairs the cytochrome C-caspase-3 apoptotic pathway induced by transforming growth factor beta in rat fetal hepatocyctes via a phosphoinositide 3-kinase-dependent pathway., Hepatol 32:528-535 (2000). 185. R. Jones, M. Parsons, M. Bonnard, et al., Protein kinase B regulates T lymphocyte survival nuclear factor kappa B activation and Bcl-X(L) levels in vivo, J Exper Med 191:1721-1734 (2000). 186. R. G. Jones, A. R. Elford, M. J. Parsons, et al., CD28-dependent activation of protein kinase B/Akt blocks Fas-mediated apoptosis by preventing death-inducing signaling complex assembly, J Exp Med 196:335-48 (2002). 187. K. Gottlob, N. Majewski, S. Kennedy, E. Kandel, R. Robey, and N. Hay, Inhibition of early apoptotic events by Akt/PKB is dependent on the first committed step of glycolysis and mitochondrial hexokinase, Genes Devel 15:1406-1418 (2001).


109

188. L. Del Peso, M. González-García, C. Page, R. Herrera, and G. Nuñez, Interleukin-3induced phosphorylation of BAD through the protein kinase Akt., Science 278:687-689 (1997). 189. P. Coffer, J. Jin, and J. Woodgett, Protein kinase B (c-Akt): a multifunctional mediator of phosphatidylinositol 3-kinase activation., Biochem J 335:1-13 (1998). 190. C. Yang, A. Murti, S. Pfeffer, J. Kim, D. Donner, and L. Pfeffer, Interferon alpha /beta promotes cell survival by activating nuclear factor kappa B through phosphatidylinositol 3-kinase and Akt, J Biol Chem 276:13756-13761 (2001). 191. M. Vander Heiden, X. Li, E. Gottleib, R. Hill, C. Thompson, and M. Colombini, Bcl-xL promotes the open configuration of the voltage-dependent anion channel and metabolite passage through the outer mitochondrial membrane, J Biol Chem 276:19414-19419 (2001). 192. E. Gottlieb, H. Vander, MG, and C. Thompson, Bcl-x(L) prevents the initial decrease in mitochondrial membrane potential and subsequent reactive oxygen species production during tumor necrosis factor alpha-induced apoptosis, Molec Cell Biol 20:5680-5689 (2000). 193. C. Scatena, Z. Stewart, D. Mays, et al., Mitotic phosphorylation of Bcl-2 during normal cell cycle progression and Taxol-induced growth arrest, J Biol Chem 273:30777-30784 (1998). 194. E. St. Clair, S. Anderson, and Z. Oltvai, Bcl-2 counters apoptosis by Bax heterodimerization-dependent and -independent mechanisms in the T-cell lineage, J Biol Chem 272:29347-29355 (1997). 195. T. Rossé, R. Olivier, L. Monney, et al., Bcl-2 prolongs cell survival after Bax-induced release of cytochrome c, Nature 391:496-499 (1998). 196. F. Li, A. Srinivasan, Y. Wang, R. Armstrong, K. Tomaselli, and L. Fritz, Cell-specific induction of apoptosis by microinjection of cytochrome C: Bcl-xL has activity independent of cytochrome C release, J Biol Chem 272:30299-30305 (1997). 197. B. Zhivotovsky, S. Orrenius, O. Brustugun, and S. Døskel, Injected cytochorme c induces apoptosis, Nature 391:449-450 (1998). 198. S. C. Ruffolo, and G. C. Shore, BCL-2 selectively interacts with the BID-induced open conformer of BAK, inhibiting BAK auto-oligomerization, J Biol Chem 278:25039-45 (2003). 199. S. Korsmeyer, X.-M. Yin, Z. Oltvai, D. Veis-Novack, and G. Linette, Reactive oxygen species and the regulation of cell death by the Bcl-2 gene family, Biochim Biophys Acta 1271:63-66(1995). 200. P. Bernardi, V. Petronilli, L. Di, F, and M. Forte, A mitochondrial perspective on cell death., Trends Biochem Sci 26:112-117 (2001). 201. A. Andreyev, B. Fahy, and G. Fiskum, Cytochrome c release from brain mitochondria is independent of the mitochondrial permeability transition., FEBS Letters 439:373-376 (1998). 202. E. Bossy-Wetzel, D. Newmeyer, and D. Green, Mitochondrial cytochrome c release in apoptosis occurs upstream of DEVD-specific caspase activation and independently of mitochondrial transmembrane depolarization., EMBO J 17:37-49 (1998). 203. M. Wei, T. Lindsten, V. Mootha, et al., tBID, a membrane-targeted death ligand, oligomerizes BAK to release cytochrome c, Genes Devel 14:2060-2071 (2000). 204. V. Gogvadze, J. Robertson, B. Zhivotovsky, and S. Orrenius, Cytochrome c release occurs via Ca2+-dependent and Ca2+-independent mechanisms that are regulated by Bax, J Biol Chem 276:19066-19071 (2001).

110

Chapter 5

205. Y. Tsujimoto, and S. Shimizu, The voltage-dependent anion channel: an essential player in apoptosis, Biochimie 84:187-93 (2002). 206. E. H. Cheng, T. V. Sheiko, J. K. Fisher, W. J. Craigen, and S. J. Korsmeyer, VDAC2 inhibits BAK activation and mitochondrial apoptosis. Science 301:513-7 (2003). 207. Y. Shi, J. Chen, C. Weng, et al., Identification of the protein-protein contact site and interaction mode of human VDAC1 with Bcl-2 family proteins, Biochem Biophys Res Commun 305:989-96 (2003). 208. E. Doran, and A. Halestrap, Cytochrome c release from isolated rat liver mitochondria can occur independently of outer-membrane rupture: possible role of contact sites., Biochem J 348:343-350 (2000). 209. J. Pastorino, N. Shulga, and J. Hoek, Mitochondrial binding of hexokinase II inhibits Bax-induced cytochrome c release and apoptosis, J Biol Chem 277:7610-7618 (2002). 210. M. Y. Vyssokikh, and D. Brdiczka, The function of complexes between the outer mitochondrial membrane pore (VDAC) and the adenine nucleotide translocase in regulation of energy metabolism and apoptosis, Acta Biochim Pol 50:389-404 (2003). 211. M. Ott, J. Robertson, V. Gogvadze, B. Zhivotovsky, and S. Orrenius, Cytochrome c release from mitochondria proceeds by a two-step process, Proc Natl Acad Sci USA 99:1259-1263 (2002). 212. L. Scorrano, M. Ashiya, K. Buttle, et al., A distinct pathway remodels mitochondrial cristae and mobilizes cytochrome c during apoptosis, Dev Cell 2:55-67 (2002). 213. M. Zhang, E. Mileykovskaya, and W. Dowhan, Gluing the respiratory chain together. Cardiolipin is required for supercomplex formation in the inner mitochondrial membrane, J Biol Chem 277:43553-6 (2002). 214. W.-X. Zong, T. Lindsten, A. Ross, G. MacGregor, and C. Thompson, BH3-only proteins that bind pro-survival Bcl-1 family members fail to induce apoptosis in the absence of Bax and Bak, Genes Devel 15:1481-1486 (2001). 215. L. Scorrano, S. A. Oakes, J. T. Opferman, et al., BAX and BAK regulation of endoplasmic reticulum Ca2+: a control point for apoptosis, Science 300:135-9 (2003). 216. T. Kuwana, M. R. Mackey, G. Perkins, et al., Bid, Bax, and lipids cooperate to form supramolecular openings in the outer mitochondrial membrane, Cell 111:331-42 (2002). 217. J.-M. Bruey, C. Ducasse, P. Bonniaud, et al., Hsp27 negatively regulates cell death by interacting with cytochrome c., Nature Cell Biol 2:645-652 (2000). 218. C. Paul, F. Manero, S. Gonin, C. Kretz-Remy, S. Virot, and A. Arrigo, Hsp27 as a negative regulator of cytochrome C release, Mol Cell Biol 22:816-834 (2002). 219. N. J. Waterhouse, J. C. Goldstein, O. von Ahsen, M. Schuler, D. D. Newmeyer, and D. R. Green, Cytochrome c maintains mitochondrial transmembrane potential and ATP generation after outer mitochondrial membrane permeabilization during the apoptotic process, J Cell Biol 153:319-28 (2001). 220. J. Cai, and D. Jones, Superoxide in apoptosis: Mitochondrial generation triggered by cytochrome c loss., J Biol Chem 273:11401-11404 (1998). 221. M. Schuler, and D. Green, Mechanisms of p53-dependent apoptosis, Biochem Soc Trans 29:684-688 (2001). 222. K. H. Vousden, Activation of the p53 tumor suppressor protein, Biochim Biophys Acta 1602:47-59 (2002). 223. U. M. Moll, and A. Zaika, Nuclear and mitochondrial apoptotic pathways of p53, FEBS Lett 493:65-9 (2001). 224. M. Oren, Decision making by p53: life, death and cancer, Cell Death Differ 10:431-42 (2003).


111

225. V. Stambolic, D. MacPherson, D. Sas, et al., Regulation of PTEN transcription by p53, Mol Cell 8:317-25 (2001). 226. N. Yahagi, H. Shimano, T. Matsuzaka, et al., p53 Activation in adipocytes of obese mice, J Biol Chem 278:25395-400 (2003). 227. P. Bouillet, and A. Strasser, BH3-only proteins - evolutionarily conserved proapoptotic Bcl-2 family members essential for initiating programmed cell death, J Cell Sci 115:156774 (2002). 228. K. Nakano, and K. Vousden, PUMA, a novel proapoptotic gene, is induced by p53, Molec Cell 7:683-694 (2001). 229. J. Yu, L. Zhang, P. Hwang, K. Kinzler, and B. Vogelstein, PUMA induces the rapid apoptosis of colorectal cancer cells, Molec Cell 7:673-682 (2001). 230. P. Li, R. Dietz, and R. von Harsdorf, p53 regulates mitochondrial membrane potential through reactive oxygen species and induces cytochrome c-independent apoptosis blocked by Bcl-2, Embo J 18:6027-6036 (1999). 231. N. Marchenko, A. Zaika, U. Moll, and D. s.-i. l. o. p. p. t. m. A. p. r. i. a. signaling, Death signal-induced localization of p53 protein to mitochondria. A potential role in apoptotic signaling, J Biol Chem 275:16202-16212 (2000). 232. M. Schuler, E. Bossy-Wetzel, J. Goldstein, P. Fitzgerald, and D. Green, p53 induces apoptosis by caspase activation through mitochondrial cytochrome c release, J Biol Chem 275:7337-7342 (2000). 233. C. Sansome, A. Zaika, N. Marchenko, and U. Moll, death, stimulus, induces, translocation, of, p53, protein, to, mitochondria, Detection by immunofluorescence on whole cells, Febs Letters 488:110-115 (2001). 234. M. Mihara, S. Erster, A. Zaika, et al., p53 Has a Direct Apoptogenic Role at the Mitochondria, Mol Cell 11:577-90 (2003). 235. G. Asher, J. Lotem, B. Cohen, L. Sachs, and Y. Shaul, Regulation of p53 stability and p53-dependent apoptosis by NADH quinone oxidoreductase 1, Proc Natl Acad Sci U S A 98:1188-93(2001). 236. P. Watcharasit, G. N. Bijur, J. W. Zmijewski, et al., Direct, activating interaction between glycogen synthase kinase-3beta and p53 after DNA damage, Proc Natl Acad Sci U S A 99:7951-5 (2002). 237. H. Li, Y. Cao, M. Berndt, J. Funder, and J. Liu, Molecular interactions between telomerase and the tumor suppressor protein p53 in vitro, Oncogene 18:6785-6794 (1999). 238. J. Liu, Telomerase: not just black and white but shades of gray, Mol Cell Biol Res Commun 3:129-135 (2000). 239. W. Fu, J. Begley, M. Killen, and M. Mattson, Anti-apoptotic role of telomerase in pheochromocytoma cells., J Biol Chem 274:7264-71 (1999). 240. S. Kang, T. Kwon, D. Kwon, and S. Do, Akt protein kinase enhances human telomerase activity through phosphorylation of telomerase reverse transcriptase subunit, J Biol Chem 274:13085-13090 (1999). 241. S. Artandi, S. Chang, S. Lee, et al., Telomere dysfunction promotes non-reciprocal translocations and epithelial cancers in mice., Nature 406:641-645 (2000). 242. D. Hanahan, Benefits of bad telomeres, Nature 406:573-574 (2000). 243. S. Shimizu, Y. Eguchi, W. Kamiike, et al., Bcl-2 prevents apoptotic mitochondrial dysfunction by regulating proton flux, Proc Natl Acad Sci USA 95:1455-1459 (1998). 244. A. Kowaltowski, S. Smaili, J. Russell, and G. Fiskum, Elevation of resting mitochondrial membrane potential of neural cells by cyclosporin A, BAPTA-AM, and Bcl-2, Am J Physiol Cell Physiol 279:C852-C859 (2000).

112

Chapter 5

245. L. Smets, J. Van den Berg, D. Acton, B. Top, H. Van Rooij, and M. Verwijs-Janssen, BCL-2 expression and mitochondrial activity in leukemic cells with different sensitivity to glucocorticoid-induced apoptosis, Blood 84:1613-1619 (1994). 246. T. Hennet, G. Bertoni, C. Richter, and E. Peterhans, Expression of BCL-2 protein enhances the survival of mouse fibrosarcoid cells in tumor necrosis factor-mediated cytotoxicity, Cancer Res 53:1456-1460 (1993A). 247. T. Graeber, C. Osmanian, T. Jacks, et al., Hypoxia-mediated selection of cells with diminished apoptotic potential in solid tumours, Nature 379:88-91 (1996). 248. D. McClintock, M. Santore, V. Lee, et al., Bcl-2 family members and functional electron transport chain regulate oxygen deprivation-induced cell death, Mol Cell Biol 22:94-104 (2002). 249. E. Wolvetang, K. Johnson, K. Krauer, S. Ralph, and A. Linnane, Mitochondrial respiratory chain inhibitors induce apoptosis, FEBS Letters 339:40-44 (1994). 250. V. Goossens, J. Grooten, and W. Fiers, The oxidative metabolism of glutamine: A modulator of reactive oxygen intermediate-mediated cytotoxicity of tumor necrosis factor in L929 fibrosarcoma cells, J Biol Chem 271:192-196 (1996). 251. J. Carretero, E. Obrador, J. Pellicer, A. Pascual, and J. Estrela, Mitochondial glutathione depletion by glutamine in growing tumor cells., Free Radical Biol Med 29:913-923 (2000). 252. M. Paumen, Y. Ishida, H. Han, et al., Direct interaction of the mitochondrial membrane protein carnitine palmitoyltransferase I with Bcl-2, Biochem Biophys Res Commun 231:523-525 (1997A). 253. A. Benedetti, A. Jezequel, and F. Orli, Preferential distribution of apoptotic bodies in acinar zone 3 of normal human and rat liver., J Hepatol 7:319-324 (1988). 254. L. Mullauer, B. Grasl-Kraupp, W. Bursch, and R. Schulte-Hermann, Transforming growth factor beta 1-induced cell death in preneoplastic foci of rat liver and sensitization by the antiestrogen tamoxifen, Hepatology 23:840-847 (1996). 255. K. Ohno, P. Ammann, R. Fasciati, and P. Maier, Transforming growth factor beta 1 preferentially induces apoptotic cell death in rat hepatocytes cultured under pericentralequivalent conditions, Toxicol Appl Pharmacol 132:227-236 (1995). 256. C. Scaffidi, I. Schmitz, J. Zha, S. Korsmeyer, P. Krammer, and M. Peter, Differential modulation of apoptosis sensitivity in CD95 type I and type II cells, J Biol Chem 274:22532-22538(1999). 257. D. Perry, and Y. Hannun, The role of ceramide in cell signaling, Biochim Biophys Acta 1436:233-243 (1998). 258. M. Krönke, Involvement of sphingomyelinases in TNF signaling pathways, Chem Phys Lipids 102:157-166 (1999). 259. Y. A. Hannun, and L. M. Obeid, The Ceramide-centric universe of lipid-mediated cell regulation: stress encounters of the lipid kind, J Biol Chem 277:25847-50 (2002). 260. R. Kolesnick, The therapeutic potential of modulating the ceramide/sphingomyelin pathway, J Clin Invest 110:3-8 (2002). 261. M. Rippo, F. Malisan, L. Ravagnan, et al., GD3 ganglioside directly targets mitochondria in a bcl-2-controlled fashion, FASEB J 14:2047-2054 (2000). 262. C. Garcia-Ruiz, A. Colell, A. Morales, M. Calvo, C. Enrich, and J. C. Fernandez-Checa, Trafficking of ganglioside GD3 to mitochondria by tumor necrosis factor-alpha, J Biol Chem 277:36443-8 (2002). 263. L. J. Siskind, R. N. Kolesnick, and M. Colombini, Ceramide channels increase the permeability of the mitochondrial outer membrane to small proteins, J Biol Chem 277:26796-803 (2002).


113

264. A. Arora, B. Jones, T. Patel, S. Bronk, and G. Gores, Ceramide induces hepatocyte cell death through disruption of mitochondrial function in the rat., Hepatology 25:958-963 (1997). 265. C. Garcia-Ruiz, A. Colell, M. Mari, A. Morales, and J. Fernez-Checa, Direct effect of ceramide on the mitochondrial electron transport chain leads to generation of reactive oxygen species: Role of mitochondrial glutathione., J Biol Chem 272:11369-11377 (1997). 266. T. Gudz, K. Tserng, and C. Hoppel, Direct inhibition of mitochondrial respiratory chain complex III by cell-permeable ceramide, J Biol Chem 272:24154-24158 (1997). 267. A. Quillet-Mary, J. Jaffrézou, V. Mansat, C. Bordier, J. Naval, and G. Laurent, Implication of mitochondrial hydrogen peroxide generation in ceramide-induced apoptosis, J Biol Chem 272:21388-21395 (1997). 268. K. Schubert, M. Scheid, and V. Duronio, Ceramide inhibits protein kinase B/Akt by promoting dephosphorylation of serine 473, J Biol Chem 275:13330-13335 (2000). 269. K. Schulze-Osthoff, A. Bakker, B. Vanhaesebroeck, R. Beyaert, W. Jacob, and W. Fiers, Cytotoxic activity of tumor necrosis factor is mediated by early damage of mitochondrial functions: Evidence for the involvement of mitochondrial radical generation, J Biol Chem 267:5317-5323 (1992). 270. H. Masaki, and H. Sakurai, Increased generation of hydrogen peroxide possibly from mitochondrial respiratory chain after UVB irradiation of murine fibroblasts, J Derm Sci 14:207-216 (1997). 271. S. Shimmura, and K. Tsubota, Ultraviolet B-induced mitochondrial dysfunction is associated with decreased cell detachment of corneal epithelial cells in vitro, Invest Ophthalmol Visual Sci 38:620-626 (1997). 272. Y. Aragane, D. Kulms, D. Metze, et al., Ultraviolet light induces apoptosis via direct activation of CD95 (Fas/APO-1) independently of its ligand CD95L., J Cell Biol 140:171182 (1998). 273. C. Belka, J. Rudner, S. Wesselborg, et al., Differential role of caspase-8 and BID activation during radiation- and CD95-induced apoptosis., Oncogene 19:1181-1190 (2000). 274. H. Miyoshi, C. Rust, P. Roberts, L. Burgart, and G. Gores, Hepatocyte apoptosis after bile duct ligation in the mouse involves Fas, Gastroenterology 117:669-677 (1999). 275. B. Yerushalmi, R. Dahl, M. Devereaux, E. Gumpricht, and R. Sokol, Bile acid-induced rat hepatocyte apoptosis is inhibited by antioxidants and blockers of the mitochondrial permeability transition, Hepatol 33:616-626 (2001). 276. Z. Han, P. Pantazis, J. Wyche, N. Kouttab, V. Kidd, and E. Hendrickson, A Fasassociated death domain protein-dependent mechanism mediates the apoptotic action of non-steroidal anti-inflammatory drugs in the human leukemic Jurkat cell line, J Biol Chem 276:38748-38754(2001). 277. O. Micheau, E. Solary, A. Hammann, and M. Dimanche-Boitrel, Fas ligand-independent, FADD-mediated activation of the Fas death pathway by anticancer drugs, J Biol Chem 274:7987-7992(1999). 278. Y. Fan, J. Zhang, R. Barhoumi, et al., Antagonism of CD95 signaling blocks butyrate induction of apoptosis in young adult mouse colonic cells., Am J Physiol 277:C310-C319 (1999). 279. T. Quan, T. He, S. Kang, J. J. Voorhees, and G. J. Fisher, Ultraviolet irradiation alters transforming growth factor beta/smad pathway in human skin in vivo, J Invest Dermatol 119:499-506 (2002).

114

Chapter 5

280. T. Maeda, M. T. Chong, R. A. Espino, et al., Role of p21 (Waf-1) in regulating the G1 and G2/M checkpoints in ultraviolet-irradiated keratinocytes, J Invest Dermatol 119:51321 (2002). 281. H. Higuchi, S. Bronk, Y. Takikawa, et al., The bile acid glycochenodeoxycholate induces trail-receptor 2/DR5 expression and apoptosis, J Biol Chem 276:38610-38618 (2001). 282. S. de la Monte, T. Neely, J. Cannon, and J. Wands, Ethanol impairs insulin-stimulated mitochondrial function in cerebellar granule neurons., Cell Mol Life Sci 58:1950-1960 (2001). 283. A. Colell, O. Coll, C. Garcia-Ruiz, et al., Tauroursodeoxycholic acid protects hepatocytes from ethanol-fed rats against tumor necrosis factor-induced cell death by replenishing mitochondrial glutathione., Hepatology 34:964-971 (2001). 284. C. Rodrigues, and C. Steer, The therapeutic effects of ursodeoxycholic acid as an antiapoptotic agent, Expert Opin Investig Drugs 10:1243-1253 (2001). 285. D. Decraene, P. Agostinis, R. Bouillon, H. Degreef, and M. Garmyn, Insulin-like growth factor-1-mediated AKT activation postpones the onset of ultraviolet B-induced apoptosis, providing more time for cyclobutane thymine dimer removal in primary human keratinocytes, J Biol Chem 277:32587-95 (2002). 286. A. Kurz, C. Block, D. Graf, S. Dahl, F. Schliess, and D. Haussinger, Phosphoinositide 3kinase-dependent Ras activation by tauroursodesoxycholate in rat liver, Biochem J 350:207-13 (2000). 287. T. Suhara, K. Fukuo, T. Sugimoto, et al., Hydrogen peroxide induces up-regulation of Fas in human endothelial cells, J Immunol 160:4042-4047 (1998). 288. L. Feldenberg, S. Thevananther, M. Del Rio, M. De Leon, and P. Devarajan, Partial ATP depletion induces Fas- and caspase-mediated apoptosis in MDCK cells., Am J Physiol 276:F837-846 (1999). 289. D. Kwon, C. Choi, J. Lee, et al., Hydrogen peroxide triggers the expression of Fas/FasL in astrocytoma cell lines and augments apoptosis, J Neuroimmunol 113:1-9 (2001). 290. H. Chan, D. P. Bartos, and L. B. Owen-Schaub, Activation-dependent transcriptional regulation of the human Fas promoter requires NF-kappaB p50-p65 recruitment, Mol Cell Biol 19:2098-108 (1999). 291. Y. Zheng, F. Ouaaz, P. Bruzzo, V. Singh, S. Gerondakis, and A. A. Beg, NF-kappa B RelA (p65) is essential for TNF-alpha-induced fas expression but dispensable for both TCR-induced expression and activation-induced cell death, J Immunol 166:4949-57 (2001). 292. Y. Yamada, K. Kajiwara, M. Yano, E. Kishida, Y. Masuzawa, and S. Kojo, Increase of ceramides and its inhibition by catalase during chemically induced apoptosis of HL-60 cells determined by electrospray ionization tandem mass spectrometry, Biochim Biophys Acta 1532:115-120 (2001). 293. A. E. Feldstein, A. Canbay, P. Angulo, et al., Hepatocyte apoptosis and Fas expression are prominent features of human nonalcoholic steatohepatitis, Gastroenterology 125:43743 (2003). 294. M. H. Schoemaker, W. M. Gommans, L. Conde de la Rosa, et al., Resistance of rat hepatocytes against bile acid-induced apoptosis in cholestatic liver injury is due to nuclear factor-kappa B activation, J Hepatol 39:153-61 (2003). 295. F. Petit, D. Arnoult, J. Lelievre, et al., Productive HIV-1 infection of primary CD4+ T cells induces mitochondrial membrane permeabilization leading to a caspase-independent cell death, J Biol Chem 277:1477-1487 (2002).


115

296. M. MacFarlane, W. Merrison, D. Dinsdale, and G. Cohen, Active caspases and cleaved cytokeratins are sequestered into cytoplasmic inclusions in TRAIL-induced apoptosis, J Cell Biol 148:1239-1254 (2000). 297. G. Velasco, D. Gómez, Pulgar, T, D. Carlin, and M. Guzmán, Evidence that the AMPactivated protein kinase stimulates rat liver carnitine palmitoyltransferase I by phosphorylating cytoskeletal components, FEBS Lett 439:317-320 (1998A). 298. G. Velasco, M. Geelen, T. Gomez del Pulgar, and M. Guzmán, Malonyl-CoAindependent acute control of hepatic carnitine palmitoyltransferase I activity: Role of Ca2+/calmodulin-dependent protein kinase II and cytoskeletal components, J Biol Chem 273:21497-21504 (1998B). 299. N. Ku, X. Zhou, D. Toivola, and M. Omary, The cytoskeleton of digestive epithelia in health and disease, Am J Physiol 277:G1108-G1137 (1999). 300. K. Zatloukal, C. Stumptner, M. Lehner, et al., Cytokeratin 8 protects from hepatotoxicity, and its ratio to cytokeratin 18 determines the ability of hepatocytes to form Mallory bodies, Am J Pathol 156:1263-1274 (2000). 301. C. Caulin, C. Ware, T. Magin, and R. Oshima, Keratin-dependent, epithelial resistance to tumor necrosis factor-induced apoptosis., J Cell Biol 149:17-22. (2000). 302. S. Gilbert, A. Loranger, N. Daigle, and N. Marceau, Simple epithelium keratins 8 and 18 provide resistance to Fas-mediated apoptosis: The protection occurs through a receptortargeting modulation, J Cell Biol 154:763-73 (2001). 303. U. Steinhusen, V. Badock, A. Bauer, et al., Apoptosis-induced cleavage of beta-catenin by caspase-3 results in proteolytic fragments with reduced transactivation potential, J Biol Chem 275:16345-16353 (2000). 304. R. Bachelder, M. Ribick, A. Marchetti, et al., p53 inhibits alpha 6 beta 4 integrin survival signaling by promoting the caspase 3-dependent cleavage of AKT/PKB., J Cell Biol 147:1063-1072 (1999). 305. G. Tang, J. Yang, Y. Minemoto, and A. Lin, Blocking caspase-3-mediated proteolysis of IKKbeta suppresses TNF-alpha-induced apoptosis, Mol Cell 8:1005-1016 (2001). 306. J. Park, K. Kim, S. Kim, and S. Lee, Caspase 3 specifically cleaves p21WAF1/CIP1 in the earlier stage of apoptosis in SK-HEP-1 human hepatoma cells, Eur J Biochem 257:242-248 (1998). 307. C. Sherr, and J. Roberts, CDK inhibitors: positive and negative regulators of G1-phase progression, Genes Dev 13:1501-12 (1999). 308. O. Coqueret, New roles for p21 and p27 cell-cycle inhibitors: a function for each cell compartment?, Trends Cell Biol 13:65-70 (2003). 309. B. Gleiss, V. Gogvadze, S. Orrenius, and B. Fadeel, Fas-triggered phosphatidylserine exposure is modulated by intracellular ATP, FEBS Lett 519:153-8 (2002). 310. G. Kass, J. Eriksson, M. Weis, S. Orrenius, and S. Chow, Chromatin condensation during apoptosis requires ATP, Biochem J 318:749-752 (1996). 311. Y. Eguchi, S. Shimizu, and Y. Tsujimoto, Intracellular ATP levels determine cell death fate by apoptosis or necrosis., Cancer Res 57:1835-1840 (1997). 312. M. Leist, B. Single, A. Castoldi, S. Kuhnle, and P. Nicotera, Intracellular adenosine triphosphate (ATP) concentration: a switch in the decision between apoptosis and necrosis, J Exp Med 185:1481-1486 (1997). 313. T. Hirsch, P. Marchetti, S. Susin, et al., The apoptosis-necrosis paradox Apoptogenic proteases activated after mitochrondrial permeability transition determine the mode of cell death, Oncogene 15:1573-1581 (1997). 314. A. Slater, C. Nobel, and S. Orrenius, The role of intracellular oxidants in apoptosis, Biochim Biophys Acta 1271:59-62 (1995).

116

Chapter 5

315. C. Richter, M. Schweizer, A. Cossarizza, and C. Franceschi, Control of apoptosis by the cellular ATP level, FEBS Lett 378:107-110 (1996). 316. W. Meikrantz, and R. Schlegel, Apoptosis and the cell cycle, J Cell Biochem 58:160-174 (1995). 317. G. Fan, X. Ma, B. Kren, and C. Steer, The retinoblastoma gene product inhibits TGFbeta1 induced apoptosis in primary rat hepatocytes and human HuH-7 hepatoma cells., Oncogene 12:1909-1919 (1996). 318. G. Gil-Gómez, A. Berns, and H. Brady, A link between cell cycle and cell death: Bax and Bcl-2 modulate Cdk2 activation during thymocyte apoptosis, EMBO J 17:7209-7218 (1998). 319. E. Thompson, The many roles of c-Myc in apoptosis, Annu Rev Physiol 60:575-600 (1998). 320. K. Hiromura, J. Pippin, M. Fero, J. Roberts, and S. Shankland, Modulation of apoptosis by the cyclin-dependent kinase inhibitor p27(Kip1), J Clin Invest 103:597-604 (1999). 321. E. Lind, J. Wayne, Q.-Z. Wang, T. Staeva, A. Stolzer, and H. Petrie, Bcl-2-induced changes in E2F regulatory complexes reveal the potential for integrated cell cycle and cell death functions, J Immunol 162:5374-5379 (1999). 322. K. Hoeflich, J. Luo, E. Rubie, M. Tsao, O. Jin, and J. Woodgett, Requirement for glycogen synthase kinase-3beta in cell survival and NF-kappaB activation, Nature 406:8690 (2000). 323. B. Bi, N. Littlewood, and I. Crispe, Cleavage of E2F-1-regulating proteins and activation of E2F-1 during CD95-induced death of thymocytes., Immunology 104:37-42 (2001). 324. I. Shin, A. V. Bakin, U. Rodeck, A. Brunet, and C. L. Arteaga, Transforming growth factor beta enhances epithelial cell survival via Akt-dependent regulation of FKHRL1, Mol Biol Cell 12:3328-39 (2001). 325. D. Ginsberg, E2F1 pathways to apoptosis, FEBS Lett 529:122-5 (2002). 326. J. Klefstrom, E. W. Verschuren, and G. Evan, c-Myc augments the apoptotic activity of cytosolic death receptor signaling proteins by engaging the mitochondrial apoptotic pathway, J Biol Chem 277:43224-32 (2002). 327. R. F. Schwabe, and D. A. Brenner, Role of glycogen synthase kinase-3 in TNF-alphainduced NF-kappaB activation and apoptosis in hepatocytes, Am J Physiol Gastrointest Liver Physiol 283:G204-11 (2002). 328. D. Liu, and L. Greene, Neuronal apoptosis at the G1/S checkpoint, Cell Tissue Res 305:217-228 (2001). 329. E. Morris, E. Keramaris, H. Rideout, et al., Cyclin-dependent kinases and P53 pathways are activated independently and mediate Bax activation in neurons after DNA damage, J Neurosci 21:5017-5026 (2001). 330. D. R. Green, and G. I. Evan, A matter of life and death, Cancer Cell 1:19-30 (2002). 331. J. Scaife, and H. Brohee, An investigation of factors influencing mitotic G2 delay in synchronous cultures of human kidney cells and X-irradiation, Can J Biochem 47:237-249 (1969). 332. S. Skog, B. Tribukait, and G. Sundius, Energy metabolism and ATP turnover time during the cell cycle of Ehrlich ascites tumour cells, ExpCell Res 141:23-29 (1982). 333. S. Skog, B. Tribukait, and G. Sundius, Energy metabolism and ATP turnover time during the cell cycle in roentgen irradiated Ehrlich ascites tumour cells, Acta Radiol Oncol 22:369-379 (1983). 334. M. Jin, Y. Shimahara, T. Yamaguchi, et al., The effect of a bolus injection of TNF-alpha and IL-1 beta on hepatic energy metabolism in rats, J Surg Res 58:509-515 (1995).


117

335. D. Hardie, D. Carling, and M. Carlson, The AMP-activated/SNF1 protein kinase subfamily: metabolic sensors of the eukaryotic cell?, Annu Rev Biochem 67:821-855 (1998). 336. G. A. Rutter, G. da Silva Xavier, and I. Leclercq, Roles of 5’-AMP-activated protein kinase (AMPK) in mammalian glucose homeostasis, Biochem J 375:1-16 (2003). 337. Z. You, L. V. Madrid, D. Saims, J. Sedivy, and C. Y. Wang, c-Myc sensitizes cells to tumor necrosis factor-mediated apoptosis by inhibiting nuclear factor kappa B transactivation, J Biol Chem 277:36671-7 (2002). 338. J. Seoane, H. V. Le, and J. Massague, Myc suppression of the p21(Cip1) Cdk inhibitor influences the outcome of the p53 response to DNA damage, Nature 419:729-34 (2002). 339. D. Ma, P. Zhou, and J. W. Harbour, Distinct mechanisms for regulating the tumor suppressor and antiapoptotic functions of Rb, J Biol Chem 278:19358-66 (2003). 340. Z. Nahle, J. Polakoff, R. V. Davuluri, et al., Direct coupling of the cell cycle and cell death machinery by E2F, Nat Cell Biol 4:859-64 (2002). 341. C. Stefanelli, I. Stanic, F. Bonavita, et al., Inhibition of glucocorticoid-induced apoptosis with 5-aminoimidazole-4-carboxamide ribonucleoside, a cell-permeable activator of AMP-activated protein kinase, Biochem Biophys Res Commun 243:821-6 (1998). 342. C. Blazquez, M. J. Geelen, G. Velasco, and M. Guzman, The AMP-activated protein kinase prevents ceramide synthesis de novo and apoptosis in astrocytes, FEBS Lett 489:149-53 (2001). 343. C. Beauloye, A. S. Marsin, L. Bertrand, et al., Insulin antagonizes AMP-activated protein kinase activation by ischemia or anoxia in rat hearts, without affecting total adenine nucleotides, FEBS Lett 505:348-52 (2001). 344. Y. Minokoshi, Y. Kim, O. Peroni, et al., Leptin stimulates fatty-acid oxidation by activating AMP-activated protein kinase, Nature 415:339-343 (2002). 345. G. R. Steinberg, J. W. Rush, and D. J. Dyck, AMPK expression and phosphorylation are increased in rodent muscle after chronic leptin treatment, Am J Physiol Endocrinol Metab 284:E648-54 (2003). 346. L. G. Fryer, A. Parbu-Patel, and D. Carling, The Anti-diabetic drugs rosiglitazone and metformin stimulate AMP-activated protein kinase through distinct signaling pathways, J Biol Chem 277:25226-32 (2002). 347. X. Wu, H. Motoshima, K. Mahadev, T. J. Stalker, R. Scalia, and B. J. Goldstein, Involvement of AMP-activated protein kinase in glucose uptake stimulated by the globular domain of adiponectin in primary rat adipocytes, Diabetes 52:1355-63 (2003). 348. M. Garcia-Gil, R. Pesi, S. Perna, et al., 5’-aminoimidazole-4-carboxamide riboside induces apoptosis in human neuroblastoma cells, Neuroscience 117:811-20 (2003). 349. J. M. Lopez, A. F. Santidrian, C. Campas, and J. Gil, 5-Aminoimidazole-4-carboxamide riboside induces apoptosis in Jurkat cells, but the AMP-activated protein kinase is not involved, Biochem J 370:1027-32 (2003). 350. G. F. Merrill, E. J. Kurth, D. G. Hardie, and W. W. Winder, AICA riboside increases AMP-activated protein kinase, fatty acid oxidation, and glucose uptake in rat muscle, Am J Physiol. Endocrinol. Metab. 273:E1107-12 (1997). 351. L: Hue, C. Beauloye, A. S. Marsin, L. Bertrand, S. Horman, and M. H. Rider, Insulin and ischemia stimulate glycolysis by acting on the same targets through different and opposing signaling pathways, J Mol Cell Cardiol 34:1091-7 (2002). 352. N. Sambandam, and G. D. Lopaschuk, AMP-activated protein kinase (AMPK) control of fatty acid and glucose metabolism in the ischemic heart, Prog Lipid Res 42:238-56 (2003).

118

Chapter 5

353. L. H. Young, Y. Renfu, R. Russell, et al., Low-flow ischemia leads to translocation of canine heart GLUT-4 and GLUT-1 glucose transporters to the sarcolemma in vivo, Circulation 95:415-22 (1997). 354. R. R. Russell, 3rd, R. Bergeron, G. I. Shulman, and L. H. Young, Translocation of myocardial GLUT-4 and increased glucose uptake through activation of AMPK by AICAR, Am J Physiol 277:H643-9 (1999). 355. E. Tomas, T.-S. Tsao, A. Saha, et al., Enhanced muscle fat oxidation and glucose transport by ACRP30 globular domain: acetyl-CoA carboxylase inhibition and AMPactivated protein kinase activation., Proc Natl Acad Sci U S A 99:16309-16313 (2002). 356. K. Koike, K. Moriya, H. Yotsuyanagi, et al., Compensatory apoptosis in preneoplastic liver of a transgenic mouse model for viral hepatocarcinogenesis, Cancer Lett 134:181 -6 (1998). 357. P. Arbuthnot, A. Capovilla, and M. Kew, Putative role of hepatitis B virus X protein in hepatocarcinogenesis: effects on apoptosis, DNA repair, mitogen-activated protein kinase and JAK/STAT pathways., J Gastroenterol Hepatol. 15:357-68 (2000). 358. M. Honda, S. Kaneko, T. Shimazaki, et al., Hepatitis C virus core protein induces apoptosis and impairs cell-cycle regulation in stably transformed Chinese hamster ovary cells, Hepatology 31:1351-1359 (2000). 359. G. Kalkeri, N. Khalap, R. Garry, C. Fermin, and S. Dash, Hepatitis C virus protein expression induces apoptosis in HepG2 cells, Virology 282:26-37 (2001). 360. C. Madden, and B. Slagle, Stimulation of cellular proliferation by hepatitis B virus X protein, Dis Markers 17:153-157 (2001). 361. A. Basu, K. Meyer, R. B. Ray, and R. Ray, Hepatitis C virus core protein is necessary for the maintenance of immortalized human hepatocytes, Virology 298:53-62 (2002). 362. A. Erhardt, M. Hassan, T. Heintges, and D. Haussinger, Hepatitis C virus core protein induces cell proliferation and activates ERK, JNK, and p38 MAP kinases together with the MAP kinase phosphatase MKP-1 in a HepG2 Tet-Off cell line., Virology 292:272-284 (2002). 363. M. N. Lee, E. Y. Jung, H. J. Kwun, et al., Hepatitis C virus core protein represses the p21 promoter through inhibition of a TGF-beta pathway, J Gen Virol 83:2145-51 (2002). 364. M. Otsuka, N. Kato, H. Taniguchi, et al., Hepatitis C virus core protein inhibits apoptosis via enhanced Bcl-xL expression, Virology 296:84-93 (2002). 365. R. B. Ray, R. Steele, A. Basu, et al., Distinct functional role of Hepatitis C virus core protein on NF-kappaB regulation is linked to genomic variation, Virus Res 87:21-9 (2002). 366. W. L. Shin, M. L. Kuo, S. E. Chuang, A. L. Cheng, and S. L. Doong, Hepatitis B virus X protein activates a survival signaling by linking Src to phosphatidylinositol 3-kinase, J Biol Chem 278:31807-13 (2003). 367. M. Dubourdeau, T. Miyamura, Y. Matsuura, L. Alric, B. Pipy, and D. Rousseau, Infection of HepG2 cells with recombinant adenovirus encoding the HCV core protein induces p21(WAF1) down-regulation - effect of transforming growth factor beta, J Hepatol 37:486 (2002). 368. H. J. Han, E. Y. Jung, W. J. Lee, and K. L. Jang, Cooperative repression of cyclindependent kinase inhibitor p21 gene expression by hepatitis B virus X protein and hepatitis C virus core protein, FEBS Lett 518:169-72 (2002). 369. J. K. Leach, L. Qiao, Y. Fang, et al., Regulation of p21 and p27 expression by the hepatitis B virus X protein and the alternate initiation site X proteins, AUG2 and AUG3, J Gastroenterol Hepatol 18:376-385 (2003).


119

370. H. Nguyen, M. Mudryj, M. Guadalupe, and S. Dandekar, Hepatitis C virus core protein expression leads to biphasic regulation of the p21 cdk inhibitor and modulation of hepatocyte cell cycle, Virology 312:245-53 (2003). 371. Y. Li, D. Dowbenko, and L. A. Lasky, AKT/PKB phosphorylation of p21Cip/WAF1 enhances protein stability of p21Cip/WAF1 and promotes cell survival, J Biol Chem 277:11352-61 (2002). 372. J. Liang, J. Zubovitz, T. Petrocelli, et al., PKB/Akt phosphorylates p27, impairs nuclear import of p27 and opposes p27-mediated G1 arrest, Nat Med 8:1153-60 (2002). 373. I. Shin, F. M. Yakes, F. Rojo, et al., PKB/Akt mediates cell-cycle progression by phosphorylation of p27(Kip1) at threonine 157 and modulation of its cellular localization, Nat Med 8:1145-52 (2002). 374. M. Vail, R. Pierce, and N. Fausto, Bcl-2 delays and alters hepatic carcinogenesis induced by transforming growth factor alpha, Cancer Res 61:594-601 (2001). 375. C. Knudson, G. Johnson, Y. Lin, and S. Korsmeyer, Bax accelerates tumorigenesis in p53-deficient mice, Cancer Res 61:659-665 (2001).

Chapter 6 Metabolic Effects of Antiproliferative Agents

6.1.

INTRODUCTION

As discussed briefly in the preceding chapter, and in addition to the fatty acids discussed earlier (see Chapter 4: Fatty Acids and Growth Regulation), three structurally and mechanistically disparate growth inhibitors rapidly induce changes in mitochondrial function. Despite the fact that two are endogenous a protein; and butyrate, a short chain fatty acid), and one a group of xenobiotics (salicylate and nonsteroidal anti-inflammatory drugs, NSAIDs), all have been shown to increase mitochondrial fatty acid oxidation and ketogenesis, absolute and/or relative to ATP biosynthesis1 (Fig. 6.1; and Fig. 6.2 – facing page 123; also, Ockner R, unpublished data). They also may act in association with or independently of p53, the diverse and complex functions of which continue to unfold (see Chapter 5.3.2.: wg/Wnt-Like Signaling: Convergence of Mitochondria, p53, and Telomerase).

6.2.

BUTYRATE

Butyrate (butyric acid anion) is a 4-carbon short chain fatty acid (SCFA)2, that inhibits growth and induces p53-independent G1 cell cycle arrest and/or apoptosis in various cell systems3-7. In the relatively aggressive HM7 colorectal carcinoma line, butyrate promotes differentiation through its effects on and E-cadherin, shifting from nuclei to intercellular junctions, thereby serving to promote cell-cell adhesion and suggesting that it suppresses wg/Wnt-like signaling8. Butyrate also induces expression of cyclin-dependent kinase inhibitors and and dephosphorylation (activation) of 121

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Clofibrate, a rodent hepatocyte mitogen

Figure 6.1. Growth modulator regulation of rodent hepatocyte fatty acid oxidation. Pathway arrow thickness approximates relative activity. Top: Inhibition of CPT-I, and thus mitochondrial fatty acid oxidation and ketogenesis, by the peroxisome proliferator clofibrate. Although cellular mitochondrial fatty acid oxidation increases absolutely, the fatty acid burden is diminished relative to an increased mitochondrial oxidative capacity; extramitochondrial FA oxidation is relatively enhanced. Bottom: Augmentation of mitochondrial fatty acid oxidation and ketogenesis by butyrate, with relative suppression of extramitochondrial FA oxidation. Abbreviations — CPT-I: carnitine palmitoyltransferase-I; FA: fatty acid; PPARct: peroxisome proliferatoractivated

retinoblastoma protein9-12; in Caco-2 cells it selectively increases the 13 expression of , as noted above (see Chapter 5.2.2.: In butyrate-exposed colon cancer cell lines, the apoptotic pathway may be activated independently of histone deacetylase inhibition14. Butyrate induction of growth arrest or apoptosis is associated with 11 decrease or collapse, respectively, of . These effects can be understood in terms of butyrate’s intramitochondrial metabolism (Fig. 6.2 – facing page 123). Thus, similar to other SCFA, butyrate readily permeates the mitochondrial inner membrane as the unesterified protonated fatty acid, independent of CPT-I. Within the mitochondrial matrix, butyrate is converted to butyryl CoA and undergoes as a fuel source which generates acetyl CoA and NADH, it contributes electrons to the electron transport chain and leads to the inhibition of pyruvate dehydrogenase and glycolysis15,l6. However, butyrate also exerts an uncoupling-like effect on oxidative phosphorylation in which increased state 4 respiration reflects a futile cycle of repeated butyryl

Figure 6.2. Sites of growth inhibitor effects on mitochondrial function. Salicylic acid (salicylate) is a classical uncoupler of oxidative phosphorylation; i.e, a protonophore; thus, it delivers protons to the mitochondrial matrix while bypassing the ATP synthase complex, thereby driving respiration and substrate utilization without generating ATP. Butyric acid (butyrate) and ketone bodies (not shown) freely enter the mitochondrial matrix as protonated acids. They are relatively weak protonophores, and further differ from classical protonophore uncouplers in that they undergo oxidation in the mitochondrial matrix, thus increasing electron transport chain flux and propensity to ROS generation. Their uncoupling-like effect derives principally from intramitochondrial ATP consumption by futile cycles of acyl CoA synthesis and hydrolysis. Sulindac is a direct inhibitor of complex I of the electron transport chain, through which approximately 75% of the electrons generated by biological oxidation reactions flow. Abbreviations — CPT-I: carnitine palmitoyltransferase-I; ketone bodies: and acetoacetate; PDH: pyruvate dehydrogenase.

6. Metabolic Effects of Antiproliferative Agents

123

CoA synthesis and hydrolysis17,18 (Fig. 6.2 – facing this page). As a result of this futile cycling, increased intramitochondrial consumption of ATP and generation of AMP drives increases in oxygen and substrate utilization, and ATP synthesis. Presumably it is through these latter effects that butyrate augments mitochondrial long chain fatty acid oxidation and ketogenesis1,19, further increasing matrix reducing potential and the predisposition to oxidative stress. These characteristics of short chain fatty acid (e.g., butyrate) metabolism differ importantly from those of long chain fatty acids, in that de novo synthesis of long chain acyl CoA thioesters (and, therefore, of their acylcarnitine derivatives) in the mitochondrial matrix is negligible. As a result, underivatized protonated long chain fatty acids which enter the matrix via nonionic diffusion are not oxidized (unlike their acylcarnitines), but instead are exported as anions across the inner mitochondrial membrane. Thus, while long chain fatty acids are relatively weak uncouplers, they are similar in this particular respect to the classical protonophore uncouplers of oxidative phosphorylation, which also are exported unoxidized across the inner mitochondrial membrane. Accordingly, while the uncoupling-like effect of short chain fatty acids is accompanied by their and therefore increased matrix reducing potential and propensity for ROS generation, classical protonophore uncouplers exert an antioxidant effect, suppressing ROS generation20-25 (also, see this Chapter 6.4: Salicylate and Other Nonsteroidal Anti-Inflammatory Drugs). It is of additional significance that enzymes which mediate synthesis and hydrolysis of ketone body CoA thioesters, and their are also present in the mitochondrial matrix18,26. This suggests that an ATPconsuming, futile cycling, electron- and ROS-generating uncoupling-like effect, similar to that of butyrate, occurs during mitochondrial ketone body oxidation. These characteristics of mitochondrial matrix ketone body metabolism may account for the in vivo suppression of tumor growth by noted above27,28 (also, see Chapter 4: Fatty Acids and Growth Regulation), for the adverse of ketone bodies on neuronal function (see Chapter 12.3.: Ketone Bodies; and Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors), and for the antiseizure effects of ketogenic diets (see Chapter 14.4.: Ketogenic Diet and Control of Seizure Activity). The mechanistic basis of the reported inhibition of butyrate-induced apoptosis by Bcl-2 overexpression29, and of butyrate downregulation of Bcl-2, are unknown. However, recent evidence that butyrate-induced apoptosis in mouse colon cells is mediated by Fas30 further supports the concept that in certain circumstances Fas

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may be activated downstream of mitochondrial events, especially related to ROS generation (see Chapter 5.3.: Mitochondrial Function in Apoptosis). In light of the foregoing evidence, normal colonic epithelium represents a special case. Thus, in non-ruminants SCFA are produced only by colonic bacterial flora, and growth of normal differentiating colonocytes appears to be supported rather than inhibited by butyrate and other SCFA. It may therefore seem surprising that in germ-free animals, in which SCFA are absent, only modest changes in colonic epithelial cell proliferation rate and cell cycle duration are observed31. In other words, there clearly is no absolute colonocyte dependence on SCFA, which therefore may be required in conventional animals only to modulate direct or indirect (e.g., pro-inflammatory) effects of the colonic microflora on the epithelium. The alternative substrates utilized in the germ-free state, and also in conventional animals, include glutamine and, to a lesser extent, glucose32,33. Through the actions of glutaminase and glutamate dehydrogenase, glutamine-derived glutamate provides 2-oxoglutarate directly to the citric acid cycle (Fig. 6.2 – facing page 123), thus driving oxidative phosphorylation while circumventing control by pyruvate dehydrogenase, the activity of which is susceptible to inhibition by the aforementioned intramitochondrial effects of the normally high ambient butyrate concentrations15. Glutamine, therefore, may support the increases in and ATP generation required by cell cycle progression in the small subset of colonic crypt cells comprising the proliferative compartment. This relatively unrestrained fueling of the citric acid cycle and electron transport chain would be consistent with glutamine’s ability to prevent apoptosis resulting from decreased glucose uptake and utilization in IL3-deprived ICDP cells34, and its recognized role as an important respiratory substrate in proliferating normal and transformed cells35-38, yet also account for its predisposition of cells to apoptosis as noted above38 (see Chapter 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration and Oxidative Phosphorylation). In contrast, in the non-proliferating compartment in which colonocytes undergo differentiation, butyrate’s adverse effects on the efficiency of ATP generation would be of lesser significance. In fact, this abundant fuel (approximately 30-40 mM in the colonic lumen) would contribute to the maintenance of a normal differentiating (i.e., non-proliferative) phenotype.


6.3.

125

TRANSFORMING GROWTH

is a 25 kDa homodimer in its fully processed mature form, which acts through its cell surface receptors to increase the expression of the cell cycle inhibitors and thereby modulating cyclin-dependent kinase activity and inducing p53-independent late G1 cell cycle arrest39-41. The antiproliferative effect of may also reflect 43 induced expression of E- and P-cadherins42, or of . This observation is consistent with, although not directly linked to, downregulation of high mobility group-I (Y) protein44, a member of the Tcf/Lef group of transcription factors involved in (and thus wg/Wnt-Like) signaling45,46. The pro-apoptotic effect of may result from diminished expression of Bcl-2 or Bcl-xL47,48, caspase-mediated cleavage/activation of Bad49, and increased ROS generation50,51 (also, see below, this section). Substantial progress has been made in the elucidation of receptor-activated signaling pathways that may contribute to these effects52-55. For several, transcriptional regulation requires cooperative interaction between receptor-activated Smads and p5356. Contrasting growth-promoting (e.g., on IMR-90 human fibroblasts) and growth-inhibiting effects (e.g., on HuCCT1 human cholangiocarcinoma cells) may, among other possible factors, reflect: 1) differential regulation by of interferon regulatory factor-1-mediated expression57; 2) generation of AP-1 forms differing in monomer composition58; 3) suppression of activation59; and/or 4) changes in abundance and/or activity of signal mediators and regulators55,60,61. Among the last of these, interaction between seemingly opposing signaling by PI3K/Akt/PKB — e.g., induced by integrin-linked kinase62,63 — and is required for epithelial-tomesenchymal transition and migration in breast tumor cell lines53,64, fetal rat hepatocytes65, and renal tubular interstitial fibrosis63. also has significant effects on cell metabolism, including increased cellular generation of ROS48,50,51. In addition, activation of latent is enhanced by NO-induced nitrosylation of latencyassociated peptide, possibly in a cell- or tissue-specific manner66, while active post-transcriptionally suppresses iNOS expression67. Within the hepatocyte and cardiac myocyte (both of which exhibit high rates of mitochondrial fatty acid oxidation), as well as in other cell types, cytoplasmic is associated predominantly with mitochondria68-72. The association involves both mature and and does not permit 68,73 rapid exchange with soluble exogenous , but is otherwise structurally and functionally uncharacterized. Appearance of in

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this cytoplasmic location, external to the lumen of the endoplasmic reticulum, implies absence of a signal peptide; accordingly, it may reflect transcription from a recognized alternative start site74. Active is also demonstrable within B cells and plasma cells, in which its cellular localization and function have not been defined75. Intracellular locations and sites of action have been observed for other humoral or paracrine 76 mediators, including . Significantly, exogenous inhibits key glycolysis-promoting processes in the hepatocyte77, consistent with its augmentation of gluconeogenesis (PEPCK)78. Both of these effects imply an increase in mitochondrial fatty acid oxidation. Furthermore, they are consistent with evidence that mature increases mitochondrial fatty acid oxidation and ketogenesis, both systemically after intracisternal (intracranial) injection79, and in hepatocytes in primary culture 1 . In the latter, the ratio of to [acetoacetate] is increased1, implying an increase in electron transport chain electron density and, therefore, a predisposition to ROS generation. Thus, changes in fatty acid oxidation would predispose to intramitochondrial oxidative stress, thereby accounting at least in part for the above-noted 50 generation of and, in turn, early activation of AP-180 and 81 PI3K/Akt/PKB . Furthermore, the linkage of to fatty acid oxidation-induced mitochondrial ROS generation provides a mechanistic basis for ROS mediation of apoptosis in fetal hepatocytes51, attenuation of ROS and activation by prior heat shock82, and prevention of apoptosis by ursodeoxycholic acid83 (also, see Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). Linkage of to fatty acid metabolism is further supported by recent evidence that fatty acids modulate activity and plasma clearance84, and by regulation of gene expression43. Moreover, this linkage may contribute to the pathogenesis of kidney disease in experimental diabetes mellitus; in this disorder, protection afforded by abrogation of signaling85,86 would be consistent with participation of fatty acid oxidation in pathogenesis. A proposed role for hyperglycemia in this process87 is addressed below (see Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). These metabolic effects of — augmented mitochondrial fatty acid oxidation and ketogenesis, increases in intramitochondrial reducing potential and ROS generation, suppression of glycolysis, and increased gluconeogenesis — are antiproliferative (see Chapter 3: Nutrient and Energy Metabolism in Cell Proliferation). They may also contribute to activation of caspase 8 and apoptosis, apparently


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independent of Fas88. Consistent with this formulation, these effects are suppressed by epidermal growth factor, acting via the PI3Kdependent signaling pathway 89 that promotes proliferation and glycolysis while inhibiting fatty acid oxidation. A proposed anti-apoptotic role for in T cells72 and mammary epithelial cells81, while seemingly in contradiction to its numerous adverse effects on cell function, growth, and survival90, may reflect cell-specific responses, and/or activation by of a cell cycle arrest that in these settings would be protective, as discussed above (see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer).

6.4.

SALICYLATES AND OTHER NONSTEROIDAL ANTI-INFLAMMATORY DRUGS (NSAIDs)

Many agents in this broad category inhibit tumorigenesis. Thus, salicylates prevent human colon adenomas91-93 and carcinomas94, and Similarly, sulindac tumors of the colon95 and liver96 in rodents. metabolites prevent polyp formation in the murine model of familial adenomatous polyposis (FAP)97, and experimental breast cancer in rodents98; studies of polyp regression in FAP are conflicting99-101. In vitro, aspirin suppresses microsatellite instability in colorectal cancer cells102, while agents from the major NSAID classes cause p53independent growth arrest and apoptosis in several cell types and colon cancer-derived cell lines. Although these agents in general inhibit cyclooxygenase 2 (COX2) and thereby the synthesis of growthpromoting prostaglandins, they also appear to act independently of altered eicosanoid metabolism103-110, or in addition to it111,112. NSAIDs also impair mitochondrial function; effects include uncoupling of oxidative phosphorylation by salicylate (Fig. 6.2 – facing page 123), salicylate derivatives, and several other NSAIDs, in vivo or in vitro113-116. The salicylate uncoupling effect is associated with diminished 117 mitochondrial , and has been correlated with antirheumatic and anti-inflammatory potency113. Although not all NSAIDs uncouple oxidative phosphorylation, this effect warrants further consideration in the context of oxidative stress and cell proliferation (also, see Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). Intramitochondrial generation of ROS, i.e., one electron reduction of molecular with formation of superoxide anion is caused by electron “leakage” from the electron transport chain, especially at

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complexes I and III and ubiquinone (Fig. 6.3). Such “leakage” may be accelerated by excessive electron entry (e.g., as a result of increased mitochondrial fatty acid oxidation) and/or diminished electron removal as the result of impaired electron transport chain function (e.g., in hypoxia). In addition to intramitochondrial defenses that are immediately available, e.g., glutathione, superoxide dismutase, and thioredoxin118-120 , moderate levels of ROS formation may activate PI3K/Akt/PKB signaling (see Chapter 3.3.2.3.: Reactive Oxygen Species) and the transcriptional regulators and AP-1121-123. The latter lead to increased expression of several genes important in cell survival and redox regulation, which thus may synergize with PI3K/Akt/PKB. These include the inducible


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form of nitric oxide synthase (iNOS)124, COX2125, Bcl-2, Bcl-xL126, and the Bcl-2 homolog Bfl-1/A1127. ROS may also contribute to expression of mitochondrial uncoupling protein-2 (UCP2) in the posthepatectomy regenerating hepatocyte128,129. activation appears to be essential for cell survival and proliferation, e.g., in response to signaling130-134 (also, see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin), although under some conditions it may exert a pro-apoptotic effect135. There is abundant evidence that uncoupling of oxidative phosphorylation suppresses intramitochondrial oxidative stress20-25. Presumably, this reflects accelerated removal of electrons through formation of (Fig. 6.3) as the result of uncoupler-induced increases in matrix proton abundance; electron “leakage” and generation of at upstream electron transport chain sites are thereby diminished. This antioxidant effect is produced by diverse uncoupling agents including underivatized long chain fatty acids, xenobiotics such as salicylate (Fig. 6.2 – facing page 123), the increased proton permeability of fatty acid-enriched mitochondrial inner membrane136 (also, see Chapter 4.3.: Omega-3 Fatty Acids), the cardioprotective agent carvedilol25, and UCP2. Moreover, activates uncoupling proteins in a fatty acid-dependent 129 manner . As a result of its uncoupling and anti-oxidant effects, 137-139 salicylate also supresses ROS-induced activation of and AP104 1 , and therefore transcriptional activation of redox-related genes. As discussed above (Chapter 6.2.: Butyrate), the antioxidant effect is not produced by those agents such as butyrate1 and ketone bodies, that exert an uncoupling-like effect, because their oxidation in the mitochondrial matrix augments electron abundance and “leakage” in the electron transport chain rather than decreasing it, as do the non-oxidized protonophore uncouplers such as salicylate. While the uncoupling effects of salicylate are protective under certain conditions, they are necessarily linked to uncoupler-induced compromise of and, therefore, of ATP generation. This may account for the otherwise paradoxical observation that salicylate protects against apoptosis in resting cells138 in which ATP requirements are low, whereas in proliferating cells in which ATP requirements are high it predisposes to 139 spontaneous108,140 and apoptosis (see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer). It also augments cell killing by inducers of the permeability transition141. Although salicylate has also been shown to bind to and inhibit the 142-144 activator , thereby suppressing insulin resistance through as yet unknown mechanisms143,144, the significance of this effect remains uncertain145. Possibly, salicylate’s enhancement of insulin

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sensitivity reflects in part its uncoupling-induced suppression of intramitochondrial ROS generation, thus favoring activation of PDH and glycolysis. Accordingly, salicylate may influence cellular responses to proliferative, antiproliferative, or apoptogenic stimuli through two closely linked mechanisms. First, it diminishes and thus compromises ATP generation. Second, it diminishes intramitochondrial ROS generation and thus suppresses activation. Both effects can be understood as the consequences of salicylate’s action as a protonophore uncoupler, and both may predispose either to protective growth arrest or to apoptosis, depending on conditions. Importantly, this mechanism of suppression by salicylate contrasts with that of the more “downstream” inhibitory effect of noted above59. NSAIDs other than salicylates may not uncouple oxidative phosphorylation but instead inhibit the electron transport chain146,147; indomethacin may do either depending on its concentration116. Evidence pertaining to the mitochondrial effects of sulindac is conflicting115,148. Our preliminary experiments (Ockner R, unpublished data) indicate that sulindac differs from salicylate in that it does not uncouple oxidative phosphorylation but instead inhibits complex I (Fig. 6.2 – facing page 123). A resulting decrease in electron transport chain flux would decrease ROS generation, consistent with sulindac’s suppression of activation149 and COX2 expression150. Similar to butyrate and however, and despite their differing effects on mitochondria, both salicylate and sulindac increase long chain fatty acid oxidation and ketogenesis, absolutely and/or relative to maximal rates of ATP synthesis (Ockner R, unpublished data). Sulindac sulfide also may activate 151 . In contrast to these effects, certain NSAIDs may cause concentration-dependent inhibition of mitochondrial fatty acid oxidation152,153, thereby inducing peroxisome proliferator-like effects153-155, and possibly accounting for an otherwise unexpected proliferative effect on normal colonic crypt cells156. NSAIDs may also activate and/or directly or indirectly157,158. Consideration of the foregoing interactions may help to clarify what has been a puzzling aspect of the relationship of COX2 and NSAIDs to cell proliferation and tumorigenesis. Among the reaction products of COX2 are prostaglandins D1 and D2 (PGD), both of which directly or 159,160 indirectly activate , and PGE2 which stimulates tumor cell 161 growth and motility . As discussed above (see Chapter 5.2.1.: a critical aspect of the peroxisome proliferator effect is that it restricts mitochondrial fatty acid oxidation as it augments, via mitochondrial mass and oxidative capacity. As a result,


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intramitochondrial electron flux, reducing potential, and therefore ROS generation are suppressed162-166, thus resembling an insulin-like effect in which remains high. Conversely, is diminished under conditions in which uncoupling agents suppress oxygen radical formation and activation. This uncoupling-induced effect would account for decreased COX2 gene transcription by salicylate and aspirin167, and for suppression of and iNOS by fatty acids136,168 (also, see Chapter 4.3.: Omega-3 Fatty Acids). In contrast, a moderate increase in intramitochondrial ROS generation (e.g., caused under certain circumstances by proinflammatory cytokines - see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin) would activate and AP-1121-123, thereby increasing expression of redox-related genes including COX2 and, thus, formation of eicosanoid activators of 169 . and In this context, the response cascade — oxidative stress augmented mitochondrial oxidative capacity — could be viewed as a mechanism that subserves feedback control of intramitochondrial oxidative stress, while promoting mitochondrial redox conditions and energetics more favorable to cell proliferation. The cascade could be interrupted at any of several loci, including uncoupling-induced suppression of ROS generation and activation, compromise of direct inhibition of the electron transport chain, or direct inhibition of COX2. As a corollary, the cascade’s ability to stimulate cell proliferation would be compromised by agents which inhibit COX2 as well as by those which do not. This formulation would account for evidence that the antiproliferative effects of COX2-selective NSAIDS may be COX2-independent170,171, or COX2-dependent but independent of PGE2172, and that they enhance the antiproliferative effect of butyrate173. It is also compatible with alternative hypotheses174, and with recent evidence that NSAIDs may inhibit either eicosanoid activation of 169 , or the latter’s interaction with its DNA response elements175,176 (also, see Chapter 5.2.3.: Furthermore, it is likely that effects of NSAIDs on cell proliferation will depend on whether the eicosanoids synthesized by COX2 are predominantly pro-inflammatory Given these (proliferative) or anti-inflammatory (anti-proliferative)177. considerations, evidence that NSAID-induced gastric mucosal injury does not depend on COX-inhibitory potency or mucosal surface exposure170,178 suggests that it may be the consequence of an antiproliferative effect on gastric crypt progenitor cells. The present formulation, if valid, would also predict that in some circumstances activation of mitochondrial proliferation may be COX2-dependent.

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6.5.

REFERENCES

1. R. Ockner, N. Lysenko, N. Wu, and N. Bass, Hepatocyte growth inhibitors modulate mitochondrial and extramitochondrial fatty acid oxidation, Hepatology 24:253A (Abstract) (1996). 2. M. Bugaut, and M. Bentéjac, Biological effects of short-chain fatty acids in nonruminant mammals., Annu Rev Nutr 13:217-241 (1993). 3. A. Toscani, D. Soprano, and K. Soprano, Molecular analysis of sodium butyrateinduced growth arrest, Oncogene Res 3:223-238 (1988). 4. A. Hague, A. Manning, K. Hanlon, L. Huschtscha, D. Hart, and C. Paraskeva, Sodium butyrate induces apoptosis in human colonic tumour cell lines in a p53-independent pathway: implications for the possible role of dietary fibre in the prevention of largebowel cancer, Intl J Cancer 55:498-505 (1993). 5. R. Gope, and M. Gope, Effect of sodium butyrate on the expression of retinoblastoma (RB1) and P53 gene and phosphorylation of retinoblastoma protein in human colon tumor cell line HT29, Cell Mol Biol 39:589-597 (1993). 6. D. Heruth, G. Zirnstein, J. Bradley, and P. Rothberg, Sodium butyrate causes an increase in the block to transcriptional elongation in the c-myc gene in SW837 rectal carcinoma cells, J Biol Chem 268:20466-20472 (1993). 7. C. Vaziri, L. Slice, and D. Faller, Butyrate-induced G1 arrest results from p21independent disruption of retinoblastoma protein-mediated signals, Cell Growth Differentiation 9:465-474 (1998). 8. M. Barshishat, S. Polak-Charcon, and B. Schwartz, Butyrate regulates E-cadherin transcription, isoform expression and intracellular position in colon cancer cells., Br J Cancer 82:195-203 (2000). 9. K. Nakano, T. Mizuno, Y. Sowa, et al., Butyrate activates the WAF1/Cip1 gene promoter through Sp1 sites in a p53-negative human colon cancer cell line, J Biol Chem 272:22199-22206 (1997). 10. S. Archer, S. Meng, A. Shei, and R. Hodin, p21WAF1 is required for butyrate-mediated growth inhibition of human colon cancer cells., Proc Natl Acad Sci USA 95:67916796 (1998). 11. B. Heerdt, M. Houston, G. Anthony, and L. Augenlicht, Mitochondrial membrane potential DYm in the coordination of p53-independent proliferation and apoptosis pathways in human colonic carcinoma cells, Cancer Res 58:2869-2875 (1998). 12. B. Schwartz, C. Avivi-Green, and S. Polak-Charcon, Sodium butyrate induces retinoblastoma protein dephosphorylation, p16 expression and growth arrest of colon cancer cells, Molec Cellular Biochem 188:21-30 (1998). 13. A. Wächtershäuser, S. Loitsch, and J. Stein, PPAR-gamma is selectively upregulated in Caco-2 cells by butyrate, Biochem Biophys Res Comm 272:380-385 (2000). 14. J. McBain, A. Eastman, C. Nobel, and G. Mueller, Apoptotic death in adenocarcinoma cell lines induced by butyrate and other histone deacetylase inhibitors, Biochem Pharmacol 53:1357-1368 (1997). 15. P. Garland, E. Newsholme, and P. Randle, Regulation of glucose uptake by muscle: Effects of fatty acids and ketone bodies, and of alloxan-diabetes and starvation, on pyruvate metabolism and on lactate/pyruvate and L-glycerol 3phosphate/dihydroxyacetone phosphate concentration ratios in rat heart and rat diaphragm muscles., Biochem J 93:665-678 (1964). 16. T. A. Hopkins, M. C. Sugden, M. J. Holness, R. Kozak, J. R. Dyck, and G. D. Lopaschuk, Control of cardiac pyruvate dehydrogenase activity in peroxisome


133

proliferator-activated receptor-alpha transgenic mice, Am J Physiol Heart Circ Physiol 285:H270-6 (2003). 17. L. Wojtczak, and P. Schönfeld, Effect of fatty acids on energy coupling processes in mitochondria, Biochim Biophys Acta 1183:41-57 (1993). 18. L. Svensson, S. Kilpeläinen, J. Hiltunen, and S. Alexson, Characterization and isolation of enzymes that hydrolyze short-chain acyl-CoA in rat-liver mitochondria, Eur J Biochem 239:526-531 (1996). 19. J. Amatruda, S. Margolis, and D. Lockwood, Regulation of ketone body production from [14C]palmitate in rat liver mitochondria: effects of cyclic nucleotides and unlabeled fatty acids., Biochem Biophys Res Commun 67:1337-1345 (1975). 20. I. Reynolds, and T. Hastings, Glutamate induces the production of reactive oxygen species in cultured forebrain neurons following NMDA receptor activation, J Neurosci 15:3318-3327 (1995). 21. V. Skulachev, Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants, Quart Rev Biophys 29:169-202 (1996). 22. A. Nègre-Salvayre, C. Hirtz, G. Carrera, et al., A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation, FASEB J 11:809-815 (1997). 23. A. Stout, H. Raphael, B. Kanterewicz, E. Klann, and I. Reynolds, Glutamate-induced neuron death requires mitochondrial calcium uptake, Nature Neurosci 1:366-373 (1998). 24. S. Korshunov, O. Kokina, E. Ruuge, V. Skulachev, and A. Starkov, Fatty acids as natural uncouplers preventing generation of and by mitochondria in the resting state, FEBS Letts 435:215-218 (1998). 25. P. Oliveira, A. Rolo, V. Sardão, P. Coxito, C. Palmeira, and A. Moreno, Carvedilol in heart mitochondria protonophore or opener of the mitochondrial KATP channels?, Life Sci 69:123-132 (2001). 26. M. Tisdale, Role of acetoacetyl-CoA synthetase in acetoacetate utilization by tumor cells, Cancer Biochem Biophys 7:101-107 (1984). 27. S. Beck, and M. Tisdale, Effect of insulin on weight loss and tumour growth in a cachexia model., Br J Cancer 59:677-681 (1989). 28. W. Zhang, S. Churchill, R. Lindahl, and P. Churchill, Regulation of D-betahydroxybutyrate dehydrogenase in rat hepatoma cell lines, Cancer Res 49:2433-2437 (1989). 29. M. Mandal, X. Wu, and R. Kumar, Bcl-2 deregulation leads to inhibition of sodium butyrate-induced apoptosis in human colorectal carcinoma cells, Carcinogenesis 18:229-232 (1997). 30. Y. Fan, J. Zhang, R. Barhoumi, et al., Antagonism of CD95 signaling blocks butyrate induction of apoptosis in young adult mouse colonic cells., Am J Physiol 277:C310-C319 (1999). 31. M. Alam, T. Midtvedt, and A. Uribe, Differential cell kinetics in the ileum and colon of germfree rats., Scand J Gastroenterol 29:445-451 (1994). 32. C. Cherbuy, D.-V. B, M. M, and P. J , , P, Duée, Effect of germfree state on the capacities of isolated rat colonocytes to metabolize n-butyrate, glucose, and glutamine., Gastroenterol. 109: 1890-1899 (1995). 33. B. Singh, A. Halestrap, and C. Paraskeva, Butyrate can act as a stimulator of growth or inducer of apoptosis in human colonic epithelial cell lines depending on the presence of alternative energy sources, Carcinogenesis 18:1265-1270 (1997).

134

Chapter 6

34. O. Kan, S. Baldwin, and A. Whetton, Apoptosis is regulated by the rate of glucose transport in an interleukin 3 dependent cell line, J Exp Med 180:917-923 (1994). 35. Z. Kovacevic, and J. Mcgivan, Mitochondrial metabolism of glutamine and glutamate and its physiological significance, Physiol Rev 63:547-605 (1983). 36. R. Nakashima, M. Paggi, and P. Pedersen, Contributions of glycolysis and oxidative phosphorylation to adenosine 5’-triphosphate production in AS-30D hepatoma cells. Cancer Res 44:5702-5706 (1984). 37. M. Board, S. Humm, and E. Newsholme, Maximum activities of key enzymes of glycolysis, glutaminolysis, pentose phosphate pathway and tricarboxylic acid cycle in normal, neoplastic and suppressed cells., Biochem J 265:503-509 (1990). 38. V. Goossens, J. Grooten, and W. Fiers, The oxidative metabolism of glutamine: A modulator of reactive oxygen intermediate-mediated cytotoxicity of tumor necrosis factor in L929 fibrosarcoma cells, J Biol Chem 271:192-196 (1996). 39. K. Polyak, J. Kato, M. Solomon, et al., p27Kipl, a cyclin-Cdk inhibitor, links transforming growth factor-beta and contact inhibition to cell cycle arrest, Genes Dev 8:9-22 (1994). 40. C. Li, L. Suardet, and J. Little, Potential role of WAF1/Cip1/p21 as a mediator of TGF-beta cytoinhibitory effect, J Biol Chem 270:4971-4974 (1995). 41. M. Datto, Y. Li, J. Panus, D. Howe, Y. Xiong, and X. Wang, Transforming growth factor beta induces the cyclin-dependent kinase inhibitor p21 through a p53independent mechanism., Proc Natl Acad Sci USA 92:5545-5549 (1995). 42. N. Muller, A. Reinacher-Schick, S. Baldus, et al., Smad4 induces the tumor suppressor E-cadherin and P-cadherin in colon carcinoma cells, Oncogene 21:6049-58 (2002). 43. U. Kintscher, S. Wakino, D. Bruemmer, et al., TGF-beta(1) induces peroxisome proliferator-activated receptor gammal and gamma2 expression in human THP-1 monocytes, Biochem Biophys Res Commun 297:794-9 (2002). 44. A. Pellacani, P. Wiesel, S. Razavi, et al., Down-regulation of high mobility groupI(Y) protein contributes to the inhibition of nitric-oxide synthase 2 by transforming growth factor-beta1, J Biol Chem 276:1653-1659 (2001). 45. B. Gumbiner, Carcinogenesis: a balance between beta-catenin and APC, Current Biol 7:R443-R446 (1997). 46. A. Yap, W. Brieher, and B. Gumbiner, Molecular and functional analysis of cadherinbased adherens junctions, Ann Rev Cell Dev Biol 13:119-146 (1997). 47. J. Francis, C. Heyworth, E. Spooncer, A. Pierce, T. Dexter, and A. Whetton, Transforming growth factor-beta 1 induces apoptosis independently of p53 and selectively reduces expression of Bcl-2 in multipotent hematopoietic cells., J Biol Chem 275:39137-39145 (2000). 48. B. Herrera, M. Fernandez, A. Alvarez, et al.. Activation of caspases occurs downstream from radical oxygen species production, Bcl-xL down-regulation, and early cytochrome C release in apoptosis induced by transforming growth factor beta in rat fetal hepatocytes, Hepatology 34:548-56 (2001A). 49. B. Kim, M. Mamura, K. Choi, B. Calabretta, and S. Kim, Transforming growth factor beta 1 induces apoptosis through cleavage of BAD in a Smad3-dependent mechanism in FaO hepatoma cells, Mol Cell Biol 22:1369-1378 (2002). 50. Y. Hong, H. Peng, F. La, and V. Liao, Hydrogen peroxide-mediated transcriptional induction of macrophage colony-stimulating factor by TGF-beta1, J Immun 159:2418-2423 (1997). 51. B. Herrera, A. M. Alvarez, A. Sanchez, et al., Reactive oxygen species (ROS) mediates the mitochondrial-dependent apoptosis induced by transforming growth factor (beta) in fetal hepatocytes, Faseb J 15:741-51 (2001B).


135

52. G. Blobe, W. Schiemann, and H. Lodish, Role of transforming growth factor b in human disease., New Engl J Med 342:1350-1358 (2000). 53. R. Akhurst, and R. Derynck, TGF-beta signaling in cancer - a double-edged sword., Trends Cell Biol 11:S44-S51 (2001). 54. L. Wakefield, and A. Roberts, TGF-beta signaling: positive and negative effects on tumorigenesis, Curr Opin Genet Dev 12:22-29 (2002). 55. Y. Shi, and J. Massague, Mechanisms of TGF-beta signaling from cell membrane to the nucleus, Cell 113:685-700 (2003). 56. M. Cordenonsi, S. Dupont, S. Maretto, A. Insinga, C. Imbriano, and S. Piccolo, Links between tumor suppressors: p53 is required for TGF-beta gene responses by cooperating with Smads, Cell 113:301-14 (2003). 57. M. Miyazaki, R. Ohashi, T. Tsuji, K. Mihara, E. Gohda, and M. Namba, Transforming growth factor-beta 1 stimulates or inhibits cell growth via down- or up-regulation of p21/Waf1, Biochem Biophys Res Comm 246:873-880 (1998). 58. A. Mauviel, K. Chung, A. Agarwal, K. Tamai, and J. Uitto, Cell-specific induction of distinct oncogenes of the Jun family is responsible for differential regulation of collagenase gene expression by transforming growth factor-beta in fibroblasts and keratinocytes, J Biol Chem 271:10917-10923 (1996). 59. M. Azuma, K. Motegi, K. Aota, T. Yamashita, H. Yoshida, and M. Sato, TGF-beta1 inhibits NF-kappaB activity through induction of IkappaB-alpha expression in human salivary gland cells: a possible mechanism of growth suppression by TGF-betal., Exp Cell Res 250:213-222 (1999). 60. K. Matsuzaki, M. Date, F. Furukawa, et al., Regulatory mechanisms for transforming growth factor _ as an autocrine inhibitor in human hepatocellular carcinoma: implications for roles of smads in its growth, Hepatol 32:218-227 (2000). 61. M. Macias-Silva, W. Li, J. I. Leu, M. A. Crissey, and R. Taub, Up-regulated transcriptional repressors SnoN and Ski bind Smad proteins to antagonize transforming growth factor-beta signals during liver regeneration, J Biol Chem 277:28483-90 (2002). 62. M. Delcommenne, C. Tan, V. Gray, L. Rue, J. Woodgett, and S. Dedhar, Phosphoinositide-3-OH kinase-dependent regulation of glycogen synthase kinase 3 and protein kinase B/AKT by the integrin-linked kinase., Proc Natl Acad Sci USA 95:11211-11216 (1998). 63. Y. Li, J. Yang, C. Dai, C. Wu, and Y. Liu, Role for integrin-linked kinase in mediating tubular epithelial to mesenchymal transition and renal interstitial fibrogenesis, J Clin Invest 112:503-16 (2003). 64. A. Bakin, A. Tomlinson, N. Bhowmick, H. Moses, and C. Arteaga, Phosphatidylinositol 3-kinase function is required for transforming growth factor bmediated epithelial to mesenchymal transition and cell migration., J Biol Chem 275:36803-36810 (2000). 65. F. Valdes, A. M. Alvarez, A. Locascio, et al., The epithelial mesenchymal transition confers resistance to the apoptotic effects of transforming growth factor beta in fetal rat hepatocytes, Mol Cancer Res 1:68-78 (2002). 66. Y. Vodovotz, L. Chesler, H. Chong, et al., Regulation of transforming growth factor beta1 by nitric oxide, Cancer Res 59:2142-2149 (1999). 67. Y. Vodovotz, C. Bogdan, J. Paik, Q. Xie, and C. Nathan, Mechanisms of suppression of macrophage nitric oxide release by transforming growth factor beta, J Exp Med 178:605-613 (1993).

136

Chapter 6

68. U. Heine, J. Burmester, K. Flanders, et al., Localization of transforming growth factor-beta 1 in mitochondria of murine heart and liver, Cell Regul 2:467-477 (1991). 69. D. Anderson, C. Guerin, G. Hageman, B. Pfeffer, and K. Flanders, Distribution of transforming growth factor-beta isoforms in the mammalian retina., J Neurosci Res 42:63-79 (1995). 70. J. Potts, S. Bassnett, and D. Beebe, Expression of transforming growth factor beta in the embryonic avian lens coincides with the presence of mitochondria, Developmental Dynamics 203:317-323 (1995). 71. S. Roth-Eichhorn, K. Kühl, and A. Gressner, Subcellular localization of (latent) transforming growth factor beta and the latent TGF-beta binding protein in rat hepatocytes and hepatic stellate cells, Hepatology 28:1588-1596 (1998). 72. W. Chen, W. Jin, H. Tian, et al., Requirement for transforming growth factor beta1 in controlling T cell apoptosis., J Exp Med 194:439-53 (2001). 73. K. Flanders, N. Thompson, D. Cissel, et al., Transforming growth factor-beta 1: histochemical localization with antibodies to different epitopes., J Cell Biol 108:653-660 (1989). 74. S. Kim, A. Glick, M. Sporn, and A. Roberts, Characterization of the promoter region of the human transforming growth factor-beta 1 gene, J Biol Chem 264:402-408 (1989). 75. T. Caver, F. O’Sullivan, L. Gold, and H. Gresham, Intracellular demonstration of active TGFbeta1 in B cells and plasma cells of autoimmune mice: IgG-Bound TGFbeta1 suppresses neutrophil function and host defense against staphylococcus aureus infection., J Clin Invest 98:2496-2506 (1996). 76. B. Grasl-Kraupp, E. Schausberger, K. Hufnagl, et al., A novel mechanism for mitogenic signaling via pro-transforming growth factor alpha within hepatocyte nuclei, Hepatology 35:1372-80 (2002). 77. M. Joaquin, J. Rosa, C. Salvado, et al., Hepatocyte growth factor and transforming growth factor beta regulate 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase gene expression in rat hepatocyte primary cultures, Biochem J 314:235-240 (1996). 78. G. Thoresen, and T. Christoffersen, Transforming growth factor beta 1 increases the phosphoenolpyruvate carboxykinase mRNA level in cultured rat hepatocytes, Cell Biol Intl 18:171-175 (1994). 79. H. Yamazaki, M. Arai, S. Matsumura, K. Inoue, and T. Fushiki, Intracranial administration of transforming growth factor-beta3 increases fat oxidation in rats, Am J Physiol Endocrinol Metab 283:E536-44 (2002). 80. Y. Yamamura, X. Hua, S. Bergelson, and H. Lodish, Critical role of smads and AP-1 complex in transforming growth factor-b-dependent apoptosis, J Biol Chem 275:36295-36302 (2000). 81. I. Shin, A. V. Bakin, U. Rodeck, A. Brunet, and C. L. Arteaga, Transforming growth factor beta enhances epithelial cell survival via Akt-dependent regulation of FKHRL1, Mol Biol Cell 12:3328-39 (2001). 82. C. Diez-Fernandez, D. Andres, and M. Cascales, Attenuating effects of heat shock against TGF-beta1-induced apoptosis in cultured rat hepatocytes, Free Radic Biol Med 33:835-46 (2002). 83. C. M. Rodrigues, X. Ma, C. Linehan-Stieers, G. Fan, B. T. Kren, and C. J. Steer, Ursodeoxycholic acid prevents cytochrome c release in apoptosis by inhibiting mitochondrial membrane depolarization and channel formation, Cell Death Differ 6:842-54 (1999).


137

84. T. Y. Ling, Y. H. Huang, M. C. Lai, S. S. Huang, and J. S. Huang, Fatty acids modulate transforming growth factor-beta activity and plasma clearance, Faseb J 17:1559-61 (2003). 85. F. N. Ziyadeh, B. B. Hoffman, D. C. Han, et al., Long-term prevention of renal insufficiency, excess matrix gene expression, and glomerular mesangial matrix expansion by treatment with monoclonal antitransforming growth factor-beta antibody in db/db diabetic mice, Proc Natl Acad Sci U S A 97:8015-20 (2000). 86. M. Fujimoto, Y. Maezawa, K. Yokote, et al., Mice lacking Smad3 are protected against streptozotocin-induced diabetic glomerulopathy, Biochem Biophys Res Commun 305:1002-7 (2003). 87. M. Isono, A. Mogyorosi, D. C. Han, B. B. Hoffman, and F. N. Ziyadeh, Stimulation of TGF-beta type II receptor by high glucose in mouse mesangial cells and in diabetic kidney, Am J Physiol Renal Physiol 278:F830-8 (2000). 88. Y. Shima, K. Nakao, T. Nakashima, et al., Activation of caspase-8 in transforming growth factor-beta-induced apoptosis of human hepatoma cells, Hepatol 30:12151222 (1999). 89. I. Fabregat, B. Herrera, M. Fernández, et al., Epidermal growth factor impairs the cytochrome C-caspase-3 apoptotic pathway induced by transforming growth factor beta in rat fetal hepatocyctes via a phosphoinositide 3-kinase-dependent pathway., Hepatol 32:528-535 (2000). 90. P. Golstein, and A. Wyllie, T cell death and transforming growth factor betal, J Exp Med 194:F19-F21 (2001). 91. N. Shaheen, W. Straus, and R. Sandler, Chemoprevention of gastrointestinal malignancies with nonsteroidal antiinflammatory drugs, Cancer 94:950-963 (2002). 92. R. S. Sandler, S. Halabi, J. A. Baron, et al., A randomized trial of aspirin to prevent colorectal adenomas in patients with previous colorectal cancer, N Engl J Med 348:883-90 (2003). 93. J. A. Baron, B. F. Cole, R. S. Sandier, et al., A randomized trial of aspirin to prevent colorectal adenomas, N Engl J Med 348:891-9 (2003). 94. E. Giovannucci, K. Egan, D. Hunter, et al., Aspirin and the risk of colorectal cancer in women, New Engl J Med 333:609-614 (1995). 95. A. Paganini-Hill, Aspirin and the prevention of colorectal cancer: a review of the evidence, Semin Surg Oncol 10:158-164 (1994). 96. A. Denda, Q. Tang, T. Endoh, et al., Prevention by acetylsalicylic acid of liver cirrhosis and carcinogenesis as well as generations of 8-hydroxydeoxyguanosine and thiobarbituric acid-reactive substances caused by a choline-deficient, L-amino aciddefined diet in rats., Carcinogenesis 15:1279-1283 (1994). 97. N. Mahmoud, S. Boolbol, A. Dannenberg, et al., The sulfide metabolite of sulindac prevents tumors and restores enterocyte apoptosis in a murine model of familial adenomatous polyposis, Carcinogenesis 19:87-91 (1998A). 98. H. Thompson, C. Jiang, J. Lu, et al., Sulfone metabolite of sulindac inhibits mammary carcinogenesis, Cancer Res 57:267-271 (1997). 99. P. Pasricha, A. Bedi, K. O’Connor, et al., The effects of sulindac on colorectal proliferation and apoptosis in familial adenomatous polyposis, Gastroenterology 109:994-998 (1995). 100. M. Cruz-Correa, L. Hylind, K. Romans, S. Booker, and F. Giardiello, Long-term treatment with sulindac in familial adenomatous polyposis: a prospective cohort study., Gastroenterology 122:641-645 (2002).

138

Chapter 6

101. F. Giardiello, V. Yang, L. Hylind, et al., Primary chemoprevention of familial adenomatous polyposis with sulindac, N Engl J Med 346:1054-1059 (2002). 102. J. Rüschoff, S. Wallinger, W. Dietmaier, et al., Aspirin suppresses the mutator phenotype associated with hereditary nonpolyposis colorectal cancer by genetic seletion, Proc Natl Acad Sci USA 95:11301-11306 (1998). 103. R. Hanif, A. Pittas, Y. Feng, et al., Effects of nonsteroidal anti-inflammatory drugs on proliferation and on induction of apoptosis in colon cancer cells by a prostaglandin-independent pathway, Biochem Pharmacol 52:237-245 (1996). 104. Z. Dong, C. Huang, R. Brown, and W. Ma, Inhibition of activator protein 1 activity and neoplastic transformation by aspirin., J Biol Chem 272:9962-9970 (1997). 105. G. Levy, Prostaglandin H synthases, nonsteroidal anti-inflammatory drugs, and colon cancer, FASEB J 11:234-247 (1997). 106. G. Piazza, A. Rahm, T. Finn, et al., Apoptosis primarily accounts for the growthinhibitory properties of sulindac metabolites and involves a mechanism that is independent of cyclooxygenase inhibition, cell cycle arrest, and p53 induction. Cancer Res 57:2452-2459 (1997). 107. D. Elder, and C. Paraskeva, NSAIDS to prevent colorectal cancer: A question of sensitivity., Gastroenterol 113:1999-2008 (1997). 108. B. Bellosillo, M. Piqué, M. Barragán, et al., Aspirin and salicylate induce apoptosis and activation of caspases in B-cell chronic lymphocytic leukemia cells., Blood 92:1406-1414 (1998). 109. E.-H. Han, N. Arber, H. Yamamoto, et al., Effects of sulindac and its metabolites on growth and apoptosis in human mammary epithelial and breast carcinoma cell lines. Breast Cancer Res Treatment 48:195-203 (1998). 110. S. Shiff, and B. Rigas, The role of cyclooxygenase inhibition in the antineoplastic effects of nonsteroidal antiinflammatory drugs (NSAIDs), J Exper Med 190:445-450 (1999). 111. N. Mahmoud, A. Dannenberg, J. Mestre, et al., Aspirin prevents tumors in a murine model of familial adenomatous polyposis, Surgery 124:225-231 (1998B). 112. R. Gupta, and R. Dubois, Aspirin, NSAIDs, and colon cancer prevention mechanisms?, Gastroenterol 114:1095-1098 (1998). 113. M. Whitehouse, Uncoupling of oxidative phosphorylation in a connective tissue (cartilage) and liver mitochondria by salicylate analogues: Relationship of structure to activity, Biochem Pharmacol 13:319-336 (1964). 114. M. Mehlman, R. Tobin, and E. Sporn, Oxidative phosphorylation and respiration by rat liver mitochondria from aspirin-treated rats, Biochem Pharmacol 21:32793285 (1972). 115. T. Mahmud, S. Rafi, D. Scott, J. Wrigglesworth, and I. Bjarnason, Nonsteroidal antiinflammatory drugs and uncoupling of mitochondrial oxidative phosphorylation. Arthritis Rheum 39:1998-2003 (1996). 116. M. Jacob, I. Bjarnason, and R. J. Simpson, Effects of indomethacin on energy metabolism in rat jejunal tissue in vivo, Clin Sci (Lond) 102:541-6 (2002). 117. R. Haas, W. Parker, Jr, D. Stumpf, and L. Eguren, Salicylate-induced loose couplinga; protonmotive force measurements, Biochem Pharmacol 34:900-902 (1985). 118. Y. Chen, J. Cai, T. J. Murphy, and D. P. Jones, Overexpressed human mitochondrial thioredoxin confers resistance to oxidant-induced apoptosis in human osteosarcoma cells, J Biol Chem 277:33242-8 (2002).


139

119. A. E. Damdimopoulos, A. Miranda-Vizuete, M. Pelto-Huikko, J. A. Gustafsson, and G. Spyrou, Human mitochondrial thioredoxin. Involvement in mitochondrial membrane potential and cell death, J Biol Chem 277:33249-57 (2002). 120. T. Tanaka, F. Hosoi, Y. Yamaguchi-Iwai, et al., Thioredoxin-2 (TRX-2) is an essential gene regulating mitochondria-dependent apoptosis, Embo J 21:1695-703 (2002). 121. C. Sen, and L. Packer, Antioxidant and redox regulation of gene transcription, FASEB J 10:709-720 (1996). 122. V. Lakshminarayanan, E. Drab-Weiss, and K. Roebuck, and tumor necrosis factor-alpha induce differential binding of the redox-responsive transcription factors AP-1 and NF-kappaB to the interleukin-8 promoter in endothelial and epithelial cells, J Biol Chem 273:32670-32678 (1998). 123. E. Shaulian, and M. Karin, AP-1 as a regulator of cell life and death, Nat Cell Biol 4:E131-6 (2002). 124. T. Vos, H. Van Goor, L. Tuyt, et al., Expression of inducible nitric oxide synthase in endotoxemic rat hepatocytes is dependent on the cellular glutathione status, Hepatology 29:421-426 (1999). 125. K. Yamamoto, T. Arakawa, N. Ueda, and S. Yamamoto, Transcriptional roles of nuclear factor kappa B and nuclear factor-interleukin-6 in the tumor necrosis factor alpha-dependent induction of cyclooxygenase-2 in MC3T3-E1 cells, J Biol Chem 270:31315-31320 (1995). 126. M. Tamatani, Y. Che, H. Matsuzaki, et al., Tumor necrosis factor induces Bcl-2 and Bcl-x expression through NFkappaB activation in primary hippocampal neurons, J Biol Chem 274:8531-8538 (1999). 127. W. Zong, L. Edelstein, C. Chen, J. Bash, and C. Gé1inas, The prosurvival Bcl-2 homolog Bfl-2/A1 is a direct transcriptional target of NF-kappaB that blocks TNFalpha-induced apoptosis, Genes Development 13:382-387 (1999). 128. F. Lee, Y. Li, H. Zhu, et al., Tumor necrosis factor increases mitochondrial oxidant production and induces expression of uncoupling protein-2 in the regenerating mouse liver, Hepatology 29:677-687 (1999). 129. K. Echtay, D. Roussel, J. St-Pierre, et al., Superoxide activates mitochondrial uncoupling proteins., Nature 415:96-99 (2002). 130. A. Beg, and D. Baltimore, An essential role for NF-kappaB in preventing TNFalpha-induced cell death., Science 274:782-784 (1996). 131. C.-Y. Wang, M. Mayo, and J. Baldwin, AS TNF- and cancer therapy-induced apoptosis: potentiation by inhibition of NF-kappaB, Science 274:784-787 (1996). 132. D. Van Antwerp, S. Martin, T. Kafri, D. Green, and I. Verma, Suppression of TNFalpha-induced apoptosis by NF-kappaB, Science 274:787-789 (1996). 133. M. Karin, and A. Lin, NF-kappaB at the crossroads of life and death, Nat Immunol 3:221-227 (2002). 134. O. Micheau, and J. Tschopp, Induction of TNF receptor I-mediated apoptosis via two sequential signaling complexes, Cell 114:181-90 (2003). 135. N. Perkins, The Rel/NF-kappa B family: friend and foe, Trends Biochem Sci 25:434440 (2000). 136. D. Rolfe, A. Hulbert, and M. Brand, Characteristics of mitochondrial proton leak and control of oxidative phosphorylation in the major oxygen-consuming tissues of the rat, Biochim Biophys Acta 1188:405-416 (1994). 137. E. Kopp, and S. Ghosh, Inhibition of NF-kappa B by sodium salicylate and aspirin, Science 265:956-959 (1994).

140

Chapter 6

138. M. Grilli, M. Pizzi, M. Memo, and P. Spano, Neuroprotection by aspirin and sodium salicylate through blockade of NF-kappaB activation, Science 274:1383-1385 (1996). 139. T. McDade, R. Perugini, F. Vittimberga, Jr, R. Carrigan, and M. Callery, Salicylates inhibit NF-kappaB activation and enhance TNF-alpha-induced apoptosis in human pancreatic cancer cells, J Surg Res 83:56-61 (1999). 140. L. Klampfer, J. Cammenga, H. Wisniewski, and S. Nimer, Sodium salicylate activates caspases and induces apoptosis of myeloid leukemia cell lines, Blood 93:2386-2394 (1999). 141. K. W. Oh, T. Qian, D. A. Brenner, and J. J. Lemasters, Salicylate enhances necrosis and apoptosis mediated by the mitochondrial permeability transition, Toxicol Sci 73:44-52 (2003). 142. M. Yin, Y. Yamamoto, and R. Gaynor, The anti-inflammatory agents aspirin and salicylate inhibit the activity of I(kappa)B kinase-beta, Nature 396:77-80 (1998). 143. J. Kim, Y. Kim, J. Fillmore, et al., Prevention of fat-induced insulin resistance by salicylate, J Clin Invest 108:437-446 (2001). 144. M. Yuan, N. Konstantopoulos, J. Lee, et al, Reversal of obesity- and diet-induced insulin resistance with salicylates or targeted disruption of Ikkbeta, Science 293:1673-1677 (2001). 145. G. Jiang, Q. Dallas-Yang, F. Liu, D. E. Moller, and B. B. Zhang, Salicylic acid reverses phorbol 12-myristate-13-acetate (PMA)- and tumor necrosis factor alpha (TNFalpha)-induced insulin receptor substrate 1 (IRS1) serine 307 phosphorylation and insulin resistance in human embryonic kidney 293 (HEK293) cells, J Biol Chem 278:180-6 (2003). 146. Y. Tokumitsu, S. Lee, and M. Ui, In vitro effects of nonsteroidal anti-inflammatory drugs on oxidative phosphorylation in rats liver mitochondria, Biochem Pharmacol 26:2101-2106 (1977). 147. K. Kawai, H. Shiojiri, H. Fukushima, and Y. Nozawa, The inhibition of mitochondrial respiration by indomethacin, a non-steroid anti-inflammatory agent possessing inhibitory effect on prostaglandin biosynthesis, Res Commun Chem Pathol Pharmacol 48:267-274 (1985). 148. P. McDougall, A. Markham, I. Cameron, and A. Sweetman, The mechanism of inhibition of mitochondrial oxidative phosphorylation by the nonsteroidal antiinflammatory agent diflunisal, Biochem Pharmacol 32:2595-2598 (1983). 149. Y. Yamamoto, M. Yin, K. Lin, and R. Gaynor, Sulindac inhibits activation of the NF-kappaB pathway, J Biol Chem 274:27307-27314 (1999). 150. M. Taylor, K. Lawson, N. Ignatenko, et al., Sulindac sulfone inhibits K-rasdependent cyclooxygenase-2 expression in human colon cancer cells, Cancer Res 60:6607-6610 (2000). 151. M. Wick, G. Hurteau, C. Dessev, et al., Peroxisome proliferator-activated receptorgamma is a target of nonsteroidal anti-inflammatory drugs mediating cyclooxygenaseindependent inhibition of lung cancer cell growth, Mol Pharmacol 62:1207-14 (2002). 152. E. Freneaux, B. Fromenty, A. Berson, et al., Stereoselective and nonstereoselective effects of ibuprofen enantiomers on mitochondrial beta-oxidation of fatty acids., J Pharmacol Exp Ther 255:529-535 (1990). 153. R. Kundu, G. Getz, and J. Tonsgard, Induction of (omega-1)-oxidation of monocarboxylic acids by acetylsalicylic acid, J Lipid Res 34:1187-1199 (1993).


141

154. Y. Cai, A. Sohlenius, K. Andersson, C. Sundberg, and J. Depierre, Effects of acetylsalicylic acid on parameters related to peroxisome proliferation in mouse liver., Biochem Pharmacol 47:2213-2219 (1994). 155. E. Rekka, E. Ayalogu, D. Lewis, G. Gibson, and C. Ioannides, Induction of hepatic microsomal CYP4A activity and of peroxisomal beta-oxidation by two non-steroidal anti-inflammatory drugs, Arch Toxicol 68:73-78 (1994). 156. P. Craven, K. Thornburg, and F. Derubertis, Sustained increase in the proliferation of rat colonic mucosa during chronic treatment with aspirin., Gastroenterology 94:567-575 (1988). 157. J. Lehmann, J. Lenhard, B. Oliver, G. Ringold, and S. Kliewer, Peroxisome proliferator-activated receptors alpha and gamma are activated by indomethacin and other non-steroidal anti-inflammatory drugs, J Biol Chem 272:3406-3410 (1997). 158. M. Jaradat, B. Wongsud, S. Phornchirasilp, et al., Activation of peroxisome proliferator-activated receptor isoforms and inhibition of prostaglandin H(2) synthases by ibuprofen, naproxen, and indomethacin, Biochem Pharmacol 62:15871595 (2001). 159. B. Forman, J. Chen, and R. Evans, Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-activated receptors alpha and delta., Proc Natl Acad Sci USA 94:4312-4317 (1997). 160. K. Yu, W. Bayona, C. Kallen, et al., Differential activation of peroxisome proliferator-activated receptors by eiconsanoids, J Biol Chem 270:23975-23983 (1995). 161. H. Sheng, J. Shao, M. Washington, and R. DuBois, Prostaglandin E2 increases growth and motility of colorectal carcinoma cells, J Biol Chem 276:18075-18081 (2001). 162. C. Prager, H. Schön, M. Nikfardjam, et al., Clofibrate elevates enzyme activities of the tricarboxylic acid cycle in rat liver, J Lipid Res 34:359-364 (1993). 163. B. Wolfe, J. Kane, R. Havel, and H. Brewster, Mechanism of the hypolipemic effect of clofibrate in postabsorptive man, J Clin Invest 52:2146-2159 (1973). 164. M. Laker, and P. Mayes, The immediate and long term effects of clofibrate on the metabolism of the perfused rat liver, Biochem Pharmacol 28:2813-2827 (1979). 165. K. Yamamoto, N. Fukuda, L. Zhang, and T. Sakai, Altered hepatic metabolism of fatty acids in rats fed a hypolipidaemic drug, fenofibrate, Pharmacol Res 33:337-342 (1996). 166. M. Poynter, and R. Daynes, Peroxisome proliferator-activated receptor alpha activation modulates cellular redox status, represses nuclear factor-kappaB signaling, and reduces inflammatory cytokine production in aging, J Biol Chem 273:3283332841 (1998). 167. X. Xu, L. Sansores-Garcia, X. Chen, N. Matijevic-Aleksic, M. Du, and K. Wu, Suppression of inducible cyclooxygenase 2 gene transcription by aspirin and sodium salicylate, Proc Natl Acad Sci USA 96:5292-5297 (1999). 168. B. A. Narayanan, N. K. Narayanan, B. Simi, and B. S. Reddy, Modulation of inducible nitric oxide synthase and related proinflammatory genes by the omega-3 fatty acid docosahexaenoic acid in human colon cancer cells, Cancer Res 63:972-9 (2003). 169. R. Gupta, J. Tan, W. Krause, et al., Prostacyclin-mediated activation of peroxisome proliferator-activated receptor in colorectal cancer, Proc Natl Acad Sci USA 97:13275-13280 (2000).

142

Chapter 6

170. M. Smith, G. Hawcroft, and M. Hull, The effect of non-steroidal anti-inflammatory drugs on human colorectal cancer cells: evidence of different mechanisms of action, Eur J Cancer 36:664-674 (2000). 171. S. Arico, S. Pattingre, C. Bauvy, et al., Celecoxib induces apoptosis by inhibiting 3-phosphoinositide-dependent protein kinase-1 activity in the human colon cancer HT-29 cell line, J Biol Chem 277:27613-21 (2002). 172. G. Sigthorsson, R. J. Simpson, M. Walley, et al., COX-1 and 2, intestinal integrity, and pathogenesis of nonsteroidal anti-inflammatory drug enteropathy in mice, Gastroenterology 122:1913-23 (2002). 173. T. Crew, D. Elder, and C. Paraskeva, A cyclooxygenase-2 (COX-2) selective nonsteroidal anti-inflammatory drug enhances the growth inhibitory effect of butyrate in colorectal carcinoma cells expressing COX-2 protein: regulation of COX-2 by butyrate., Carcinogenesis 21:69-77 (2000). 174. B. Rigas, and S. Shiff, Is inhibition of cyclooxygenase required for the chemopreventive effect of NSAIDs in colon cancer? A model reconciling the current contradiction, Med Hypotheses 54:210-215 (2000). 175. T. He, T. Chan, B. Vogelstein, and K. Kinzler, PPARdelta is an APC-regulated target of nonsteroidal anti-inflammatory drugs, Cell 99:335-345 (1999). 176. J. Peters, S. Lee, W. Li, et al., Growth, adipose, brain, and skin alterations resulting from targeted disruption of the mouse peroxisome proliferator-activated receptor beta(delta), Molec Cell Biol 20:5119-5128 (2000B). 177. B. Poligone, and A. Baldwin, Positive and negative regulation of NF-kappa B by COX-2: Roles of different prostaglandins, J Biol Chem 276:38658-38664 (2001). 178. J. Scheiman, and G. Elta, Gastroduodenal mucosal damage with salsalate versus aspirin: results of experimental models and endoscopic studies in humans, Semin Arthritis Rheumatism 20:121-127 (1990).

Chapter 7 Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications

7.1.

INTRODUCTION

At various points and in several contexts in the discussion thus far, the potentially adverse effects of excessive mitochondrial fatty acid oxidation have been emphasized, especially in terms of their possible influence on cell proliferation and oxidative stress. The intent of the present chapter is to summarize these effects and, in particular, to develop more fully the biochemical and metabolic foundations of their pathophysiologic and clinical significance. Although certain of the concepts are to some extent hypothetical, they are supported directly and indirectly by a substantial body of evidence. As a result, they may provide insight into, and contribute to the elucidation of, an unexpectedly diverse array of incompletely understood scientific and clinical issues.

7.2.

ADVERSE EFFECTS OF FATTY ACIDS ON MITOCHONDRIAL FUNCTION AND CELL REDOX BALANCE

Butyrate, and certain NSAIDs increase mitochondrial fatty acid oxidation, absolutely and/or relative to ATP biosynthesis. These effects are thus similar to those of medium chain and fatty acids, but they act by diverse mechanisms. The changes caused by these agents, or by other factors which augment fatty acid oxidation — such as increased plasma [FFA] associated with obesity, insulin resistance, diabetes mellitus, 143

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starvation, or HIV infection — may adversely affect mitochondrial function in several ways: 1) Fatty acids1-12 and obesity13 inhibit PI3K-mediated insulin (wg/Wntlike) signaling; increased tissue lipid abundance and mitochondrial fatty acid oxidation inhibit glucose uptake, glycolysis, and pyruvate dehydrogenase activity 7,10,14-16. 2) Fatty acids in excess may override regulatory control of CPT-I by malonyl CoA10,17-20, or abrogate such control by inducing expression of malonyl CoA decarboxylase21. The resulting augmentation of fatty acid predisposes to electron transport chain overload, ROS formation, and mitochondrial thiol depletion, i.e., oxidative stress. Consistent with this metabolic predisposition to conditions that are conducive to apoptosis, caspases are activated in intact cells and in a cellfree system by conditions favoring mitochondrial fatty acid oxidation22. In addition, peroxisomal of fatty acids may be augmented by excess fatty acids generally, as well as by those specific fatty acids that are inherently less readily oxidized in mitochondria, such as trans fatty acids23-25 (also, see this Chapter 7.4.1.: Atherosclerosis and Arterial Hypertension). 3) Fatty acids may disrupt VDAC and membrane contact sites, and favor opening of the permeability transition pore26,27. 4) Saturated fatty acids may be more likely than monounsaturated fatty acids to enter oxidative pathways and induce lipotoxicity in cultured cells28. Excesses in cytoplasmic palmitate or stearate also increase de novo (sphingomyelinase-independent) biosynthesis of ceramide, and apoptosis, in murine hematopoietic cell lines29. 5) Underivatized long chain fatty acids uncouple oxidative phosphorylation directly30-34. Fatty acids may also uncouple indirectly, through fatty acid increased expression of UCP235,36, or as the result of increased inner membrane proton permeability37 induced by fatty acid-enriched membrane lipids. While the uncoupling effect of underivatized long chain fatty acids tends to mitigate the generation and consequences of ROS38-42, the latter dominates at higher rates of fatty acid oxidation. 6) In addition to direct mitochondrial effects, excess fatty acids may enter and induce ROS-generating extramitochondrial fatty acid oxidation pathways, i.e., peroxisomal and and oxidation in the SER. These and other observations suggest that for any given cell, its position and direction in the broad spectrum of growth regulation — from proliferation to growth arrest to apoptosis — is substantially influenced by mitochondrial function and substrate utilization. Thus,

7. Fatty Acids and Mitochondria, Cell Growth and Injury

145

mitochondrial fatty acid oxidation and ketogenesis are: 1) suppressed in dividing cells; 2) inhibited by structurally diverse mitogens and growth promoters such as insulin43, (reflecting activation of the PI3K/Akt/PKB survival pathway), and the peroxisome proliterators; and 3) relatively or absolutely increased by structurally diverse growth inhibitors such as butyrate, salicylate and certain NSAIDs, and fatty acids. In general, then, mitochondrial fatty acid oxidation is inversely related to growth-associated changes in and ATP production, which are augmented in aerobic proliferation and diminshed in growth arrest or apoptosis. It is clear that the interaction between fatty acids and mitochondria — and its functional consequences — are varied, and may be regarded as “favorable” or “unfavorable” depending on their predominant characteristics and cellular context, e.g., growth arrest as opposed to mitosis. Such interactions are subject to modulation by cellular fatty acid abundance and compartmentalization, the latter influenced by extramitochondrial metabolism and cytosolic binding proteins. Within mitochondria, fatty acid may support ketogenesis and/or the citric acid cycle, thereby providing energy required, e.g., for hepatocyte gluconeogenesis or myocyte contraction. However, high rates of mitochondrial fatty acid caused by antiproliferative agents or fatty acid override of the CPT-I control point (the latter effect driven by elevated plasma [FFA] associated, e.g., with obesity, insulin resistance, diabetes mellitus, and HIV infection) may predispose to ROS generation and oxidative stress.

7.3.

ORIGINS OF INTRAMITOCHONDRIAL OXIDATIVE STRESS

At any given moment, entry of electrons into the electron transport chain principally reflects the rate at which they are derived from oxidation of metabolic fuels. As discussed above (see Fig. 7.1, and Chapter 6.4.: Salicylates and Other Nonsteroidal Anti-Inflammatory Drugs), intramitochondrial generation of ROS largely reflects “leakage” of some of these electrons from the electron transport chain, resulting in one-electron reduction of molecular oxygen to form superoxide anion and, through the action of superoxide dismutase, (ROS generation may also become prominent in states of high mitochondrial energization and as will be discussed below; see this Chapter 7.4.2.2.: Paradoxical Benefits of Adversity.) Consistent with this critical

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role of substrate oxidation-derived electron flux, caloric restriction decreases body fat stores and (fatty acid-related) intramitochondrial generation44, undoubtedly contributing to its anti-apoptotic, antitumorigenic, and longevity effects45-47. The major potentially oxidizable fuels (thus, the major electron donors) include fatty acids, glucose and its derivatives, and certain amino acids, especially those that may serve as substrates for gluconeogenesis (e.g., alanine), or whose derivatives directly enter the citric acid cycle (e.g., glutamine-derived 2-oxoglutarate). Each of these warrants consideration with regard to its propensity to generate excessive electron flux and thereby ROS and oxidative stress. Non-nutrient inducers of intramitochondrial ROS, such


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as ethanol and thermal stress, are discussed below (see this Chapter 7.4.2.2.: Paradoxical Benefits of Adversity.) Pyruvate, derived from glycolysis, lactate, glycerol, or gluconeogenic amino acids such as alanine, is a major electron donor. However, pyruvate and the glucose molecule from which it is usually derived are unlikely sources of electron transport chain-related oxidative stress because the rate at which they supply electrons is tightly controlled through feedback regulation of pyruvate dehydrogenase by the very conditions in the mitochondrial matrix that would otherwise predispose to ROS generation. These include increased electron abundance and reducing potential ([NADH]/[NAD]), high [acetyl CoA]/[CoA], and high [ATP]/[ADP]; the last of these three directly indicates relative deficiency of the ADP substrate that is required if electrons are to be removed efficiently through their consumption (effectively, neutralization) in ATP synthase-mediated oxidative phosphorylation. Hyperglycemia has been implicated as a cause of oxidative stressinduced tissue injury48,49. However, in vitro studies of this process often employ high glucose concentrations under conditions in which FFA may be absent or present at unphysiologically low concentrations. This is an important issue, as plasma [FFA] is almost invariably elevated in poorly controlled diabetes mellitus in vivo. In general, moreover, cellular uptake of glucose is opposed by the insulin resistant state to which these fatty acids contribute7,10, as well as by an autoregulatory effect of glucose itself50. Additionally, certain inconsistent findings, e.g., cell survival51 vs. cell death52, the complex and conflicting evidence concerning glucose and fatty acid interactions with the pancreatic cell53,54, and the absence of an effect of simvistatin on glycemic control despite its beneficial effect on vascular disease in diabetics55, suggest that high glucose concentrations need not be inherently injurious. Moreover, formation of advanced glycation endproducts (AGE) via alternative oxidative stress-dependent pathways has been demonstrated56,57, including their formation under euglycemic conditions56,58. Furthermore, streptozotocin diabetes mellitusinduced abnormalities in nerve conduction and were normalized by Finally, insulin, independent of glycemia and protein glycation59. macrovascular complications of type II diabetes mellitus often antedate or otherwise correlate poorly with hyperglycemia60,61, and may reflect Together, these enhancement of AGE toxicity by plasma FFA62. observations imply that factors other than (or in addition to) hyperglycemia, including fatty acids, play an important determining role in the pathogenesis of diabetes mellitus-associated tissue injury and outcome. As a corollary, it is possible that increases in plasma [FFA] and fatty acid oxidation are major causes of diabetes-related pro-oxidant

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effects, and/or that FFA and hyperglycemia are synergistic. Indeed, a basis for such synergy may be provided by the “Crabtree effect”, an incompletely understood glucose-induced inhibition of mitochondrial respiration63-66. Thus, consequent to an impairment of respiration and electron flux induced by the Crabtree effect, fatty acid-driven CPT-I 17-20,67 override and augmented would be expected to force even greater electron density in the electron transport chain, thereby promoting ROS generation, oxidative stress, and cell injury. Oxidation of other substrates such as glutamate or glutamine, which can enter the citric acid cycle independent of pyruvate dehydrogenase, may be limited, and therefore less likely to generate oxidative stress. For example, the liver is a net producer, not consumer, of glutamate 68 , and approximately 90% of hepatic glutamate uptake that does occur is localized to cells in the immediate pericentral region in support of glutamine synthesis69. Glutamine clearly can support citric acid cycle activity and oxidative phosphorylation in rapidly proliferating cells such as tumors and normal enterocytes70. However, glutamine is also committed to a diversity of metabolic functions 69,70 , not least being its established importance as the precursor of exported glutamate 68 and a substrate for gluconeogenesis71; significantly, requisite energy for the latter process is provided by mitochondrial fatty acid oxidation.

7.4.

CONSEQUENCES OF INTRAMITOCHONDRIAL OXIDATIVE STRESS: ATHEROSCLEROSIS AND BEYOND

The foregoing analysis suggests that in the intact organism, increased mitochondrial fatty acid may be a particularly important and perhaps dominant inducer of excessive, potentially ROS- and oxidative stress-generating, mitochondrial electron transport chain flux, in addition to the contribution of peroxisomal generation of This would apply especially to conditions associated with sustained increases in plasma and tissue fatty acids, such as obesity, insulin resistance, and diabetes mellitus. Plasma [FFA] is also increased by the adrenergic effects of smoking-related nicotine administration 72-75 . Moreover, it is of critical significance that FFA generated by lipolysis in the metabolically highly active and relatively insulin-resistant visceral adipose tissue76 are released into portal venous plasma. As a result, they may exert substantial influence on hepatocellular metabolism, yet remain relatively unrepresented in peripheral plasma FFA concentrations.


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These considerations suggest that the consequences of excessive fatty acid oxidation may be widely distributed, affecting directly or indirectly a diversity of proliferative and non-proliferative cells, and contributing importantly to the broader problem of oxidative stress and its sequelae, including atherosclerosis (see below), aging77-79, carcinogenesis80 (also, see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable), and certain forms of dementia and neurodegenerative disease (see Chapter 13.4.2.: HIV Dementia; and Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). They may also contribute to elucidation of the somewhat unexpected beneficial effects of caloric restriction44-47. Evidence in support of this general concept is provided by the following selected examples.

7.4.1. Atherosclerosis and Arterial Hypertension Current evidence indicates that development and progression of the endothelial lesion of atherosclerosis is promoted by preferential entry of oxidized low density lipoprotein (LDL) from plasma into the arterial endothelium and subendothelial compartment. In the vessel wall, oxidized LDL and other sources contribute to accumulation of lipid in endothelial macrophages, generating the “foam cell” and leading to the subsequent evolution of plaque formation, plaque rupture, and thrombosis. Two components of this complex process, i.e., formation of oxidized LDL and the endothelial lesion itself, are appropriately considered in the present context because of evidence suggesting that they may reflect at least in part pathological effects of fatty acid-related oxidative stress. 7.4.1.1. Oxidized LDL Several mechanisms have been identified by which oxidized LDL may be generated, but the origin of these particles in vivo remains unclear81-83. Because of this persisting uncertainty, a previously unexamined yet potential source of oxidized LDL may deserve consideration, namely, the hepatocellular endoplasmic reticulum (ER) in which the triacylglycerolrich VLDL precursors of LDL are assembled. There is strong evidence that the ER lumen is oxidizing relative to cytosol84-86, such that luminal GSH/GSSG ratio is approximately two This oxidizing orders of magnitude less than that of cytosol84. environment is important for proper folding of secretory proteins. It primarily reflects flavin adenine dinucleotide- and activity of the ER membrane protein Ero1p, and is subject to modulation by

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cytosolic conditions86. Molecular oxygen is the ultimate electron acceptor. As generation approximates 5% of total disulfide formation86, the redox process is similar to the electron transport chain in terms of proportionate ROS generation. In addition to oxidizing conditions generated within the ER lumen itself, ER resident proteins may also be subject to oxidative modification by low concentrations of exogenous (thus, cytosolic) as well87. Importantly, co-translational translocation of apolipoprotein B into the ER lumen may be delayed88, suggesting that it may exist at least transiently as a resident protein equivalent. As a result, nascent apolipoprotein B may be subject to modification by endogenous (intraluminal) or exogenous (cytosolic) oxidants. The concept that oxidizing conditions in the ER lumen, either normal or heightened by cellular or cytosolic oxidative stress, fosters oxidative changes in lipid or protein components of nascent triacylglycerol-rich VLDL is clearly hypothetical. However, such a scenario would be consistent with hepatocellular conditions in obesity and type II diabetes mellitus, and trans fatty acid metabolism; in these, entry of excessive fatty acid oxidation-generated mitochondrial or peroxisomal ROS into hepatocyte cytosol would contribute to the pathogenesis of non-alcoholic steatohepatitis and secondarily to ER oxidizing potential (see this Chapter 7.2.: Adverse Effects of Fatty Acids on Mitochondrial Function; and, 7.3.: Origins of Intramitochondrial Oxidative Stress). As partially oxidized dietary fatty acids have been shown to be incorporated into triacylglycerol-rich lipoproteins by both enterocyte and hepatocyte89,90, ER luminal oxidizing conditions could also induce and/or sustain a partially oxidized state of fatty acids destined for incorporation into VLDL lipids. Presumably, an effect on LDL longevity in the plasma compartment and endothelial uptake would require that the oxidized moiety (lipid or protein) be located at the surface of the LDL particle. Therefore, although oxidized TG- or cholesterol ester-fatty acids that are primarily associated with the lipid core may have relatively little effect on oxidized LDL clearance, they would contribute importantly to the burden of intracellular fatty acids that enter oxidative pathways after uptake by endothelial macrophages91. 7.4.1.2. The Endothelial Lesion In the evolving endothelial lesion of atherosclerosis92, and in experimental endothelial cholesterol loading93, increases in uptake of FFA and oxidized LDL are mediated at least in part by the CD36 scavenger receptor94. This process encumbers endothelial cells and endothelial


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macrophages with excess lipid (fatty acids as well as cholesterol); excess fatty acids drive increased mitochondrial and presumably — especially for trans fatty acids — peroxisomal fatty acid oxidation23-25. Fatty acids and their oxidative derivatives, released from oxidized LDL after uptake by endothelial macrophages, induce CD36 expression, thus activating a vicious cycle in which lipids in oxidized LDL augment their own uptake95,96. Oxidative fatty acid products here include not only hydroxy-derivatives97, but also those generated by Cellular abundance of fatty acids and therefore their availability to enter oxidative pathways in this setting is also increased by phospholipase A2 activation98. Moreover, among its diverse actions99, leptin exerts important and highly relevant effects on cell metabolism (see also Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior). These include: increased endothelial cell mitochondrial fatty acid oxidation, mediated via activation of AMPK100 or protein kinase A101; increased endothelial cell oxidative stress and INK activation102; and increased fatty acid oxidation in skeletal103,104 and cardiac muscle105. Plasma concentration of leptin, and therefore its potential activation of such oxidative processes in the endothelial cell, are increased in common forms of obesity. The increases in fatty acid oxidation-related ROS formation in mitochondria and MnSOD-generated and peroxisomes (direct formation of that are associated with these processes would:1) increase damage to mitochondrial DNA as an early event in atherogenesis106; 2) enhance monocyte/macrophage recruitment and expression of proinflammatory cytokines107-118 and C-reactive protein117-119; 3) provide on a sustained basis at least a portion of the required for the augmented myeloperoxidase activity that has been implicated in lesion pathogenesis57,120-122; 4) increase susceptibility to hyperglycemiaand AGE-associated endothelial cell injury 62 (also see this Chapter 7.3.: Origins of Intramitochondrial Oxidative Stress); and 5) predispose to the increase in macrophage apoptosis that is linked to plaque rupture123. In addition, unsaturated fatty acids increase degradation of the ATP-binding cassette transporter ABCA1, upon which macrophage cholesterol export depends124-126. The linkage of fatty acids to pathogenesis of the endothelial lesion is consistent with the anti-inflammatory effect of activation on endothelial cells127,128, and is further supported by evidence: 1) that fibrate activators, which inhibit CPT-I — and thereby fatty acid oxidation and generation of intramitochondrial oxidative stress129 — are anti-atherosclerotic109,130,131; 2) that deletion of the adipocyte FABP isoform ALBP/aP2 that is normally expressed in macrophages and is upregulated in foam cells132, is protective against atherosclerosis133-135; 3)

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that plasma levels of C-reactive protein reflect obesity and abdominal adipose tissue136, and are strongly associated with body mass index, insulin resistance, type II diabetes mellitus, and the “metabolic syndrome”119, and cigarette smoking137, while C-reactive protein itself exerts a direct proinflammatory effect on endothelial cells138; and 4) that the protective fatty acids diminish macrophage scavenger receptor expression139, thereby potentially suppressing uptake of cholesterol- and fatty acid-containing lipoproteins. In view of this substantial evidence, the mechanism underlying a recently reported anti-inflammatory response following lipoprotein lipase-induced triacylglycerol-rich lipoprotein hydrolysis140 is unclear. 7.4.1.3. Beneficial Effects of “Statin” Hypolipidemic Agents.

Against this background, it is particularly significant that the antiatherosclerotic effects of statins increasingly appear to transcend the decreases in serum cholesterol levels that result from inhibition of cholesterol biosynthesis at the level of the HMG CoA reductase reaction55. These broader benefits are primarily antiproliferative and antioxidant in nature. Thus, statin-induced growth arrest and/or apoptosis have been demonstrated in diverse cell types, including vascular endothelium141 and smooth muscle142, T lymphocytes143, and invasive breast cancer lines144. Equally broad antioxidant effects of statins are reflected in their inhibition of activation145,146 and of amyloid formation147 (also, see Chapter 15.5.2.: Hypothesized Role of Fatty Acid Metabolism in Neurodegeneration), and by diminished secretion of proinflammatory cytokines143,145 and C-reactive protein148. Although these surprisingly broad actions importantly affect the arterial wall, their similar influences in diverse cells and tissues are indicative of their more fundamental nature. Consideration of their potential mechanistic basis provides equally unexpected insights. Central to an understanding of the statin effects is the SREBP family of transcriptional regulators, which serves to coordinate expression of genes related to cellular sterol balance and fatty acid biosynthesis (see Chapter 4.4.: Fatty Acids, Modulation of Cell Growth, and the AMPActivated Protein Kinase). Statin inhibition of HMG CoA reductase blocks the biosynthesis of mevalonate, of its downstream intermediates including the key isoprenoids farnesyl pyrophosphate and geranylgeranyl pyrophosphate, and of sterols including cholesterol. The resulting cellular deficiency of cholesterol activates SREBP transcriptional regulation, but because SREBP-1c activation requires antecedent activation of LXR by the now less available oxysterols149, SREBP isoforms


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become independently regulated150 and SREBP-2 becomes the dominant activated species150,151. Both antiproliferative and antioxidant effects of HMG CoA reductase inhibition may be understandable in the context of these responses. Cell proliferation requires availability not only of fatty acids and sterols (especially cholesterol), but also of critical isoprenoid intermediates, the latter effecting, e.g., plasma membrane anchorage of RhoA/Ras/Raf/MAP kinase signaling pathway components141,144,152 (also, see Chapter 4.4.: Fatty Acids, Modulation of Cell Growth, and the AMPActivated Protein Kinase). Generation of these isoprenoids is blocked by statin inhibition of HMG CoA reductase. Although deficiency of cellular cholesterol may be mitigated by SREBP-2-induced increases in LDL receptor expression153-155, this mechanism does not ameliorate deficiency of the isoprenoid intermediates. The growth inhibitory effects of this deficiency, moreover, are augmented by SREBP-2 induced increases in 156 expression of the antiproliferative nuclear receptor . is also activated, possibly reflecting changes in cellular fatty acid economy146,152. Through an unknown mechanism, cell cycle inhibition by mevastatin reflects in part its suppression of phosphorylation-dependent activation of cdk2157. Antioxidant effects of statins may be understood at least in part as the result of SREBP-2-mediated transcriptional regulation of critical reactions in the fatty acid biosynthetic pathway. Generally this pathway is more specifically and vigorously regulated by SREBP-1c, but abundant evidence indicates that SREBP-2 also increases transcription and activity of the key mediators of fatty acid biosynthesis, i.e., acetyl CoA carboxylase and fatty acid synthase, at least in liver153,155, Hep G2 cells154, and intestine151. SREBP-2 mRNA abundance is also correlated with that of fatty acid synthase in breast cancer cells158. Thus, SREBP-induced activation of fatty acid biosynthesis, with its obligatory increase in malonyl CoA abundance and inhibition of CPT-I, both implies and effects suppression of mitochondrial fatty acid oxidation. As a corollary, fatty acid oxidation-induced generation of intramitochondrial ROS would also be suppressed, consistent with the observation that statins inhibit Creactive protein expression, unrelated to LDL cholesterol levels159. Statininduced inhibition of Rho GTPase as a result of geranyl-geranyl pyrophosphate deficiency also results in post-transcriptional upregulation of endothelial nitric oxide synthase160-162, likely contributing to improved vascular reactivity163. Antiproliferative and antioxidant effects of the statins may also interact144,145. The significance of vascular mitochondrial fatty acid oxidation-induced ROS generation in pathogenesis is further supported by the protective effects of the fibrate drugs, as noted above.

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Thus, while statins are hypothesized here to effect indirect suppression of mitochondrial CPT-I and fatty acid oxidation by increasing acetyl CoA carboxylase-mediated malonyl CoA generation, the fibrates accomplish this through direct inhibition of CPT-I itself. The isoprenoid-related antiproliferative effect is consistent with available evidence. In contrast, while the hypothesized, indirect, fatty acid biosynthesis-mediated antioxidant effect of SREBP-2 is highly plausible, it awaits experimental testing. If validated, it would not only help to clarify the mechanism underlying this increasingly important pharmacological effect of the statin hypolipidemic agents (see Chapter 15.5.: Metabolic Determinants in Alzheimer Disease: Synthesis), but in so doing would provide additional strong support for a broadly important connection between fatty acid oxidation and oxidative stress. Moreover, the hypothesized mechanism could account for the unexpected and incompletely understood inhibition of atherosclerosis in mice by a synthetic LXR ligand despite only minimal changes in plasma lipids164; in this instance LXR ligand-induced SREBP-1c activation would promote fatty acid synthesis-associated, malonyl CoA-mediated, suppression of mitochondrial fatty acid oxidation. Finally, atorvastatin was shown to inhibit the expression of NADPH oxidase components as a consequence of impaired prenylation165. In addition to their relationship to atherogenesis, fatty acid effects on vessel wall function may also contribute to the complex pathogenesis of arterial hypertension. Important linkages are suggested by the association of CD36 inactivation with both insulin resistance and arterial hypertension in the spontaneously hypertensive rat, in which elevated plasma [FFA] contribute to insulin resistance166,167. While less conclusively established, FFA effects on vascular tone and reactivity may be mediated in part through fatty acid oxidation-induced ROS generation with attenuation of nitric oxide synthase-dependent vasodilation, interaction with the angiotensin II receptor, and glucocorticoid effects5,165,168-170.

7.4.2. Adverse Effects of Fatty Acid Oxidation-Induced ROS in Various Organs Increases in plasma [FFA] — and thus in cellular fatty acid uptake and abundance — foster increased mitochondrial fatty acid oxidation and ROS generation. By this mechanism, they may contribute over time to the dysfunction of both cardiac myocytes21,105,171-176 and pancreatic islet beta-cells177-179 that are associated with obesity, insulin resistance, cardiac failure, and diabetes mellitus. This pathogenetic mechanism would also account for the protective antioxidant effect of agents that uncouple


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oxidative phosphorylation38-42, such as the protonophore carvedilol in the cardiac myocyte180 and UCP2 in the pancreatic beta cell181. Similar effects of fatty acids may mediate the altered islet cell metabolism and energetics that characterize type II diabetes mellitus, that are induced by 182 , and that are opposed by sulfonylurea-induced inhibition of CPT-I183. It also is consistent with the protection afforded by the wg/Wnt-like mediator of PI3K signaling Akt/PKB184, possibly including its recently recognized linkage to SREBP activation158,185 (also, see Chapter 4.4.: Fatty acids, Modulation of Cell Growth, and the AMP-Activated Protein Kinase). Fatty acid oxidationinduced mitochondrial oxidative stress may also contribute to the progressive type II diabetes mellitus-like syndrome that precedes hyperglycemia and diminished insulin secretory capacity associated with impaired frataxin activity in Friedreich’s ataxia and an animal model of the disease186. Although non-oxidative fatty acid metabolism has been implicated in “lipotoxicity”, i.e., lipid-associated cell injury187-189, lipotoxicity in recent cell culture studies was inversely correlated with fatty acid incorporation into triacylglycerols28; while this implies a direct relationship between lipotoxicity and fatty acid oxidation, the latter was not quantified in that report. Fatty acyl CoA thioesters may also perturb beta-cell function and insulin secretion non-oxidatively, through activation of the ATP-regulated channel190. In liver, oxidative stress generated at both mitochondrial and extramitochondrial (e.g., peroxisomal) sites as the result of excess hepatocellular abundance and oxidation of fatty acids is likely to contribute importantly to the pathogenesis of nonalcoholic steatohepatitis35,191-193. In addition, oxidative stress associated with excessive fatty acid oxidation may induce genomic instability — potentially self-perpetuating194 — promoting mutagenesis, and thereby contributing to the demonstrated association of hepatocellular carcinoma with diabetes mellitus80,195,196, and of cancer in general with obesity197. Tumorigenesis would be further augmented by In brain, nicotine-induced activation of PI3K/Akt/PKB signaling198. available evidence is consistent with the hypothesis that sustained increases in plasma FFA, acting directly and via augmented ketogenesis in brain astrocytes, may play an important role in the pathogenesis of Alzheimer disease (see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). 7.4.2.1. Ischemia-Reperfusion Injury and Heat Shock/Stress Response Fatty acid oxidation-induced intramitochondrial ROS generation may account to a large extent for the oxidative stress associated with ischemia-

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reperfusion injury199-202, and for the ROS-dependent mitochondrial channel activation that is associated with preconditioning-induced protection203. Fatty acid oxidation-dependent generation of ROS would also be consistent with the protection afforded both by inhibition of mitochondrial fatty acid oxidation204-207 and by the heat shock/stress response. The latter reflects in part a PI3K-mediated transcriptional activation of heat shock factor (HSF1)208 and the interaction between small heat shock proteins (Hsp) and intermediate filaments209-211. The latter linkage would be expected to modulate CPT-I activity, and thereby to suppress mitochondrial fatty acid oxidation212,213 (also, see Chapter 4.4.: Fatty acids, Modulation of Cell Growth, and the AMP-Activated Protein Kinase), consistent in the case of mild heat shock with cyclin D1 expression and cell cycle progression214. In addition, induction of Hsp expression215, together with that of UCP36,216, would serve to mitigate the adverse effects of ROS, likely reflecting in part stabilization of Akt/PKB by Hsp27217 and Hsp90218,219, and contributing importantly to the protective effect of preconditioning. As noted above (see Chapter 5.2.1.: further evidence of the importance of a mitochondrial origin of ROS in macrophages is provided in dramatic fashion by the inverse relationship between ROS and expression of UCP: deletion of UCP expression augments both ROS generation and killing of parasites220. 7.4.2.2. Paradoxical Benefits of Adversity In addtion to ischemic preconditioning, the foregoing concepts also provide an explanation for certain unexpected phenomena, such as the protection against ischemia/reperfusion injury that is afforded by brief antecedent heat shock as well as the longer term cardioprotective221 and neuroprotective222,223 benefits provided by regular intake of small amounts of ethanol. A plausible explanation for such effects may be that small quantities of ethanol224, heat shock, and ischemic preconditioning cause immediate, limited, and transient increases in mitochondrial electron transport chain electron abundance, thus predisposing to ROS generation225. With ethanol, the electron excess derives principally from alcohol dehydrogenase- and aldehyde dehydrogenase-generated NADH224, whereas in ischemic preconditioning insufficient oxygen is available to dispose of what otherwise would represent a relatively normal electron load. The pivotal role of ROS in preconditioning is strongly supported by the concentration-dependent protective effect of itself 226,227 . In the special case of heat shock, against ischemia/reperfusion injury increases in electron transport chain flux and ROS generation are to be expected as the result of the recognized exponential effect of temperature


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on biochemical reaction rates generally. Thus, assuming an approximate doubling of reaction rates for each 10°C increase in temperature – i.e., a of 2228 – a 5°C increase to 42°C, as is commonly employed in such experiments, could increase electron flux by approximately 70% relative to that at 37°C, with corresponding increases in ROS generation. Similar considerations may also account for the neuroprotective effects of 32°C hypothermia (a 5°C decrease), in which an anticipated 30% reduction in electron transport chain flux would be expected to correspondingly diminish ROS generation229. Because the otherwise adverse consequences of ischemic preconditioning, brief hyperthermia, and moderate ethanol ingestion are limited in both duration and intensity, they are not lethal. Instead, their dominant effect may be to elicit ROS-mediated induction of defensive survival responses as discussed above, that in limited ethanol use may be repeated and sustained. These consist of: 1) activation of PI3K- and survival signaling230,231 (also, see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable, and Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin), including stabilization of Akt/PKB by Hsp27217 and Hsp90218,219, and PI3K/Akt/PKB-activated glycolytic fueling of the citric acid cycle (e.g., via mitochondrial-bound hexokinase and inhibition of and 2) expression of uncoupling and heat shock proteins, resulting in dissipation of intramitochondrial ROS (via uncoupling proteins), and suppression of the mitochondrial permeability transition232 and of fatty acid oxidation-induced ROS (via heat shock protein- and intermediate filament-mediated constraints on CPT-I). A recently reported protective effect of low-dose against 231 ischemia/reperfusion injury may involve similar mechanisms. It is of additional interest that heat stress-induced Hsp70 expression attenuates changes that include decreased hepatocellular increased ROS generation, activation of and apoptosis233. These protective responses against effects provide additional evidence consistent with the concept that they depend at least in part on its augmentation of mitochondrial fatty acid oxidation234 (also, see Chapter 6.3.: Transforming Growth Importantly, the foregoing formulation, even if valid, would presumably apply only to those cells and tissues that are capable of mounting the requisite antioxidant defensive response, and to those circumstances in which increased ROS abundance is induced by an endogenous departure from physiological conditions (e.g., temporary thermal or ischemic stress) or by an exogenous agent that is locally and rapidly degradable (e.g., ethanol in moderate amounts). In contrast, for

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those circumstances in which cellular antioxidant defensive response is limited (e.g., brain neurons), or in which oxidative stress is imposed by a locally non-degradable xenobiotic which impairs mitochondrial function (e.g., many sedative/hypnotic drugs), repeated exposure even if brief may exert cumulative effects that are deleterious, rather than beneficial. The special case of neuron-astrocyte interaction in antioxidant defense is discussed later in this work (see Chapter 13.4.1.: Astrocyte-Neuron Interaction and Oxidative Stress; and, Chapter 15.5.: Metabolic Determinants in Alzheimer Disease: Synthesis). These and earlier considerations also serve to highlight the potentially contrasting effects of ROS on cell function and survival. Thus, ROS in low abundance may induce cell proliferation235,236, and are important activators of the survival-promoting transcription factors and AP1 237-239. In hypoxia, ROS effect concentration-dependent inhibition of protein tyrosine phosphatase 1B240,241, leading to activation of survival responses, e.g., wg/Wnt-like PI3K/Akt/PKB signaling, and Moreover, NO-derived peroxynitrite may activate activation242-244. glucose-6-phosphate dehydrogenase and thus the pentose phosphate pathway, thereby increasing formation of NADPH245, a critical cofactor in antiooxidant defense and in the biosynthesis of fatty acids, isoprenoids, and sterols (see Chapter 3.2.: Intermediary Metabolism: General Considerations). In contrast, generation of ROS in greater abundance and/or duration may induce apoptotic or necrotic cell death235,246-249 or, more gradually — over years or decades — to the diverse agingassociated complications discussed in the foregoing paragraphs (also, see Chapter 15.5.: Metabolic Determinants in Alzheimer Disease: Synthesis). Integrated effects of HSF-1 and Forkhead may mediate protective responses to these more sustained insults, e.g., fatty acid-associated mitochondrial dysfunction 250 , through their activation of heat shock proteins (HSF-1) and of antioxidant defenses and cell cycle arrest (Forkhead)251. Finally, it may seem paradoxical that at peak mitochondrial energization and its associated high and ATP abundance, electron transport chain “leakage” and ROS generation actually increase252-254. This well-documented phenomenon is predictable, given the constraints on electron transport chain flux imposed by the implicit decrease in ADP availability. Conceivably, it represents an evolutionarily ancient “signal”, communicated via the effects of low-level ROS on PI3K and wg/Wnt-like signaling discussed above. The signal would indicate that mitochondria are maximally energized, and therefore optimally prepared to support the enormous increases in energy generation and expenditure


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required by progression through the mitotic cycle (see Chapter 3.3.2.3.: Reactive Oxygen Species). As noted, long chain fatty acids may also uncouple mitochondrial oxidative phosphorylation through the protonophoric effect of the underivatized carboxylic acid30,32-34. Unlike short chain fatty acids (e.g., butyrate, ketone bodies) long chain fatty acids are not converted to acyl CoA thioesters in the mitochondrial matrix and therefore do not directly undergo As is true of other uncouplers, therefore, underivatized long chain fatty acids may diminish intramitochondrial ROS generation and oxidative stress. In so doing, however, they may compromise attainment of the increases in and thus in ATP synthesis, that are needed to support mitosis. At high rates of mitochondrial fatty acid oxidation, moreover, prominent pro-oxidant consequences of electron transport chain overload and electron “leakage” appear to override the relatively weak antioxidant uncoupling effect of the underivatized long chain fatty acids.

Figure 7.2. Growth modulator regulation of rodent hepatocyte fatty acid oxidation. Pathway arrow thickness approximates relative activity. Top: Inhibition of CPT-I, and thus mitochondrial fatty acid oxidation and ketogenesis, by the peroxisome proliferator clofibrate. Although cellular mitochondrial fatty acid oxidation increases absolutely, the fatty acid burden is diminished relative to an increased mitochondrial oxidative capacity; extramitochondrial FA oxidation is relatively enhanced. Bottom: Augmentation of mitochondrial fatty acid oxidation and ketogenesis by butyrate, with relative suppression of extramitochondrial FA oxidation. Abbreviations — CPT-I: carnitine palmitoyltransferase-I; FA: fatty acid; peroxisome proliferatoractivated

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Together, the concepts developed here support a model in which mitochondrial substrate oxidation and energization are central correlates and decisive determinants in the control of cell growth and cell injury. Mitochondrial generation of ROS, importantly influenced by fatty acid oxidation, may exert a broad spectrum of effects, depending on its intensity and duration. In growth (Fig. 7.2), the model holds that an inverse relationship exists between cell proliferation and mitochondrial fatty acid oxidation, absolute and/or relative to ATP biosynthesis. As a corollary, mitochondrial fatty acid oxidation is subject to modulation by agents which influence growth, including both small molecules (e.g., clofibrate, butyrate, NSAIDs, fatty acids) and macromolecules (e.g., insulin, and The model provides for the metabolic versatility and responsiveness implicitly required for precise control of growth-related processes under rapidly changing conditions. Its operation is demonstrated perhaps most impressively by the consequences of 70% hepatectomy. In this setting, an abruptly imposed metabolic crisis elicits a response in which previously quiescent hepatocytes execute a rapid and timely entry into, and subsequent exit from, the mitotic cycle.

7.5.

REFERENCES

1. X. Chen, N. Iqbal, and G. Boden, The effects of free fatty acids on gluconeogenesis and glycogenolysis in normal subjects., J Clin Invest 103:365-372 (1999). 2. A. Dresner, D. Laurent, M. Marcucci, et al., Effects of free fatty acids on glucose transport and IRS-1-associated phosphatidylinositol 3-kinase activity., J Clin Invest 103:253-259 (1999). 3. M. E. Griffin, M. J. Marcucci, G. W. Cline, et al., Free fatty acid-induced insulin resistance is associated with activation of protein kinase C theta and alterations in the insulin signaling cascade, Diabetes 48:1270-4 (1999). 4. K. Cusi, K. Maezono, A. Osman, et al., Insulin resistance differentially affects the PI 3kinase- and MAP kinase-mediated signaling in human muscle., J Clin Invest 105:311-320 (2000). 5. H. Steinberg, G. Paradisi, G. Hook, K. Crowder, J. Cronin, and A. Baron, Free fatty acid elevation impairs insulin-mediated vasodilation and nitric oxide production, Diabetes 49:1231-1238 (2000). 6. P. Arner, Insulin resistance in type 2 diabetes: role of fatty acids, Diabetes Metab Res Rev 18 Suppl 2:S5-9 (2002). 7. G. Boden, and G. I. Shulman, Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction, Eur J Clin Invest 32 Suppl 3:14-23 (2002). 8. Y. Kruszynska, D. Worrall, J. Ofrecio, J. Frias, G. Macaraeg, and J. Olefsky, Fatty acid-induced insulin resistance: decreased muscle PI3K activation but unchanged Akt phosphorylation, J Clin Endocrinol Metab 87:226-234 (2002).


161

9. T. K. Lam, H. Yoshii, C. A. Haber, et al., Free fatty acid-induced hepatic insulin resistance: a potential role for protein kinase C-delta, Am J Physiol Endocrinol Metab 283:E682-91 (2002). 10. J. McGarry, Banting lecture, 2001: dysregulation of fatty acid metabolism in the etiology of type 2 diabetes, Diabetes 51:7-18 (2002). 11. C. L. Soltys, L. Buchholz, M. Gandhi, A. S. Clanachan, K. Walsh, and J. R. Dyck, Phosphorylation of cardiac protein kinase B is regulated by palmitate, Am J Physiol Heart Circ Physiol 283:H1056-64 (2002). 12. C. Yu, Y. Chen, G. W. Cline, et al., Mechanism by which fatty acids inhibit insulin activation of insulin receptor substrate-1 (IRS-1)-associated phosphatidylinositol 3kinase activity in muscle, J Biol Chem 277:50230-50236 (2002). 13. Z. Jiang, Y. Lin, A. Clemont, et al., Characterization of selective resistance to insulin signaling in the vasculature of obese Zucker (fa/fa) rats, J Clin Invest 104:447-457 (1999). 14. R. Behal, D. Buxton, J. Robertson, and M. Olson, Regulation of the pyruvate dehydrogenase multienzyme complex., Annu Rev Nutr 13:497-520 (1993). 15. R. Scholz, M. Olson, A. Schwab, U. Schwabe, C. Noell, and W. Braun, The effect of fatty acids on the regulation of pyruvate dehydrogenase in perfused rat liver, Eur J Biochem 86:519-530 (1978). 16. T. A. Hopkins, M. C. Sugden, M. J. Holness, R. Kozak, J. R. Dyck, and G. D. Lopaschuk, Control of cardiac pyruvate dehydrogenase activity in peroxisome proliferator-activated receptor-alpha transgenic mice, Am J Physiol Heart Circ Physiol 285:H270-6 (2003). 17. S. Mills, D. Foster, and J. Mcgarry, Interaction of malonyl-CoA and related compounds with mitochondria from different rat tissues: Relationship between ligand binding and inhibition of carnitine palmitoyltransferase I, Biochem J 214:83-91 (1983). 18. L. Drynan, P. Quant, and V. Zammit, Flux control exerted by mitochondrial outer membrane carnitine palmitoyltransferase over beta-oxidation, ketogenesis and tricarboxylic acid cycle activity in hepatocytes isolated from rats in different metabolic states., Biochem J 317:791-795 (1996A). 19. L. Drynan, P. Quant, and V. Zammit, The role of changes in the sensitivity of hepatic mitochondrial overt carnitine palmitoyltransferase in determining the onset of the ketosis of starvation in the rat., Biochem J 318:767-770 (1996B). 20. J. Sleboda, K. Risan, O. Spydevold, and J. Bremer, Short-term regulation of carnitine palmitoyltransferase I in cultured rat hepatocytes: spontaneous inactivation and reactivation by fatty acids, Biochim Biophys Acta 1436:541-549 (1999). 21. M. Young, G. Goodwin, J. Ying, et al., Regulation of cardiac and skeletal muscle malonyl-CoA decarboxylase by fatty acids, Am J Physiol Endocrinol Metab 280: E471-E479 (2001 A). 22. M. Mutomba, H, Yan, M. Konyavko, et al., Regulation of the activity of caspases by L-carnitine and palmitoylcarnitine, FEBS Letters 478:19-25 (2000). 23. L. D. Lawson, and F. A. Kummerow, beta-Oxidation of the coenzyme A esters of elaidic, oleic, and stearic acids and their full-cycle intermediates by rat heart mitochondria, Biochim Biophys Acta 573:245-54 (1979). 24. T. Flatmark, A. Nilsson, J. Kvannes, et al., On the mechanism of induction of the enzyme systems for peroxisomal beta-oxidation of fatty acids in rat liver by diets rich in partially hydrogenated fish oil, Biochim Biophys Acta 962:122-30 (1988).

162

Chapter 7

25. A. Ascherio, M. B. Katan, P. L. Zock, M. J. Stampfer, and W. C. Willett, Trans fatty acids and coronary heart disease, N Engl J Med 340:1994-8 (1999). 26. G. Klug, J. Krause, A. Ostlund, O. Knoll, and D. Brdiczka, Alterations in liver mitochondrial function as a result of fasting and exhaustive exercise, Biochim Biophys Acta 764:272-282 (1984). 27. G. Beutner, A. Rück, B. Riede, and D. Brdiczka, Complexes between porin, hexokinase, mitochondrial creatine kinase and adenylate translocates display properties of the permeability transition pore: Implication for regulation of permeability transition by the kinases., Biochim Biophys Acta 1368:7-18 (1998). 28. L. L. Listenberger, X. Han, S. E. Lewis, et al., Triglyceride accumulation protects against fatty acid-induced lipotoxicity, Proc Natl Acad Sci U S A 100:3077-82 (2003). 29. M. Paumen, Y. Ishida, M. Muramatsu, M. Yamamoto, and T. Honjo, Inhibition of carnitine palmitoyltransferase I augments sphingolipid synthesis and palmitateinduced apoptosis, J Biol Chem 272:3324-3329 (1997B). 30. V. Skulachev, Fatty acid circuit as a physiological mechanism of uncoupling of oxidative phosphorylation, FEBS Lett 294:158-162 (1991). 31. L. Wojtczak, and P. Schönfeld, Effect of fatty acids on energy coupling processes in mitochondria, Biochim Biophys Acta 1183:41-57 (1993). 32. L. Wojtczak, and M. Wiêckowski, The mechanisms of fatty acid-induced proton permeability of the inner mitochondrial membrane, J Bioenerg Biomembranes 31:447-455 (1999A). 33. P. Schönfeld, M. Wiêckowski, and L. Wojtczak, Thyroid hormone-induced expression of the ADP/ATP carrier and its effect on fatty acid-induced uncoupling of oxidative phosphorylation, FEBS Lett 416:19-22 (1997A). 34. P. Schönfeld, and R. Bohnensack, Fatty acid-promoted mitochondrial permeability transition by membrane depolarization and binding to the ADP/ATP carrier, FEBS Lett 420:167-170 (1997B). 35. K. Chavin, S. Yang, H. Lin, et al., Obesity induces expression of uncoupling protein2 in hepatocytes and promotes liver ATP depletion., J Biol Chem 274:5692-5700 (1999). 36. K. Echtay, D. Roussel, J. St-Pierre, et al., Superoxide activates mitochondrial uncoupling proteins., Nature 415:96-99 (2002). 37. D. Rolfe, A. Hulbert, and M. Brand, Characteristics of mitochondrial proton leak and control of oxidative phosphorylation in the major oxygen-consuming tissues of the rat, Biochim Biophys Acta 1188:405-416 (1994). 38. I. Reynolds, and T. Hastings, Glutamate induces the production of reactive oxygen species in cultured forebrain neurons following NMDA receptor activation, J Neurosci 15:3318-3327 (1995). 39. V. Skulachev, Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants, Quart Rev Biophys 29:169-202 (1996). 40. A. Nègre-Salvayre, C. Hirtz, G. Carrera, et al., A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation, FASEB J 11:809-815 (1997). 41. A. Stout, H. Raphael, B. Kanterewicz, E. Klann, and I. Reynolds, Glutamate-induced neuron death requires mitochondrial calcium uptake, Nature Neurosci 1:366-373 (1998).


163

42. S. Korshunov, O. Kokina, E. Ruuge, V. Skulachev, and A. Starkov, Fatty acids as natural uncouplers preventing generation of and by mitochondria in the resting state, FEBS Letts 435:215-218 (1998). 43. S. Tanaka, L. Mohr, E. Schmidt, K. Sugimachi, and J. Wands, Biological effects of human insulin receptor substrate-1 overexpression in hepatocytes, Hepatol 26:598604 (1997). 44. R. Gredilla, G. Barja, and M. Lopez-Torres, Effect of short-term caloric restriction on production and oxidative DNA damage in rat liver mitochondria and location of the free radical source, J Bioenerg Biomembr 33:279-287 (2001). 45. B. Grasl-Kraupp, W. Bursch, B. Ruttkay-Nedecky, A. Wagner, B. Lauer, and R. Schulte-Hermann, Food restriction eliminates preneoplastic cells through apoptosis and antagonizes carcinogenesis in rat liver, Proc Natl Acad Sci USA 91:9995-9999 (1994). 46. K. Kolaja, K. Bunting, and J. Klaunig, Inhibition of tumor promotion and hepatocellular growth by dietary restriction in mice, Carcinogenesis 17:1657-1664 (1996). 47. V. D. Longo, and C. E. Finch, Evolutionary medicine: from dwarf model systems to healthy centenarians?, Science 299:1342-6 (2003). 48. X. Du, D. Edelstein, L. Rossetti, et al., Hyperglycemia-induced mitochondrial superoxide overproduction activates the hexosamine pathway and induces plasminogen activator inhibitor-1 expression by increasing Sp1 glycosylation., Proc Natl Acad Sci USA 97:12222-12226 (2000). 49. T. Nishikawa, D. Edelstein, X. Du, et al., Normalizing mitochondrial superoxide production blocks three pathways of hyperglycaemic damage, Nature 404:787-790 (2000). 50. S. I. Itani, A. K. Saha, T. G. Kurowski, H. R. Coffin, K. Tornheim, and N. B. Ruderman, Glucose autoregulates its uptake in skeletal muscle: involvement of AMPactivated protein kinase, Diabetes 52:1635-40 (2003). 51. S. Srinivasan, E. Bernal-Mizrachi, M. Ohsugi, and M. A. Permutt, Glucose promotes pancreatic islet beta-cell survival through a PI 3-kinase/Akt-signaling pathway, Am J Physiol Endocrinol Metab 283:E784-93 (2002). 52. K. Maedler, P. Sergeev, F. Ris, et al., Glucose-induced beta cell production of IL-lbeta contributes to glucotoxicity in human pancreatic islets, J Clin Invest 110:851-60 (2002). 53. D. T. Stein, B. E. Stevenson, M. W. Chester, et al., The insulinotropic potency of fatty acids is influenced profoundly by their chain length and degree of saturation, J Clin Invest 100:398-403 (1997). 54. K. Maedler, J. Oberholzer, P. Bucher, G. A. Spinas, and M. Y. Donath, Monounsaturated Fatty Acids Prevent the Deleterious Effects of Palmitate and High Glucose on Human Pancreatic beta-Cell Turnover and Function, Diabetes 52:726-33 (2003). 55. R. Collins, J. Armitage, S. Parish, P. Sleigh, and R. Peto, MRC/BHF Heart Protection Study of cholesterol-lowering with simvastatin in 5963 people with diabetes: a randomised placebo-controlled trial, Lancet 361:2005-16 (2003). 56. W. Palinski, T. Koschinsky, S. W. Butler, et al., Immunological evidence for the presence of advanced glycosylation end products in atherosclerotic lesions of euglycemic rabbits, Arterioscler Thromb Vase Biol 15:571-82 (1995). 57. M. Anderson, J. Requena, J. Crowley, S. Thorpe, and J. Heinecke, The myeloperoxidase system of human phagocytes generates N-epsilon-

164

Chapter 7

(carboxymethyl)lysine on proteins: a mechanism for producing advanced glycation end products at sites of inflammation., J Clin Invest 104:103-113 (1999). 58. T. Sakaguchi, S. F. Van, S. D. Yan, et al., Central role of RAGE-dependent neointimal expansion in arterial restenosis, J Clin Invest 111:959-72 (2003). 59. T. J. Huang, S. A. Price, L. Chilton, et al., Insulin prevents depolarization of the mitochondrial inner membrane in sensory neurons of type 1 diabetic rats in the presence of sustained hyperglycemia, Diabetes 52:2129-36 (2003). 60. W. C. Duckworth, M. McCarren, and C. Abraira, Glucose control and cardiovascular complications: the VA Diabetes Trial, Diabetes Care 24:942-5 (2001 A). 61. W. C. Duckworth, Hyperglycemia and cardiovascular disease, Curr Atheroscler Rep 3:383-91 (2001B). 62. S. Yamagishi, T. Okamoto, S. Amano, et al., Palmitate-induced apoptosis of microvascular endothelial cells and pericytes, Mol Med 8:179-84 (2002). 63. I. Sussman, M. Erecinska, and D. F. Wilson, Regulation of cellular energy metabolism: the Crabtree effect, Biochim Biophys Acta 591:209-23 (1980). 64. L. Wojtczak, V. Teplova, K. Bogucka, et al., Effect of glucose and deoxyglucose on the redistribution of calcium in ehrlich ascites tumour and Zajdela hepatoma cells and its consequences for mitochondrial energetics: Further arguments for the role of Ca(2+) in the mechanism of the crabtree effect, Eur J Biochem 263:495-501 (1999B). 65. S. Rodriguez-Enriquez, O. Juarez, J. Rodriguez-Zavala, and R. Moreno-Sanchez, Multisite control of the Crabtree effect in ascites hepatoma cells, Eur J Biochem 268:2512-2519 (2001). 66. V. Lemeshko, Model of the outer membrane potential generation by the inner membrane of mitochondria, Biophys J 82:684-692 (2002). 67. J. McGarry, and N. Brown, Reconstitution of purified, active and malonyl-CoAsensitive rat liver carnitine palmitoyltransferase I: relationship between membrane environment and malonyl-CoA sensitivity, Biochem J 349:179-187 (2000). 68. M. Hediger, and T. Welbourne, Glutamate transport and metabolism Introduction: Glutamate transport, metabolism, and physiological responses, Am J Physiol 277:F477-F480 (1999). 69. D. Häussinger, Hepatic glutamine transport and metabolism, Adv Enzymol Rel Areas Molec Biol 72:43-86 (1998). 70. Z. Kovacevic, and J. Mcgivan, Mitochondrial metabolism of glutamine and glutamate and its physiological significance, Physiol Rev 63:547-605 (1983). 71. R. Hankard, M. Haymond, and D. Darmaun, Role of glutamine as a glucose precursor in fasting humans, Diabetes 46:1535-1541 (1997). 72. P. Cryer, M. Haymond, J. Santiago, and S. Shah, Norepinephrine and epinephrine release and adrenergic mediation of smoking-associated hemodynamic and metabolic events., New Engl J Med 295:573-7 (1976). 73. M. Hellerstein, N. Benowitz, R. Neese, et al., Effects of cigarette smoking and its cessation on lipid metabolism and energy expenditure in heavy smokers, J Clin Invest 93:265-72 (1994). 74. K. Fattinger, D. Verotta, and N. Benowitz, Pharmacodynamics of acute tolerance to multiple nicotinic effects in humans., J Pharmacol Exp Therapeutics 281:1238-46 (1997). 75. J. Rincón, A. Krook, D. Galuska, H. Wallberg-Henriksson, and J. Zierath, Altered skeletal muscle glucose transport and blood lipid levels in habitual cigarette smokers, Clin Physiol 19:135-142 (1999).


165

76. S. Mittelman, G. Van Citters, E. Kirkman, and R. Bergman, Extreme insulin resistance of the central adipose depot in vivo, Diabetes 51:755-761 (2002). 77. T. Finkel, and N. Holbrook, Oxidants, oxidative stress and the biology of ageing., Nature 408:239-247 (2000B). 78. J. Liu, D. W. Killilea, and B. N. Ames, Age-associated mitochondrial oxidative decay: improvement of carnitine acetyltransferase substrate-binding affinity and activity in brain by feeding old rats acetyl-L- carnitine and/or R-alpha -lipoic acid, Proc Natl Acad Sci U S A 99:1876-81 (2002). 79. S. Hekimi, and L. Guarente, Genetics and the specificity of the aging process, Science 299:1351-4 (2003). 80. R. Ockner, R. Kaikaus, and N. Bass, Fatty acid metabolism and the pathogenesis of hepatocellular carcinoma: review and hypothesis, Hepatology 18:669-676 (1993). 81. G. M. Chisolm, and D. Steinberg, The oxidative modification hypothesis of atherogenesis: an overview, Free Radic Biol Med 28:1815-26 (2000). 82. J. P. Gaut, and J. W. Heinecke, Mechanisms for oxidizing low-density lipoprotein. Insights from patterns of oxidation products in the artery wall and from mouse models of atherosclerosis, Trends Cardiovasc Med 11:103-12 (2001). 83. J. L. Witztum, and D. Steinberg, The oxidative modification hypothesis of atherosclerosis: does it hold for humans?, Trends Cardiovasc Med 11:93-102 (2001). 84. C. Hwang, A. J. Sinskey, and H. F. Lodish, Oxidized redox state of glutathione in the endoplasmic reticulum, Science 257:1496-502 (1992). 85. A. R. Frand, J. W. Cuozzo, and C. A. Kaiser, Pathways for protein disulphide bond formation, Trends Cell Biol 10:203-10 (2000). 86. B. P. Tu, and J. S. Weissman, The FAD- and O(2)-Dependent Reaction Cycle of ErolMediated Oxidative Protein Folding in the Endoplasmic Reticulum, Mol Cell 10:98394 (2002). 87. D. van der Vlies, E. H. Pap, J. A. Post, J. E. Celis, and K. W. Wirtz, Endoplasmic reticulum resident proteins of normal human dermal fibroblasts are the major targets for oxidative stress induced by hydrogen peroxide, Biochem J 366:825-30 (2002). 88. R. A. Davis, and T. Y. Hui, 2000 George Lyman Duff Memorial Lecture: atherosclerosis is a liver disease of the heart, Arterioscler Thromb Vasc Biol 21:88798 (2001). 89.I. Staprans, J. H. Rapp, X. M. Pan, and K. R. Feingold, Oxidized lipids in the diet are incorporated by the liver into very low density lipoprotein in rats, J Lipid Res 37:420-30 (1996). 90. M. Penumetcha, N. Khan, and S. Parthasarathy, Dietary oxidized fatty acids: an atherogenic risk?, J Lipid Res 41:1473-80 (2000). 91. R. Salvayre, N. Auge, H. Benoist, and A. Negre-Salvayre, Oxidized low-density lipoprotein-induced apoptosis, Biochim Biophys Acta 1585:213-21 (2002). 92. I. Tabas, Consequences of cellular cholesterol accumulation: basic concepts and physiological implications, J Clin Invest 110:905-11 (2002). 93. P. Yao, and I. Tabas, Free cholesterol loading of macrophages is associated with widespread mitochondrial dysfunction and activation of the mitochondrial apoptosis pathway, J Biol Chem 276:42468-42476 (2001). 94. M. Febbraio, D. Hajjar, and R. Silverstein, CD36: a class B scavenger receptor involved in angiogenesis, atherosclerosis, inflammation, and lipid metabolism., J Clin Invest 108:785-791 (2001).

166

Chapter 7

95. L. Nagy, P. Tontonoz, J. G. Alvarez, H. Chen, and R. M. Evans, Oxidized LDL regulates macrophage gene expression through ligand activation of PPARgamma, Cell 93:229-40 (1998). 96. P. Tontonoz, L. Nagy, J. G. Alvarez, V. A. Thomazy, and R. M. Evans, PPARgamma promotes monocyte/macrophage differentiation and uptake of oxidized LDL, Cell 93:241-52 (1998). 97. E. I. Waddington, K. D. Croft, K. Sienuarine, B. Latham, and I. B. Puddey, Fatty acid oxidation products in human atherosclerotic plaque: an analysis of clinical and histopathological correlates, Atherosclerosis 167:111-20 (2003). 98. S. Panini, L. Yang, A. Rusinol, M. Sinensky, J. Bonventre, and C. Leslie, Arachidonate metabolism and the signaling pathway of induction of apoptosis by oxidized LDL/oxysterol, J Lipid Res 42:1678-1686 (2001). 99. A. Z. Zhao, M. M. Shinohara, D. Huang, et al., Leptin induces insulin-like signaling that antagonizes cAMP elevation by glucagon in hepatocytes, J Biol Chem 275:11348-54 (2000). 100. Z. Dagher, N. Ruderman, K. Tornheim, and Y. Ido, Acute regulation of fatty acid oxidation and AMP-activated protein kinase in human umbilical vein endothelial cells., Circ Res 88:1276-82 (2001). 101. S. Yamagishi, D. Edelstein, X. Du, Y. Kaneda, M. Guzman, and M. Brownlee, Leptin induces mitochondrial superoxide production and monocyte chemoattractant protein-1 expression in aortic endothelial cells by increasing fatty acid oxidation via protein kinase A, J Biol Chem 276:25096-25100 (2001). 102. A. Bouloumie, T. Marumo, M. Lafontan, and R. Busse, Leptin induces oxidative stress in human endothelial cells, Faseb J 13:1231-8 (1999). 103. G. Steinberg, A. Bonen, and D. Dyck, Fatty acid oxidation and triacylglycerol hydrolysis are enhanced after chronic leptin treatment in rats, AJP - Endocrinol Metab 282:E593-E600 (2002). 104. Y. Minokoshi, Y. Kim, O. Peroni, et al., Leptin stimulates fatty-acid oxidation by activating AMP-activated protein kinase, Nature 415:339-343 (2002). 105. L. L. Atkinson, M. A. Fischer, and G. D. Lopaschuk, Leptin activates cardiac fatty acid oxidation independent of changes in the AMP-activated protein kinase-acetylCoA carboxylase-malonyl-CoA axis, J Biol Chem 277:29424-30 (2002). 106. S. W. Ballinger, C. Patterson, C. A. Knight-Lozano, et al., Mitochondrial integrity and function in atherogenesis, Circulation 106:544-9 (2002). 107. G. Chinetti, S. Griglio, M. Antonucci, et al., Activation of proliferator-activated receptors alpha and gamma induces apoptosis of human monocyte-derived macropages., J Biol Chem 273:25573-25580 (1998). 108. P. Delerive, B, De, K, S. Besnard, et al., Peroxisome proliferated-activated receptor alpha negatively regulates the vascular inflammatory gene response by negative cross-talk with transcription factors NF-kappaB and AP-1., J Biol Chem 274:3204832054 (1999). 109. N. Marx, G. Sukhova, T. Collins, P. Libby, and J. Plutzky, PPARalpha activators inhibit cytokine-induced vascular cell adhesion molecule-1 expression in human endothelial cells, Circulation 99:3125-3131 (1999). 110. I. Rusyn, C. Bradham, L. Cohn, et al., Corn oil rapidly activates nuclear factorkappaB in hepatic Kupffer cells by oxidant-dependent mechanisms, Carcinogenesis 20:2095-2100 (1999). 111. R. De Caterina, J. Liao, and P. Libby, Fatty acid modulation of endothelial activation., Am J Clin Nutr 71:213S-23S (2000).


167

112. P. Delerive, C. Furman, E. Teissier, J.-C. Fruchart, P. Duriez, and B. Staels, Oxidized phosopholipids activate PPARa in a phospholipase A2-dependent manner., FEBS Letters 471:34-38 (2000A). 113. B. Hennig, P. Meerarani, P. Ramadass, B. Watkins, and M. Toborek, Fatty acidmediated activation of vascular endothelial cells, Metab Clin Exper 49:1006-1013 (2000). 114. B. Neve, J.-C. Fruchart, and B. Staels, Role of the peroxisome proliferator-activated receptors (PPAR) in atherosclerosis, Biochem Pharmacol 60:1245-1250 (2000). 115. C. Glass, Potential roles of the peroxisome proliferator-activated receptor-gamma in macrophage biology and atherosclerosis, J Endocrinol 169:461-464 (2001). 116. W. Dichtl, M. Ares, A. Jonson, et al., Linoleic Linoleic acid-stimulated vascular adhesion molecule-1 expression in endothelial cells depends on nuclear factor-kappaB activation., Metabolism 51:327-333 (2002). 117. P. Libby, P. Ridker, and A. Maseri, Inflammation and atherosclerosis, Circulation 105:1135-1143 (2002). 118. A. D. Pradhan, J. E. Manson, J. E. Rossouw, et al., Inflammatory biomarkers, hormone replacement therapy, and incident coronary heart disease: prospective analysis from the Women’s Health Initiative Observational Study, Jama 288:980-7 (2002). 119. M. B. Pepys, and G. M. Hirschfield, C-reactive protein: a critical update, J Clin Invest 111:1805-12 (2003). 120. S. Klebanoff, Myeloperoxidase, Proceedings of the Association of American Physicians 111:383-389 (1999). 121. E. Podrez, H. Abu-Soud, and S. Hazen, Myeloperoxidase-generated oxidants and atherosclerosis, Free Radical Biol Med 28:1717-1725 (2000). 122. W. Li, F. Miller, Jr, H. Zhang, D. Spitz, L. Oberley, and N. Weintraub, production by a non-phagocytic NAD(P)H oxidase causes oxidant injury, J Biol Chem 276:29251-29256 (2001). 123. Z. Mallat, and A. Tedgui, Current perspective on the role of apoptosis in atherothrombotic disease, Circulation Res 88:998-1003 (2001). 124. R. R. Singaraja, C. Fievet, G. Castro, et al., Increased ABCA1 activity protects against atherosclerosis, J Clin Invest 110:35-42 (2002). 125. Y. Wang, and J. Oram, Unsaturated fatty acids inhibit cholesterol efflux from macrophages by increasing degradation of ATP-binding cassette transporter A1, J Biol Chem 277:5692-5697 (2002). 126. A. R. Tall, P. Costet, and N. Wang, Regulation and mechanisms of macrophage cholesterol efflux, J Clin Invest 110:899-904 (2002). 127. N. Wang, L. Verna, N. G. Chen, et al., Constitutive activation of peroxisome proliferator-activated receptor-gamma suppresses pro-inflammatory adhesion molecules in human vascular endothelial cells, J Biol Chem 277:34176-81 (2002). 128. C.-H. Lee, and R. Evans, Peroxisome proliferator-activated receptor-gamma in macrophage lipid homeostasis, Trends Endocrinol Metab 13:331-335 (2002). 129. M. Poynter, and R. Daynes, Peroxisome proliferator-activated receptor alpha activation modulates cellular redox status, represses nuclear factor-kappaB signaling, and reduces inflammatory cytokine production in aging, J Biol Chem 273:3283332841 (1998). 130. H. Duez, Y. S. Chao, M. Hernandez, et al., Reduction of atherosclerosis by the peroxisome proliferator-activated receptor alpha agonist fenofibrate in mice, J Biol Chem 277:48051-7 (2002).

168

Chapter 7

131. G. A. Francis, J. S. Annicotte, and J. Auwerx, PPAR-alpha effects on the heart and other vascular tissues, Am J Physiol Heart Circ Physiol 285:H1-9 (2003). 132. Y. Fu, N. Luo, M. F. Lopes-Virella, and W. T. Garvey, The adipocyte lipid binding protein (ALBP/aP2) gene facilitates foam cell formation in human THP-1 macrophages, Atherosclerosis 165:259-69 (2002). 133. L. Makowski, J. Boord, K. Maeda, et al., Lack of macrophage fatty-acid-binding protein aP2 protects mice deficient in apolipoprotein E against atherosclerosis, Nature Med 7:699-705 (2001). 134. M. Perrella, A. Pellacani, M. Layne, et al., Absence of adipocyte fatty acid binding protein prevents the development of accelerated atherosclerosis in hypercholesterolemic mice, FASEB J 15:1774-1776 (2001). 135. J. B. Boord, K. Maeda, L. Makowski, et al., Adipocyte fatty acid-binding protein, aP2, alters late atherosclerotic lesion formation in severe hypercholesterolemia, Arterioscler Thromb Vasc Biol 22:1686-91 (2002). 136. I. Lemieux, A. Pascot, D. Prud’homme, et al., Elevated C-reactive protein: another component of the atherothrombotic profile of abdominal obesity, Arterioscler Thromb Vasc Biol 21:961-7 (2001). 137. L. A. Bazzano, J. He, P. Muntner, S. Vupputuri, and P. K. Whelton, Relationship between cigarette smoking and novel risk factors for cardiovascular disease in the United States, Ann Intern Med 138:891-7 (2003). 138. V. Pasceri, J. T. Willerson, and E. T. Yeh, Direct proinflammatory effect of Creactive protein on human endothelial cells, Circulation 102:2165-8 (2000). 139. E. Miles, F. Wallace, and P. Calder, Dietary fish oil reduces intercellular adhesion molecule 1 and scavenger receptor expression on murine macrophages, Athersclerosis 152:43-50 (2000). 140. O. Ziouzenkova, S. Perrey, L. Asatryan, et al., Lipolysis of triglyceride-rich lipoproteins generates PPAR ligands: evidence for an antiinflammatory role for lipoprotein lipase, Proc Natl Acad Sci U S A 100:2730-35 (2003). 141. L. Vincent, C. Soria, F. Mirshahi, et al., Cerivastatin, an inhibitor of 3-hydroxy-3methylglutaryl coenzyme a reductase, inhibits endothelial cell proliferation induced by angiogenic factors in vitro and angiogenesis in in vivo models, Arterioscler Thromb Vasc Biol 22:623-9 (2002). 142. C. B. Xu, E. Stenman, and L. Edvinsson, Reduction of bFGF-induced smooth muscle cell proliferation and endothelin receptor mRNA expression by mevastatin and atorvastatin, Biochem Pharmacol 64:497-505 (2002). 143. S. Youssef, O. Stuve, J. C. Patarroyo, et al., The HMG-CoA reductase inhibitor, atorvastatin, promotes a Th2 bias and reverses paralysis in central nervous system autoimmune disease, Nature 420:78-84 (2002). 144. C. Denoyelle, M. Vasse, M. Korner, et al., Cerivastatin, an inhibitor of HMG-CoA reductase, inhibits the signaling pathways involved in the invasiveness and metastatic properties of highly invasive breast cancer cell lines: an in vitro study, Carcinogenesis 22:1139-48 (2001). 145. K. Pahan, F. G. Sheikh, A. M. Namboodiri, and I. Singh, Lovastatin and phenylacetate inhibit the induction of nitric oxide synthase and cytokines in rat primary astrocytes, microglia, and macrophages, J Clin Invest 100:2671-9 (1997). 146. I. Inoue, F. Itoh, S. Aoyagi, et al., Fibrate and statin synergistically increase the transcriptional activities of PPARalpha/RXRalpha and decrease the transactivation of NFkappaB, Biochem Biophys Res Commun 290:131-9 (2002).


169

147. K. Fassbender, M. Simons, C. Bergmann, et al., Simvastatin strongly reduces levels of Alzheimer’s disease beta -amyloid peptides Abeta 42 and Abeta 40 in vitro and in vivo, Proc Natl Acad Sci U S A 98:5856-61 (2001). 148. M. A. Albert, E. Danielson, N. Rifai, and P. M. Ridker, Effect of statin therapy on Creactive protein levels: the pravastatin inflammation/CRP evaluation (PRINCE): a randomized trial and cohort study, Jama 286:64-70 (2001). 149. R. A. DeBose-Boyd, J. Ou, J. L. Goldstein, and M. S. Brown, Expression of sterol regulatory element-binding protein 1c (SREBP-1c) mRNA in rat hepatoma cells requires endogenous LXR ligands, Proc Natl Acad Sci U S A 98:1477-82 (2001). 150. Z. Sheng, H. Otani, M. S. Brown, and J. L. Goldstein, Independent regulation of sterol regulatory element-binding proteins 1 and 2 in hamster liver, Proc Natl Acad Sci U S A 92:935-8 (1995). 151. F. J. Field, E. Born, S. Murthy, and S. N. Mathur, Regulation of sterol regulatory element-binding proteins in hamster intestine by changes in cholesterol flux, J Biol Chem 276:17576-83 (2001). 152. G. Martin, H. Duez, C. Blanquart, et al., Statin-induced inhibition of the Rhosignaling pathway activates PPARalpha and induces HDL apoA-I, J Clin Invest 107:1423-32 (2001). 153. J. D. Horton, I. Shimomura, M. S. Brown, R. E. Hammer, J. L. Goldstein, and H. Shimano, Activation of cholesterol synthesis in preference to fatty acid synthesis in liver and adipose tissue of transgenic mice overproducing sterol regulatory elementbinding protein-2, J Clin Invest 101:2331-9 (1998). 154. H. Scharnagl, R. Schinker, H. Gierens, M. Nauck, H. Wieland, and W. Marz, Effect of atorvastatin, simvastatin, and lovastatin on the metabolism of cholesterol and triacylglycerides in HepG2 cells, Biochem Pharmacol 62:1545-55 (2001). 155. J. D. Horton, J. L. Goldstein, and M. S. Brown, SREBPs: activators of the complete program of cholesterol and fatty acid synthesis in the liver, J Clin Invest 109:112531 (2002). 156. L. Fajas, K. Schoonjans, L. Gelman, et al., Regulation of peroxisome proliferatoractivated receptor gamma expression by adipocyte differentiation and determination factor 1/sterol regulatory element binding protein 1: implications for adipocyte differentiation and metabolism, Mol Cell Biol 19:5495-503 (1999). 157. C. Ukomadu, and A. Dutta, Inhibition of cdk2 activating phosphorylation by mevastatin, J Biol Chem 278:4840-46 (2003). 158. Y. A. Yang, P. J. Morin, W. F. Han, et al., Regulation of fatty acid synthase expression in breast cancer by sterol regulatory element binding protein-1c, Exp Cell Res 282:132-7 (2003). 159. I. Jialal, D. Stein, D. Balis, S. M. Grundy, B. Adams-Huet, and S. Devaraj, Effect of hydroxymethyl glutaryl coenzyme a reductase inhibitor therapy on high sensitive Creactive protein levels, Circulation 103:1933-5 (2001). 160. U. Laufs, and J. K. Liao, Post-transcriptional regulation of endothelial nitric oxide synthase mRNA stability by Rho GTPase, J Biol Chem 273:24266-71 (1998). 161. A. M. Lefer, R. Scalia, and D. J. Lefer, Vascular effects of HMG CoA-reductase inhibitors (statins) unrelated to cholesterol lowering: new concepts for cardiovascular disease, Cardiovasc Res 49:281-7 (2001). 162. H. Harada, K. P. Pavlick, I. N. Hines, et al., Sexual dimorphism in reduced-size liver ischemia and reperfusion injury in mice: role of endothelial cell nitric oxide synthase, Proc Natl Acad Sci U S A 100:739-44 (2003).

170

Chapter 7

163. J. Malik, V. Melenovsky, D. Wichterle, et al.. Both fenofibrate and atorvastatin improve vascular reactivity in combined hyperlipidaemia (fenofibrate versus atorvastatin trial-FAT), Cardiovasc Res 52:290-8 (2001). 164. S. B. Joseph, E. McKilligin, L. Pei, et al., Synthetic LXR ligand inhibits the development of atherosclerosis in mice, Proc Natl Acad Sci U S A 99:7604-9 (2002B). 165. S. Wassmann, U. Laufs, K. Muller, et al., Cellular antioxidant effects of atorvastatin in vitro and in vivo, Arterioscler Thromb Vasc Biol 22:300-5 (2002). 166. T. J. Aitman, A. M. Glazier, C. A. Wallace, et al., Identification of Cd36 (Fat) as an insulin-resistance gene causing defective fatty acid and glucose metabolism in hypertensive rats, Nat Genet 21:76-83 (1999). 167. N. Qi, L. Kazdova, V. Zidek, et al., Pharmacogenetic evidence that Cd36 is a key determinant of the metabolic effects of pioglitazone, J Biol Chem 277:48501-7 (2002). 168. B. M. Egan, E. L. Greene, and T. L. Goodfriend, Nonesterified fatty acids in blood pressure control and cardiovascular complications, Curr Hypertens Rep 3:107-16 (2001). 169. T. Ichiki, K. Takeda, T. Tokunou, et al., Downregulation of angiotensin II type 1 receptor by hydrophobic 3-hydroxy-3-methylglutaryl coenzyme A reductase inhibitors in vascular smooth muscle cells, Arterioscler Thromb Vasc Biol 21:1896901 (2001). 170. H. Masuzaki, H. Yamamoto, C. J. Kenyon, et al., Transgenic amplification of glucocorticoid action in adipose tissue causes high blood pressure in mice, J Clin Invest 112:83-90 (2003). 171. Y. Zhou, P. Grayburn, A. Karim, et al., Lipotoxic heart disease in obese rats: implications for human obesity, Proc Natl Acad Sci USA 97:1784-1789 (2000). 172. G. M. Corbi, S. Carbone, P. Ziccardi, et al., FFAs and QT intervals in obese women with visceral adiposity: effects of sustained weight loss over 1 year, J Clin Endocrinol Metab 87:2080-3 (2002). 173. J. R. Dyck, and G. D. Lopaschuk, Malonyl CoA control of fatty acid oxidation in the ischemic heart, J Mol Cell Cardiol 34:1099-109 (2002). 174. B. Finck, J. Lehman, T. Leone, et al., The cardiac phenotype induced by PPARalpha overexpression mimics that caused by diabetes mellitus., J Clin Invest 109:121-130 (2002). 175. T. Ramanathan, K. Shirota, S. Morita, et al., Left ventricular oxygen utilization efficiency is impaired in chronic streptozotocin-diabetic sheep, Cardiovasc Res 55:749-56 (2002). 176. S. Boudina, P. Mazumder, R. Cooksey, D. McClain, and E. Abel, Mitochondrial impairment contributes to cardiac dysfunction in obese leptin-deficient (ob/ob) mice, Circulation 108:IV-16 (Abstract) (2003). 177. M. Higa, Y. Zhou, M. Ravazzola, D. Baetens, L. Orci, and R. Unger, Troglitazone prevents mitochondrial alterations, beta cell destruction, and diabetes in obese prediabetic rats, Proc Natl Acad Sci USA 96:11513-11518 (1999). 178. S. Piro, M. Anello, C. Di Pietro, et al., Chronic exposure to free fatty acids or high glucose induces apoptosis in rat pancreatic islets: possible role of oxidative stress, Metabolism 51:1340-7 (2002). 179. K. Sakai, K. Matsumoto, T. Nishikawa, et al., Mitochondrial reactive oxygen species reduce insulin secretion by pancreatic beta-cells, Biochem Biophys Res Commun 300:216-22 (2003).


171

180. P. Oliveira, A. Rolo, V. Sardão, P. Coxito, C. Palmeira, and A. Moreno, Carvedilol in heart mitochondria protonophore or opener of the mitochondrial KATP channels?, Life Sci 69:123-132 (2001). 181. L. X. Li, F. Skorpen, K. Egeberg, I. H. Jorgensen, and V. Grill, Induction of uncoupling protein 2 mRNA in beta-cells is stimulated by oxidation of fatty acids but not by nutrient oversupply, Endocrinology 143:1371-7 (2002). 182. J. C. Yoon, G. Xu, J. T. Deeney, et al., Suppression of beta cell energy metabolism and insulin release by PGC-1alpha, Dev Cell 5:73-83 (2003). 183. M. Lehtihet, N. Welsh, P. O. Berggren, G. A. Cook, and A. Sjoholm, Glibenclamide inhibits islet carnitine palmitoyltransferase 1 activity, leading to PKC-dependent insulin exocytosis, Am J Physiol Endocrinol Metab 285:E438-46 (2003). 184. C. E. Wrede, L. M. Dickson, M. K. Lingohr, I. Briaud, and C. J. Rhodes, Protein kinase B/Akt prevents fatty acid-induced apoptosis in pancreatic beta-cells (INS-1), J Biol Chem 277:49676-84 (2002). 185. Y. A. Yang, W. F. Han, P. J. Morin, F. J. Chrest, and E. S. Pizer, Activation of fatty acid synthesis during neoplastic transformation: role of mitogen-activated protein kinase and phosphatidylinositol 3-kinase, Exp Cell Res 279:80-90 (2002). 186. M. Ristow, H. Mulder, D. Pomplun, et al., Frataxin deficiency in pancreatic islets causes diabetes due to loss of beta cell mass, J Clin Invest 112:527-34 (2003). 187. J. E. Schaffer, Lipotoxicity: when tissues overeat, Curr Opin Lipidol 14:281-7 (2003). 188. R. H. Unger, and L. Orci, Lipoapoptosis: its mechanism and its diseases, Biochim Biophys Acta 1585:202-12 (2002). 189. W. El-Assad, J. Buteau, M.-L. Peyot, et al., Saturated fatty acids synergize with elevated glucose to cause pancreatic beta-cell death., Endocrinology 144:4154-4163 (2003). 190. R. Branstrom, I. B. Leibiger, B. Leibiger, B. E. Corkey, P. O. Berggren, and O. Larsson, Long chain coenzyme A esters activate the pore-forming subunit (Kir6. 2) of the ATP-regulated potassium channel, J Biol Chem 273:31395-400 (1998). 191. I. Leclercq, G. Farrell, J. Field, D. Bell, F. Gonzalez, and G. Robertson, CYP2E1 and CYP4A as microsomal catalysts of lipid peroxides in murine nonalcoholic steatohepatitis, J Clin Invest 105:1067-1075 (2000). 192. A. Sanyal, C. Campbell-Sargent, F. Mirshahi, et al., Nonalcoholic steatohepatitis: association of insulin resistance and mitochondrial abnormalities, Gastroenterol 120:1281-1285 (2001). 193. B. A. Neuschwander-Tetri, and S. H. Caldwell, Nonalcoholic steatohepatitis: summary of an AASLD Single Topic Conference, Hepatology 37:1202-19 (2003). 194. C. L. Limoli, E. Giedzinski, W. F. Morgan, S. G. Swarts, G. D. Jones, and W. Hyun, Persistent oxidative stress in chromosomally unstable cells, Cancer Res 63:3107-11 (2003). 195. M. M. Hassan, L. Y. Hwang, C. J. Hatten, et al., Risk factors for hepatocellular carcinoma: synergism of alcohol with viral hepatitis and diabetes mellitus, Hepatology 36:1206-13 (2002). 196. J. A. Marrero, R. J. Fontana, G. L. Su, H. S. Conjeevaram, D. M. Emick, and A. S. Lok, NAFLD may be a common underlying liver disease in patients with hepatocellular carcinoma in the United States, Hepatology 36:1349-54 (2002). 197. E. E. Calle, C. Rodriguez, K. Walker-Thurmond, and M. J. Thun, Overweight, obesity, and mortality from cancer in a prospectively studied cohort of U.S. adults, N Engl J Med 348:1625-38 (2003).

172

Chapter 7

198. K. West, J. Borgnard, A. Clark, et al., Rapid Akt activation by nicotine and a tobacco carcinogen modulates the phenotype of normal human airway epithelial cells, J Clin Invest 111:81-90 (2003). 199. G. D. Lopaschuk, Alterations in fatty acid oxidation during reperfusion of the heart after myocardial ischemia, Am J Cardiol 80:11A-16A (1997). 200. T. Vanden Hoek, L. Becker, Z. Shao, C. Li, and P. Schumacker, Reactive oxygen species released from mitochondria during brief hypoxia induce preconditioning in cardiomyocytes, J Biol Chem 273:18092-18098 (1998). 201. Y. Yue, M. Krenz, M. Cohen, J. Downey, and S. Critz, Menadione mimics the infarct-limiting effect of preconditioning in isolated rat hearts, Am J Physiol Heart Circ Physiol 281:H590-595 (2001). 202. H. Jaeschke, Molecular mechanisms of hepatic ischemia-reperfusion injury and preconditioning, Am J Physiol Gastrointest Liver Physiol 284:G15-26 (2003). 203. G. Lebuffe, P. T. Schumacker, Z. H. Shao, T. Anderson, H. Iwase, and T. L. Vanden Hoek, ROS and NO trigger early preconditioning: relationship to mitochondrial KATP channel, Am J Physiol Heart Circ Physiol 284:H299-308 (2003). 204. P. Kantor, A. Lucien, R. Kozak, and G. Lopaschuk, The antianginal drug trimetazidine shifts cardiac energy metabolism from fatty acid oxidation to glucose oxidation by inhibiting mitochondrial long-chain 3-ketoacyl coenzyme A thiolase, Circ Res 86:580-588 (2000). 205. D. Portilla, G. Dai, J. Peters, F. Gonzalez, M. Crew, and A. Proia, Etomoxir-induced PPARa-modulated enzymes protect during acute renal failure, Am J Physiol Renal Physiol 278:F667-F675 (2000). 206. R. Cornelussen, G. Van Der Vusse, T. Roemen, and L. Snoeckx, Heat pretreatment differentially affects cardiac fatty acid accumulation during ischemia and postischemic reperfusion., Am J Physiol Heart Circ Physiol 280:H1736-H1743 (2001). 207. M. Taniguchi, C. Wilson, C. Hunter, D. Pehowich, A. Clanachan, and G. Lopaschuk, Dichloroacetate improves cardiac efficiency after ischemia independent of changes in mitochondrial proton leak, Am J Physiol Heart Circ Physiol 280:H1762-H1769 (2001). 208. I. Xavier, P. Mercier, C. McLoughlin, A. Ali, J. Woodgett, and N. Ovsenek, Glycogen synthase kinase 3b negatively regulates both DNA-binding and transcriptional activities of heat shock factor 1, J Biol Chem 275:29147-29152 (2000). 209. N. Collier, M. Sheetz, and M. Schlesinger, Concomitant changes in mitochondria and intermediate filaments during heat shock and recovery of chicken embryo fibroblasts., J Cellular Biochem 52:297-307 (1993). 210. K. Djabali, N. De, B, F. Lon, and M. Portier, AlphaB-crystallin interacts with intermediate filaments in response to stress., J Cell Sci 110:2759-2769 (1997). 211. I. Izawa, M. Nishizawa, K. Ohtakara, K. Ohtsuka, H. Inada, and M. Inagaki, Identification of Mrj, a DnaJ/Hsp40 family protein, as a keratin 8/18 filament regulatory protein, J Biol Chem 275:34521-7 (2000). 212. G. Velasco, D. Gómez, Pulgar, T, D. Carlin, and M. Guzmán, Evidence that the AMPactivated protein kinase stimulates rat liver carnitine palmitoyltransferase I by phosphorylating cytoskeletal components, FEBS Lett 439:317-320 (1998A). 213. G. Velasco, M. Geelen, T. Gomez del Pulgar, and M. Guzmán, Malonyl-CoAindependent acute control of hepatic carnitine palmitoyltransferase I activity: Role of Ca2+/calmodulin-dependent protein kinase II and cytoskeletal components, J Biol Chem 273:21497-21504 (1998B).


173

214. S. I. Han, S. Y. Oh, W. J. Jeon, et al., Mild heat shock induces cyclin D1 synthesis through multiple Ras signal pathways, FEBS Lett 515:141-5 (2002). 215. K. Takuma, K. Mori, E. Lee, R. Enomoto, A. Baba, and T. Matsuda, Heat shock inhibits hydrogen peroxide-induced apoptosis in cultured astrocytes, Brain Res 946:232-8 (2002). 216. F. Lee, Y. Li, H. Zhu, et al., Tumor necrosis factor increases mitochondrial oxidant production and induces expression of uncoupling protein-2 in the regenerating mouse liver, Hepatology 29:677-687 (1999). 217. K. M. Mearow, M. E. Dodge, M. Rahimtula, and C. Yegappan, Stress-mediated signaling in PC12 cells - the role of the small heat shock protein, Hsp27, and Akt in protecting cells from heat stress and nerve growth factor withdrawal, J Neurochem 83:452-62 (2002). 218. S. Sato, N. Fujita, and T. Tsuruo, Modulation of Akt kinase activity by binding to Hsp90, Proc Natl Acad Sci USA 97:10832-10837 (2000). 219. A. D. Basso, D. B. Solit, G. Chiosis, B. Giri, P. Tsichlis, and N. Rosen, Akt forms an intracellular complex with heat shock protein 90 (Hsp90) and Cdc37 and is destabilized by inhibitors of Hsp90 function, J Biol Chem 277:39858-66 (2002). 220. D. Arsenijevic, H. Onuma, C. Pecqueur, et al., Disruption of the uncoupling protein2 gene in mice reveals a role in immunity and reactive oxygen species production, Nat Genet 26:435-9 (2000). 221. K. J. Mukamal, K. M. Conigrave, M. A. Mittleman, et al., Roles of drinking pattern and type of alcohol consumed in coronary heart disease in men, N Engl J Med 348:109-18 (2003A). 222. A. Ruitenberg, J. C. van Swieten, J. C. Witteman, et al., Alcohol consumption and risk of dementia: the Rotterdam Study, Lancet 359:281-6 (2002). 223. K. J. Mukamal, L. H. Kuller, A. L. Fitzpatrick, W. T. Longstreth, Jr., M. A. Mittleman, and D. S. Siscovick, Prospective study of alcohol consumption and risk of dementia in older adults, Jama 289:1405-13 (2003B). 224. J. B. Hoek, A. Cahill, and J. G. Pastorino, Alcohol and mitochondria: a dysfunctional relationship, Gastroenterology 122:2049-63 (2002). 225. M. M. da Silva, A. Sartori, E. Belisle, and A. J. Kowaltowski, Ischemic preconditioning inhibits mitochondrial respiration, increases H2O2 release, and enhances K+ transport, Am J Physiol Heart Circ Physiol 285:H154-62 (2003). 226. D. M. Valks, T. J. Kemp, and A. Clerk, Regulation of Bcl-xL expression by H2O2 in cardiac myocytes, J Biol Chem 278:25542-7 (2003). 227. Y. Yaguchi, H. Satoh, N. Wakahara, et al., Protective effects of hydrogen peroxide against ischemia/reperfusion injury in perfused rat hearts, Circ J 67:253-8 (2003). 228. K. R. Valeur, and R. degli Agosti, Simulations of temperature sensitivity of the peroxidase-oxidase oscillator, Biophys Chem 99:259-70 (2002). 229. M. Sakoh, and A. Gjedde, Neuroprotection in hypothermia linked to redistribution of oxygen in brain, Am J Physiol Heart Circ Physiol 285:H17-25 (2003). 230. H. Z. Zhou, J. S. Karliner, and M. O. Gray, Moderate alcohol consumption induces sustained cardiac protection by activating PKC-epsilon and Akt, Am J Physiol Heart Circ Physiol 283:H165-74 (2002). 231. N. Teoh, A. Dela Pena, and G. Farrell, Hepatic ischemic preconditioning in mice is associated with activation of NF-kappaB, p38 kinase, and cell cycle entry, Hepatology 36:94-102 (2002). 232. L. He, and J. J. Lemasters, Heat shock suppresses the permeability transition in rat liver mitochondria, J Biol Chem 278:16755-60 (2003).

174

Chapter 7

233. C. Diez-Fernandez, D. Andres, and M. Cascales, Attenuating effects of heat shock against TGF-betal-induced apoptosis in cultured rat hepatocytes, Free Radic Biol Med 33:835-46 (2002). 234. R. Ockner, N. Lysenko, N. Wu, and N. Bass, Hepatocyte growth inhibitors modulate mitochondrial and extramitochondrial fatty acid oxidation, Hepatology 24:253A (Abstract) (1996). 235. J. Dypbukt, M. Ankarcrona, M. Burkitt, et al., Different prooxidant levels stimulate growth, trigger apoptosis, or produce necrosis of insulin-secreting RINm5F cells: The role of intracellular polyamines., J Biol Chem 269:30553-30560 (1994). 236. M. Benhar, D. Engelberg, and A. Levitzki, ROS, stress-activated kinases and stress signaling in cancer, EMBO Rep 3:420-5 (2002). 237. C. Sen, and L. Packer, Antioxidant and redox regulation of gene transcription, FASEB J 10:709-720 (1996). 238. V. Lakshminarayanan, E. Drab-Weiss, and K. Roebuck, and tumor necrosis factor-alpha induce differential binding of the redox-responsive transcription factors AP-1 and NF-kappaB to the interleukin-8 promoter in endothelial and epithelial cells, J Biol Chem 273:32670-32678 (1998). 239. E. Shaulian, and M. Karin, AP-1 as a regulator of cell life and death, Nat Cell Biol 4:E131-6 (2002). 240. S. Lee, K. Kwon, S. Kim, and S. Rhee, Reversible inactivation of protein-tyrosine phosphatase 1B in A431 cells stimulated with epidermal growth factor, J Biol Chem 273:15366-15372 (1998). 241. W. Barrett, J. DeGnore, Y. Keng, Z. Zhang, M. Yim, and P. Chock, Roles of superoxide radical anion in signal transduction mediated by reversible regulation of protein-tyrosine phosphatase IB., J Biol Chem 274:34543-34546 (1999). 242. N. Chandel, D. McClintock, C. Feliciano, et al., Reactive oxygfen species generated at mitochondrial complex III stabilize hypoxia-inducible factor-1alpha during hypoxia: a mechanism of O2 sensing., J Biol Chem 275:25130-25138 (2000). 243. C. Schroedl, D. S. McClintock, G. R. Budinger, and N. S. Chandel, Hypoxic but not anoxic stabilization of HIF-1alpha requires mitochondrial reactive oxygen species, Am J Physiol Lung Cell Mol Physiol 283:L922-31 (2002). 244. G. Semenza, HIF-1 and tumor progression: pathophysiology and therapeutics., Trends Molec Med 8:S62-S67 (2002). 245. P. Garcia-Nogales, A. Almeida, and J. Bolaños, Peroxynitrite protects neurons against nitric oxide-mediated apoptosis. A key role for glucose-6-phosphate dehydrogenase activity in neuroprotection., J Biol Chem 278:864-874 (2003). 246. V. Nilakantan, B. Spear, and H. Glauert, Liver-specific catalase expression in transgenic mice inhibits NF-kappaB activation and DNA synthesis induced by the peroxisome proliferator ciprofibrate, Carcinogenesis 19:631-637 (1998). 247. T. Finkel, Redox-dependent signal transduction., FEBS Letters 476:52-54 (2000A). 248. V. Thannickal, R. Day, S. Klinz, M. Bastien, J. Larios, and B. Fanburg, Rasdependent and -independent regulation of reactive oxygen species by mitogenic growth factors and TGF-b1, FASEB J 14:1741-1748 (2000). 249. S. Nemoto, and T. Finkel, Redox regulation of forkhead proteins through a p66shcdependent signaling pathway, Science 295:2450-2452 (2002). 250. K. F. Petersen, D. Befroy, S. Dufour, et al., Mitochondrial dysfunction in the elderly: possible role in insulin resistance, Science 300:1140-2 (2003). 251. A. L. Hsu, C. T. Murphy, and C. Kenyon, Regulation of aging and age-related disease by DAF-16 and heat-shock factor, Science 300:1142-5 (2003).


175

252. S. S. Korshunov, V. P. Skulachev, and A. A. Starkov, High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria, FEBS Lett 416:15-8 (1997). 253. S.-S. Liu, Cooperation of a “reactive oxygen cycle” with the Q cycle and the proton cycle in the respiratory chain--superoxide generating and cycling mechanisms in mitochondria, J Bioenerg Biomembr 31:367-376 (1999). 254. I. Lee, E. Bender, and B. Kadenbach, Control of mitochondrial membrane potential and ROS formation by reversible phosphorylation of cytochrome c oxidase, Mol Cell Biochem 234-235:63-70 (2002).

Chapter 8 Metabolism and Gene Expression in Liver Regeneration

8.1.

INTRODUCTION

EGF, HGF, and transforming growth are potent hepatocyte mitogens, but their effect in normal animals in vivo is limited or inapparent except under conditions in which a not fully characterized “priming” factor has rendered hepatocytes competent to proliferate1. Clues to the nature of this priming, and of the competence which it confers, are provided by the metabolic and molecular responses to 70% hepatectomy, studied in greatest detail in the rat. Analysis of these responses suggests their separation into two more or less temporally and functionally distinct phases, the relationship of which to eventual hepatocyte mitosis can be understood in terms of the overall model of growth regulation that has been presented and developed in the preceding chapters.

8.2.

SYSTEMIC METABOLIC RESPONSES TO 70% HEPATECTOMY

The organism is threatened immediately by a precipitous reduction in both the major repository of readily available glucose (hepatic glycogen), and the major mechanism for its replenishment (hepatic gluconeogenesis). Plasma glucose and insulin fall rapidly, while concentrations and secretion rates of glucagon, catecholamines, corticosteroids, and other counterregulatory hormones rise2,3. These responses increase adipocyte hormone-sensitive lipase activity and FFA 177

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Figure 8.1. Fatty acid oxidation and gluconeogenesis; suppressed glycolysis and fatty acid synthesis. Enclosed areas represent mitochondria. This pattern of substrate utilization is antiproliferative. Arrow thickness approximates relative pathway a c t i v i t y ; dotted line: pathway suppressed during growth arrest. Numbers in parentheses indicate approximate yields of ATP from complete oxidation of palmitate (-100), compared (in a ketogenic cell, e.g., hepatocyte) with complete diversion to ketogenesis (-20). Fatty acids activate transcriptionally upregulating both mitochondrial and extramitochondrial fatty acid oxidation. When present in excess, fatty acids may override malonyl CoA inhibition of CPT-I. Abbreviations — ATP: adenosine triphosphate; CPTI: carnitine palmitoyltransferase-I; EMFO: extramitochondrial fatty acid oxidation in peroxisomes, and in endoplasmic reticulum); F26BP: fructose-2,6-bisphosphate; PEPCK: phosphoenolpyruvate carboxykinase; PL: phospholipids, i.e., glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

release into plasma, a process that is augmented by increased plasma [HGF]4. The rapid early rise in plasma [FFA]5 drives an increase in FFA uptake per unit mass of the hepatic remnant6. This starvation-like response is accompanied by changes in glucose metabolism in the remaining hepatocytes7. Glycogenolysis increases2.3, and hepatocellular consumption of glucose (glycolysis) is curtailed. These immediately available mechanisms to support increased glucose export, and thus plasma glucose concentration, are augmented by increased gluconeogenesis (Fig. 8.1). Gluconeogenesis utilizes as

8. Metabolism and Gene Expression in Liver Regeneration

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substrates amino acids, lactate, and glycerol mobilized from the periphery, and is fueled energetically by greatly increased mitochondrial oxidation of fatty acids, now abundantly available to the hepatocyte. Together, these changes serve to limit the fall in plasma glucose and to provide for its rapid restoration toward levels needed to maintain the function of the central nervous system. Under these conditions as during fasting8 and in the tumor-bearing mouse studies cited above9 (also, see Chapter 4.2.: Fatty Acids), the central nervous system would be expected to become the major consumer of plasma glucose. This reflects suppression of glucose utilization in skeletal muscle and other peripheral sites after partial hepatectomy by the combined effects of low plasma insulin and high concentrations of counterregulatory hormones and FFA10-21. Renal gluconeogenesis and glucose export 22 also contribute to this early response, as these increase significantly after partial hepatectomy23 and are less susceptible to suppression by insulin than in liver24. The renal contribution thus accelerates normalization of plasma glucose and insulin, and may be essential for the subsequent hepatocellular shift from gluconeogenesis to glycolysis that is required for cell cycle entry (see this Chapter 8.4.: Ensuing Modulation of Systemic and Hepatocellular Metabolism). Thus, acute post-hepatectomy starvation-like conditions limit systemic glucose consumption as fatty acids become the dominant fuel except in the central nervous system. Plasma glucose and insulin levels may be improved subsequently in animals with access to food25, but restoration of glucose supply depends to a large extent on hepatic and renal gluconeogenesis, fueled by increased FFA mobilization and increased fatty acid uptake and mitochondrial oxidation. Visceral adipose tissue is relatively active metabolically and insulin resistant26; thus, despite rising insulin levels, it may assume special significance by continuing to provide FFA directly to liver via the portal vein. Moreover, although the excess of fatty acids with which the hepatocyte is confronted suppresses glycolysis and activates AMPK-associated antiproliferative signaling27, augmented FFA flux may be a necessary antecedent to subsequent hepatocyte proliferation. This possibility is suggested by the observation that while early administration of exogenous glucose normalizes hepatocellular ATP and plasma glucose, insulin, and glucagon, thus suppressing lipolysis and plasma FFA concentration5, it paradoxically delays both DNA synthesis and restoration of hepatic mass28,29. An obvious effect of exogenous glucose under these conditions is its augmentation of plasma insulin levels; in turn, insulin’s supression of adipose tissue hormonesensitive lipase would decrease the availability of essential FFA required for membrane and eicosanoid biogenesis. The opposite result

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follows treatment of rats with clofibrate, a peroxisome proliferator that through its inhibitory effect on CPT-I activity suppresses mitochondrial fatty acid oxidation relative to an increasing mitochondrial oxidative capacity (see Chapter 5.2.1.: In this case, the resulting clofibrate-induced limitation in intramitochondrial generation of ROS, and increased hepatocyte extramitochondrial abundance of fatty acids (including accelerates the onset of DNA synthesis and the restoration of mitochondrial protein and liver mass after hepatectomy30. 31 L-FABP expression, required for activation of , is also increased by generally parallels that of the extramitochondrial fatty acid oxidation enzymes32,33, and is increased during regenerative hepatocyte mitosis34,35. Expression of the cytochrome P450 4A is strongly increased36, thereby augmenting generation of the long chain dicarboxylic fatty acids that also activate the overall response32,33. The importance of these pathways is supported by the delay in regeneration that is observed in mice36. Thus, the initial stages of regeneration after partial hepatectomy depend importantly upon the availability of increased quantities of FFA, including the essential fatty acids derived from storage in adipose tissue TG, and presumably the long chain dicarboxylic fatty acid activators of that are generated via extramitochondrial oxidative pathways in the hepatocyte.

8.3.

EARLY CHANGES IN HEPATOCELLULAR METABOLISM AND GENE EXPRESSION

The initial starvation-like increases in systemic availability of fatty acids and gluconeogenic precursors (e.g., glycerol, amino acids) are paralleled by major changes in hepatocellular metabolism and gene expression (Figs. 8.1, 8.2). Consistent with diminished insulin effect37, increased mitochondrial fatty acid oxidation-fueled gluconeogenesis and glucose export reflect rapid increases in the expression of PEPCK and glucose-6-phosphatase mRNAs38,39, which peak within a few hours40. Conversely, hepatic utilization of glucose (glycolysis), and expression of glycolytic enzymes, including glucokinase, PFK2, pyruvate kinase, and pyruvate dehydrogenase are suppressed27,38-41. As discussed earlier, this metabolic pattern is directly opposite to that associated with, and required for, cell proliferation. In this setting, the significance of an early transient increase in nuclear abundance 42 is unclear; its later, more sustained, elevation is consistent with the onset of regenerative mitosis.


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Figure 8.2. Contrasting patterns of intermediary metabolism in cancer cachexia and liver regeneration. In early liver regeneration and in hepatocytes of cachectic cancer patients, increased mitochondrial fatty acid oxidation is associated with increased gluconeogenesis and oxidative stress, and decreased and ATP. In contrast, in late regeneration (proliferative) hepatocytes and in cancer cells, glycolysis, ATP abundance, and fatty acid synthesis are increased, while fatty acid oxidation is suppressed. Under hypoxic conditions, glycolysis in normal and cancer cells is largely anaerobic, survival mechanisms are activated, and suppressed fatty acid oxidation implicit (see Chapter 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation). Abbreviations — mitochondrial inner membrane potential; FA: fatty acid; NA: data not available

The now virtually exclusive hepatocellular dependence on fatty acid oxidation for ATP generation5 is met by increases in FFA supply and CPT-I expression43,44, and an almost certainly decreased CPT-I responsiveness to inhibition by malonyl CoA45-49, itself limited in abundance50,51. “Wasteful” export of potentially oxidizable fatty acid carbon atoms in the form of ketone bodies is curtailed43,52,53, reflecting decreased expression of mitochondrial HMGCoA synthase43, and preferential utilization of acetyl CoA in citrate synthesis and citric acid cycle-dependent ATP generation52. However, this abundance of long chain fatty acids exerts adverse effects on mitochondrial function, including an uncoupling of oxidative phosphorylation54-58, as well as decreases in the subunit of the ATP synthase59-61, and the efficiency of ATP generation59,61. Thus, cellular energy charge falls5,62,63. Perturbed electron transport chain function under these conditions of high fatty acid oxidation rates59-61,64-66 overwhelms amelioration of ROS formation that might otherwise result from the fatty acid uncoupling effect67-71. Intramitochondrial oxidative stress and lipid peroxidation are increased, consistent with fatty acid

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override of CPT-I control45-49 and thus increased abundance of electrons in the electron transport chain. The resulting increase in electron “leakage” and generation is reflected by increases in malondialdehyde concentration in mitochondria 51 and whole liver50, and in activity of mitochondrial managanese superoxide dismutase51, and NO generation 72 , as well as by a decrease in mitochondrial glutathione 51,60 . In addition, there is evidence of peroxisome proliferation 73 and an increase in peroxisomal palmitoyl CoA oxidase activity within 12 hr 74 , supporting the concept that augmented cellular fatty acid flux has enhanced extramitochondrial fatty acid oxidation pathways (i.e., peroxisomal and cytochrome P450 4A and P450 2E1 in the endoplasmic reticulum). As a result, oxidative stress generated at the electron transport chain is compounded by that produced by extramitochondrial generation of ROS75,76. In addition to the changes in fatty acid oxidative metabolism, increased hepatocellular fatty acid flux after partial hepatectomy is reflected in substantially increased TG biosynthesis and VLDL secretion in the remaining hepatocytes77. The initial increase in mitochondrial fatty acid oxidation also provides the energy required for gluconeogenesis, while inhibiting pyruvate dehydrogenase and glycolysis. Together, these conditions sustain the approximately 12-16 hour-long state of growth arrest that precedes the start of replicative DNA synthesis. Given the energetically precarious state of the hepatocyte that is implied by this constellation of adverse effects, it is not difficult to understand the mechanistic basis for the lethal consequences of excessive hepatic resection, or of transplantation of insufficient liver mass.

8.4.

ENSUING MODULATION OF SYSTEMIC AND HEPATOCELLULAR METABOLISM

Fostered by elevations in plasma FFA, hepatic and renal gluconeogenesis are increased and systemic glucose consumption curtailed, thus normalizing plasma levels of glucose and insulin62,78 , and hepatic PEPCK expression79, within a few hours. By approximately 6 to 12 hours, hepatocellular energy charge has reached a nadir 62 and also begins to increase as indicators of mitochondrial oxidative stress decrease51, further suggesting that the relationship of fatty acid availability and oxidation to overall mitochondrial fatty acid oxidative capacity also may be normalizing (Fig. 8.2). Consistent with this concept, wg/Wnt-like signaling is augmented42; mitochondrial DNA, state 3 respiration, coupling of oxidative phosphorylation, and the activities of various


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mitochondrial enzymes increase64,80, approximately paralleled by ubiquinone abundance65. At 12 hours post-hepatectomy, rat liver glutathione concentration and synthesis are increased81. The evolving changes in redox balance as hepatocytes progress toward mitosiscompetence are consistent with evidence that high levels of oxidative stress that inhibit respiratory chain components including ubiquinone 66 cause cell necrosis or apoptosis; in contrast, lower levels may induce growth82,86 (also, see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable).

Figure 8.3. Glycolysis- and citric acid cycle-fueled biosynthesis of ATP and fatty acid; suppressed fatty acid oxidation and gluconeogenesis. This figure represents the predominant pattern of substrate utilization in proliferating cells; enclosed areas represent mitochondria. Arrow thickness approximates relative pathway activity; dotted lines: pathways that are suppressed during cell proliferation. Fatty acids also may be derived from plasma. Abbreviations — CPT-I: carnitine palmitoyltransferase-I (shown as inhibited by malonyl CoA); EMFO: extramitochondrial fatty acid oxidation in peroxisomes, and in endoplasmic reticulum); F26BP: fructose-2,6bisphosphate; PEPCK: phosphoenolpyruvate carboxykinase; PL: glycerophosphatides; peroxisome proliferator-activated TG: triacylglycerols; VLDL: very low density lipoproteins.

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Augmented hepatocellular utilization of glucose by 24 hours is suggested by an increase in de novo fatty acid synthesis52, as this process depends on both glycolytic and pentose phosphate pathway activity (Fig. 8.3). Increased fatty acid synthesis despite ongoing fatty acid oxidation would appear to constitute a potentially wasteful futile cycle, inconsistent with the usually reciprocal activation of these opposing pathways. However, the methods utilized in the development of these important data52 would have precluded recognition of the metabolic heterogeneity among hepatocytes that reflects sequential initiation of DNA synthesis and mitosis from periportal to pericentral regions of the hepatic lobule87,88. By 24 hours, mitochondrial ATP production, coupling of oxidative phosphorylation, and actually exceed normal resting levels64,89-91. The associated increase in hepatic oxygen utilization is supported by increased extraction of oxygen from sinusoidal blood, and is reflected in depression of hepatic venous oxygen saturation to levels below basal92. The importance of accelerated ATP biosynthesis is supported by the observation that in transgenic mice expressing creatine kinase in liver (thus providing for an auxiliary cytosolic high-energy phosphate reserve), ATP levels, DNA synthesis, and liver weight gain after 70% hepatectomy are enhanced93. The hepatocyte, up to this stage having been committed to fatty acid oxidation-fueled gluconeogenesis and growth arrest, is now poised energetically for what is in effect a 180 degree shift, to glycolysis, fatty acid synthesis, and cell cycle progression (Figs. 8.1, 8.2, 8.3). Not surprisingly, the regenerative process may be delayed in obese animals and in patients whose livers are steatotic prior to partial hepatectomy94; in both settings, prolongation of a fatty acid oxidation-induced impairment of growth may be anticipated.

8.4.1. Effects of Proinflammatory Cytokines; Insulin

vs.

The mechanism by which hepatocytes survive after 70% hepatectomy despite the adverse effects of increased fatty acid flux on mitochondrial function, and thus avoid the progressive deterioration that is characteristic of more extensive hepatic resection5, has been unclear. Recent evidence suggests that proinflammatory cytokines contribute importantly to this survival. Thus, expression and secretion of and other proinflammatory cytokines including IL-1 and IL-6 are increased after 70% hepatectomy, and appear to be fundamental to the regenerative process95,96, while IL-6 increases survival of transplanted steatotic livers97. Kupffer cell release of these mediators is promoted by ICAM98. A role


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for proinflammatory cytokines is also suggested by enhancement of liver size and incorporation in intact rats99, and IL-6 induction of mouse hepatocyte proliferation in primary culture100. Additionally, antibodies to inhibit hepatocyte proliferation induced by partial hepatectomy101, and abrogate the protection against apoptosis otherwise afforded by peroxisome proliferators in hepatocyte cultures102. Regeneration is also inhibited by deficiency of IL-6103 or of receptors for (TNFR-1), the latter correctable by administration of exogenous IL-6104. Gut-derived bacterial lipopolysaccharide (LPS) enhances post-hepatectomy Kupffer cell expression, perhaps providing an explanation for the delayed hepatic regeneration observed in germ-free animals95. However, clearly has cytotoxic potential105,106, mediates the adverse effects of lipopolysaccharide105,106, and disrupts mitochondrial electron transport107. Thus, positive participation of the proinflammatory cytokines in liver regeneration would seem difficult to reconcile with the energy requirements of cell proliferation. Advances in the understanding of proinflammatory cytokine biology help to clarify this matter108. and related proinflammatory cytokines are functionally linked to systemic fatty acid metabolism16, and adipocyte expression is increased in obesity. Along with elevated plasma [FFA], contributes to systemic insulin resistance, at least in part by inhibiting insulin receptor signaling109. A more explicit connection to fatty acid metabolism is suggested by evidence that adipocyte expression requires expression of aP2, the adipocyte form of cytosolic FABP110. In view of the importance of Kupffer cells in overall economy, increased expression after partial hepatectomy may be related, as noted, to increased Kupffer cell uptake and utilization of FFA (see Chapter 5.2.1.: and Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). IL-1, and IL-6 (and through these mediators LPS) exert profound and variably overlapping effects on hepatocellular fatty acid metabolism108, both in vivo108,111,112 and in vitro113,114. These agents stimulate lipolysis in adipose tissue111, thereby increasing plasma [FFA]. The rise in [FFA] drives increased hepatocellular fatty acid uptake, The despite acutely decreased hepatic expression of L-FABP115. 108,112,116-118, cytokines also enhance hepatocyte glycogenolysis, glycolysis and de novo lipogenesis108; as a result, cytosolic malonyl CoA is increased112, thus inhibiting CPT-I, mitochondrial fatty acid oxidation and ketogenesis112,117. The cytokine-induced decrease in mitochondrial fatty acid oxidation also reflects differential regulation of acyl CoA synthase expression, which decreases in mitochondria and increases in the

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endoplasmic reticulum119. After a single acute administration of to normal rats, peroxisomal fatty acid mRNA, protein, and enzyme activity, and mRNA and protein were inhibited120,121, a finding that appears to be inconsistent with the previously cited evidence of increased peroxisomal acyl CoA oxidase activity at 12 hr after partial hepatectomy74. Accordingly, while such inhibition of the peroxisomal system would serve to decrease the utilization of alternative (non-ATPgenerating) pathways for oxidative metabolism of long chain fatty acids, the relevance of these observations in normal animals to the complex evolution of events following hepatectomy is unclear. Thus, under the influence of the proinflammatory cytokines and greater availability of glucose, hepatocellular cytosolic fatty acid abundance is increased: augmented FFA uptake from plasma and de novo biosynthesis acting in concert with decreased mitochondrial fatty acid oxidation, favor diversion of fatty acids toward esterification pathways. In the setting of systemic infection, similar compartmentalization accounts for the recognized proinflammatory cytokine-induced increases in TG biosynthesis, VLDL TG secretion, and hypertriglyceridemia111. In the residual post-hepatectomy hepatocyte, such changes would serve to promote formation of the glycerophosphatides114 required for membrane biogenesis and lipid signaling122. Moreover, similar to the effects of insulin and peroxisome proliferators, proinflammatory cytokines decrease mitochondrial matrix reducing potential, as reflected in decreased ratios of to [acetoacetate] and of [NADH] to [NAD]112,118; these favorable redox changes are associated with increases in state 3 These circumstances, which are respiration and ATP generation118. consistent with proinflammatory cytokine-induced inhibition of mitochondrial fatty acid oxidation, imply a relative or absolute increase in glycolysis and approximate the metabolic requirements for cell proliferation. In the setting of partial hepatectomy, however, sustained increases in hepatocellular fatty acid abundance could override malonyl CoA inhibition of CPT- 45-49, suggesting that one or more additional factors, e.g., malonyl CoA-independent cytoskeletal regulation of CPTI123,124, may be required to constrain mitochondrial fatty acid oxidation in this situation. In the aggregate, available evidence indicates that and other proinflammatory cytokines inhibit fatty acid while enhancing glycolysis, mitochondrial energetics, lipogenesis, and fatty acid esterification. In these respects they resemble the effects of other growth promoters such as insulin and the peroxisome proliferators. However, the cytokine effects are unlikely to be mediated by changes in plasma insulin.


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as they are also observed in cell culture 113 and because high concentrations of are required to increase insulin levels in 117,125 vivo . Thus, the hepatocellular effects of proinflammatory cytokines are directly conducive to cell proliferation: they support the state of increased glycolysis and lipogenesis that prevails when DNA replication begins after partial hepatectomy, while contributing to suppression of the high rates of mitochondrial fatty acid oxidation and gluconeogenesis that prevail during the initial period of growth arrest. These effects of — increased glycolysis and lipogenesis, with suppression of mitochondrial fatty acid oxidation — are also among the most characteristic and important consequences of insulin signaling. Given this, the fact that actually interferes with insulin signaling and induces insulin resistance suggests that may be acting as a substitute for insulin. This unusual regulatory interaction may serve to take advantage of what may be the most significant and profound difference between the two agents in this context: that is, in contrast to insulin’s suppressive effect on adipocyte lipolysis and FFA mobilization, potently activates these processes. In so doing, promotes the implementation of two fundamental and obligatory components of cell proliferation: 1) similar to insulin, a supportive intracellular metabolic and energetic milieu; and 2) in contrast to insulin, an augmented availability of adipocyte-derived essential and other fatty acids required for membrane and eicosanoid biogenesis. The fact that and other proinflammatory cytokines have salutary effects on hepatocyte metabolism, energetics, and growth potential, despite the ability of to disrupt mitochondrial electron transport and induce cell death107,126-129, implies that the cellular impact of these agents must be the resultant of several factors. These likely include a protective effect of cytokeratins 8 and18130,131, cytokine doseage116,132,133, modulation by various growth factors129 and/or NO72,133-136, effects of dietary fatty acids and possibly eicosanoids137,138, species/strain, age139, and in vivo vs. in vitro conditions. Of particular significance in this 140-142 context is evidence that activates . As discussed above (see Chapter 5.3.: Mitochondrial Function in Apoptosis), this presumably reflects signaling-induced activation of PI3K and Akt/PKB143, and/or non-lethal impairment of the electron transport chain with resulting generation of low level ROS144-146 . . in turn transcriptionally upregulates expression of genes important in redox regulation including iNOS147, COX2148, Bcl-2, Bcl-xL149 and the Bcl-2 homolog Bfl-1/A1150; it may also contribute to the expression of UCP2 in the regenerating post-hepatectomy liver151,152. These responses protect diverse cells and cell lines from

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cytotoxicity. Moreover, activation of participates in mediating the anti-apoptotic effect of insulin in CHO cells153 and of insulin-like growth factor-1 in neuronal cells154. also mediates sphingomyelinase/ceramide induction of iNOS, and thereby inhibition of caspases by NO in astrocytes, MCF-7 adenocarcinoma cells, and hepatocytes155-157. Indeed, iNOS is essential for normal liver regeneration after partial hepatectomy136, indicating the critical importance of its role in suppressing apoptosis158. Its diverse effects suggest that NO may function directly or indirectly to buffer redox changes, perhaps in part through modulation of the electron transport chain159. The critical dependence of the survival and proliferative arm of TNFR1 signaling on PI3K/Akt/PIB143 may render it vulnerable to inhibition by fatty acid excess, as is the case with insulin receptor signaling11-15,17-21,160 (also, see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). It is likely that such inhibitory interaction would be strongly manifest during the early hours after partial hepatectomy, when hepatocellular fatty acid abundance, fatty acid oxidation, and oxidative stress are prominent (see this Chapter 8.3.: Early Changes in Hepatocellular Metabolism and Gene Expression), and that this would contribute to the delayed initiation of cell cycle progression and replicative DNA synthesis that characterize this period. As a corollary, it may generally be the case that selection between alternative TNFR1 signaling pathways — i.e., survival- and growthpromoting vs. cell death-initiating161, reflects an important modulating influence of varying fatty acid-induced suppression of PI3K/AKT/PKB (see Chapter 5.3.: Mitochondrial Function in Apoptosis). As activation is an early consequence of both partial hepatectomy162 and, less consistently, peroxisome proliferator-induced mitogenesis163,164, its largely transcriptionally activated redox-related protective effect may account for the growth enhancement by that otherwise would be difficult to comprehend. Consistent with this formulation is evidence that inhibition of binding to DNA decreased mitosis and increased apoptosis during liver regeneration165. Not explainable by this interpretation, however, is the obervation that, while IL-6 administration corrected the impaired regenerative DNA synthesis in TNFR-1 knockout mice, it failed to increase binding to 104 DNA . This suggests that other factors, e.g., redox balance, or activity of Bcl-2 and/or Bcl-xL149,166,167, may have contributed to a putative protective effect. If so, such an effect would depend not only on abundance of the anti-apoptotic protein, but also on its phosphorylation state and/or its interaction with other moieties, possibly including other members of the Bcl-2 protein family 168 . The availability of diverse alternative protective


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mechanisms may account for the recent observation that hepatocytespecific inhibition of does not lead to apoptosis in posthepatectomy regenerating hepatocytes169.

8.4.2. Completion of Regeneration and Cell Cycle Exit After up to three rounds of mitosis, hepatocytes exit the cell cycle. There is no evidence that the return to a state of growth arrest is initiated by a specific mechanism or signal, and therefore may at least in part represent gradual normalization of those factors that initially led to cell proliferation. Given their large number, including changes in plasma levels of glucose, FFA, insulin and other hormones, growth factors, and cytokines, and in functions more specifically hepatocellular, normalization of any or all could be involved. Among them, however, evolving systemic and hepatocellular fatty acid metabolism may be dominant. In particular, whereas mitochondrial fatty acid oxidation increased initially, in response to dramatically increased FFA mobilization and in support of systemic requirements for augmented gluconeogenesis, the subsequent insulin-mediated suppression of FFA mobilization decreased hepatocellular fatty acid availability. As the antilipolytic effect of insulin diminishes availability to the hepatocyte of essential fatty acids stored in adipose tissue, it may contribute significantly to the antiproliferative effects of glucose administration on hepatic tumorigenesis in glycogen storage disease type I170,171 and early after partial hepatectomy28,29. The manner in which the overall regenerative process may be subject to modulation by an apparent interplay between insulin and awaits more complete characterization. Full understanding of these complex interactions will require further elucidation of the mechanism by which cell metabolism and energetics are transduced to effect cell cycle regulation. The significance of this evolutionarily conserved connection is exemplified in yeast by the SNF-1 gene172. It is strongly suggested in mammalian cells by the SNF-1 homologue AMPK and wg/Wnt-like signaling cascade, by numerous ATP-dependent protein synthetic, phosphorylative, and degradative reactions required for cell cycle progression, e.g., those involving the cyclins, retinoblastoma protein, and the actin cytoskeleton173-177, and by the conservation of substrate and energy implicit in normal Mphase phosphorylation-induced inactivation of poly(A) polymerase and thus of protein synthesis178.

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8.5

REFERENCES

1. N. Fausto. Hepatic regeneration, in: Hepatology: a Textbook of Liver Disease, T. Boyer, ed., Saunders, Philadelphia (1996), pp. 32-58. 2. U. Junge, and W. Creutzfeldt, Hepatotrophic effects of pancreatic and gastrointestinal hormones in the rat in vivo and in vitro, Hepatotrophic Factors, CIBA Symp #53:269-283 (1978). 3. H. Leffert, K. Koch, T. Moran, and B. Rubalcava, Hormonal control of rat liver regeneration, Gastroenterology 76:1470-1482 (1979). 4. M. Kaibori, A. Kwon, M. Nakagawa, et al., Stimulation of liver regeneration and function after partial hepatectomy in cirrhotic rats by continuous infusion of recombinant human hepatocyte growth factor, J Hepatol 27:381-390 (1997). 5. T. Nakatani, K. Ozawa, M. Asano, M. Ukikusa, Y. Kamiyama, and T. Tobe, Differences in predominant energy substrate in relation to the resected hepatic mass in the phase immediately after hepatectomy, J Lab Clin Med 97:887-898 (1981). 6. T. Olivecrona, and G. Fex, Metabolism of plasma lipids in partially hepatectomized rats, Biochim Biophys Acta 202:259-268 (1970). 7. S. Pilkis, and D. Granner, Molecular physiology of the regulation of hepatic gluconeogenesis and glycolysis, Annu Rev Physiol 54:885-909 (1992). 8. O. Owen, A. Morgan, H. Kemp, J. Sullivan, M. Herrera, and G. Cahill, Jr, Brain metabolism during fasting, J Clin Invest 46:1589-1595 (1967). 9. H. Mulligan, and M. Tisdale, Metabolic substrate utilization by tumour and host tissues in cancer cachexia, Biochem J 277:321-326 (1991B). 10. L. Stryer, Biochemistry, W.H. Freeman and Co, New York (1995). 11. X. Chen, N. Iqbal, and G. Boden, The effects of free fatty acids on gluconeogenesis and glycogenolysis in normal subjects., J Clin Invest 103:365-372 (1999). 12. A. Dresner, D. Laurent, M. Marcucci, et al., Effects of free fatty acids on glucose transport and IRS-1-associated phosphatidylinositol 3-kinase activity., J Clin Invest 103:253-259 (1999). 13. M. E. Griffin, M. J. Marcucci, G. W. Cline, et al., Free fatty acid-induced insulin resistance is associated with activation of protein kinase C theta and alterations in the insulin signaling cascade, Diabetes 48:1270-4 (1999). 14. K. Cusi, K. Maezono, A. Osman, et al., Insulin resistance differentially affects the PI 3-kinase- and MAP kinase-mediated signaling in human muscle., J Clin Invest 105:311-320 (2000). 15. P. Arner, Insulin resistance in type 2 diabetes: role of fatty acids, Diabetes Metab Res Rev 18 Suppl 2:S5-9 (2002). 16. G. Boden, and G. I. Shulman, Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction, Eur J Clin Invest 32 Suppl 3:14-23 (2002). 17. Y. Kruszynska, D. Worrall, J. Ofrecio, J. Frias, G. Macaraeg, and J. Olefsky, Fatty acid-induced insulin resistance: decreased muscle PI3K activation but unchanged Akt phosphorylation, J Clin Endocrinol Metab 87:226-234 (2002). 18. T. K. Lam, H. Yoshii, C. A. Haber, et al., Free fatty acid-induced hepatic insulin resistance: a potential role for protein kinase C-delta, Am J Physiol Endocrinol Metab 283:E682-91 (2002). 19. J. McGarry, Banting lecture, 2001: dysregulation of fatty acid metabolism in the etiology of type 2 diabetes, Diabetes 51:7-18 (2002).


191

20. C. L. Soltys, L. Buchholz, M. Gandhi, A. S. Clanachan, K. Walsh, and J. R. Dyck, Phosphorylation of cardiac protein kinase B is regulated by palmitate, Am J Physiol Heart Circ Physiol 283:H1056-64 (2002). 21. C. Yu, Y. Chen, G. W. Cline, et al., Mechanism by which fatty acids inhibit insulin activation of insulin receptor substrate-1 (IRS-1)-associated phosphatidylinositol 3kinase activity in muscle, J Biol Chem 277:50230-50236 (2002). 22. O. Owen, P. Felig, A. Morgan, J. Wahren, and G. Cahill, Jr, Liver and kidney metabolism during prolonged starvation, J Clin Invest 48:574-583 (1969). 23. N. Katz, A. Brinkmann, and K. Jungermann, Compensatory increase of the gluconeogenic capacity of rat kidney after partial hepatectomy, Hoppe Seylers Z Physiol Chem 360:51-57 (1979). 24. H. J. Woerle, E. Popa, J. Dostou, S. Welle, J. Gerich, and C. Meyer, Exogenous insulin replacement in type 2 diabetes reverses excessive hepatic glucose release, but not excessive renal glucose release and impaired free fatty acid clearance, Metabolism 51:1494-500 (2002). 25. C. Morley, S. Kuku, A. Rubenstein, and J. Boyer, Serum hormone levels following partial hepatectomy in the rat, Biochem Biophys Res Commun 67:653-661 (1975). 26. S. Mittelman, G. Van Citters, E. Kirkman, and R. Bergman, Extreme insulin resistance of the central adipose depot in vivo, Diabetes 51:755-761 (2002). 27. T. Kawaguchi, K. Osatomi, H. Yamashita, T. Kabashima, and K. Uyeda, Mechanism for fatty acid “sparing” effect on glucose-induced transcription: regulation of carbohydrate-responsive element-binding protein by AMP-activated protein kinase, J Biol Chem 277:3829-3835 (2002). 28. J. Simek, V. Chmelar, J. Melka, J. Pazderka, and Z. Charvat, Influence of protracted infusion of glucose and insulin on the composition and regeneration activity of liver after partial hepatectomy in rats, Nature 213:910-911 (1967). 29. J. Ngala Kenda, B. De Hemptinne, and L. Lambotte, Role of metabolic overload in the initiation of DNA synthesis following partial hepatectomy in the rat, Eur Surg Res 16:294-302 (1984). 30. B. Mittal, and C. Kurup, Influence of clofibrate administration on the rate of synthesis of macromolecules in regeneration rat liver, Biochim Biophys Acta 609:475-482 (1980). 31. C. Wolfrum, C. Borrmann, T. Börchers, and F. Spener, Fatty acids and hypolipidemic drugs regulate peroxisome proliferator-activated receptors a- and g-mediated gene expression via liver fatty acid binding protein: A signaling path to the nucleus, Proc Natl Acad Sci USA 98:2323-2328 (2001). 32. R. Kaikaus, W. Chan, N. Lysenko, R. Ray, P. Ortiz de Montellano, and N. Bass, Induction of peroxisomal fatty acid beta-oxidation and liver fatty acid binding protein by peroxisome proliferators: Mediation via the cytochrome P-450 4A1 omegahydroxylase pathway, J Biol Chem 268:9592-9603 (1993A). 33. R. Kaikaus, Z. Sui, N. Lysenko, et al., Regulation of pathways of extramitochondrial fatty acid oxidation and liver fatty acid-binding protein by long-chain monocarboxylic fatty acids in hepatocytes: effect of inhibition of carnitine palmitoyltransferase I, J Biol Chem 268:26866-26871 (1993B). 34. J. Bassuk, P. Tsichlis, and S. Sorof, Liver fatty acid binding protein is the mitosisassociated polypeptide target of a carcinogen in rat hepatocytes., Proc Natl Acad Sci USA 84:7547-7551 (1987). 35. S. Sorof, Modulation of mitogenesis by liver fatty acid binding protein, Cancer Metast Rev 13:317-336 (1994).

192

Chapter 8

36. S. P. Anderson, L. Yoon, E. B. Richard, C. S. Dunn, R. C. Cattley, and J. C. Corton, Delayed liver regeneration in peroxisome proliferator-activated receptor-alpha-null mice, Hepatology 36:544-54 (2002). 37. K. Sasaki, T. Cripe, S. Koch, et al., Multihormonal regulation of phosphoenolpyruvate carboxykinase gene transcription: The dominant role of insulin, J Biol Chem 259:15242-15251 (1984). 38. J. Rosa, R. Bartrons, and A. Tauler, Gene expression of regulatory enzymes of glycolysis/gluconeogenesis in regenerating rat liver, Biochem J 287:113-116 (1992A). 39. J. Rosa, A. Tauler, A. Lange, S. Pilkis, and R. Bartrons, Transcriptional and posttranscriptional regulation of 6-phosphofructo-2-kinase/fructose-2,6bisphosphatase during liver regeneration, Proc Natl Acad Sci USA 89:3746-3750 (1992B). 40. R. Taub, Liver regeneration 4: transcriptional control of liver regeneration, FASEB J 10:413-427 (1996). 41. P. Schofield, A. Kerbey, and M. Sugden, Hepatic pyruvate metabolism during liver regeneration after partial hepatectomy in the rat, Intl J Biochem 18:453-458 (1986). 42. S. Monga, P. Pediaditakis, K. Mule, D. Stolz, and G. Michalopoulos, Changes in WNT/beta-catenin pathway during regulated growth in rat liver regeneration, Hepatol 33:1098-1109 (2001). 43. G. Asins, J. Rosa, D. Serra, et al., Gene expression of enzymes regulating ketogenesis and fatty acid metabolism in regenerating rat liver., Biochem J 299:6569 (1994). 44. E. Park, R. Mynatt, G. Cook, and K. Kashfi, Insulin regulates enzyme activity, malonyl-CoA sensitivity and mRNA abundance of hepatic carnitine palmitoyltransferase-I, Biochem J 310:853-858 (1995). 45. S. Mills, D. Foster, and J. Mcgarry, Interaction of malonyl-CoA and related compounds with mitochondria from different rat tissues: Relationship between ligand binding and inhibition of carnitine palmitoyltransferase I, Biochem J 214:83-91 (1983). 46. L. Drynan, P. Quant, and V. Zammit, Flux control exerted by mitochondrial outer membrane carnitine palmitoyltransferase over beta-oxidation, ketogenesis and tricarboxylic acid cycle activity in hepatocytes isolated from rats in different metabolic states., Biochem J 317:791-795 (1996A). 47. L. Drynan, P. Quant, and V. Zammit, The role of changes in the sensitivity of hepatic mitochondrial overt carnitine palmitoyltransferase in determining the onset of the ketosis of starvation in the rat., Biochem J 318:767-770 (1996B). 48. J. Sleboda, K. Risan, O. Spydevold, and J. Bremer, Short-term regulation of carnitine palmitoyltransferase I in cultured rat hepatocytes: spontaneous inactivation and reactivation by fatty acids, Biochim Biophys Acta 1436:541-549 (1999). 49. J. McGarry, and N. Brown, Reconstitution of purified, active and malonyl-CoAsensitive rat liver carnitine palmitoyltransferase I: relationship between membrane environment and malonyl-CoA sensitivity, Biochem J 349:179-187 (2000). 50. K. Ueda, T. Yoshioka, Y. Takehara, and K. Abe, Lipoperoxides, vitamin E, and activities of superoxide dismutase, glutathione peroxidase, and catalase in regenerating rat liver, Biochem Intl 7:663-669 (1983). 51. J.-L. Tsai, K.-L. King, C.-C. Chang, and Y.-H. Wei, Changes of mitochondrial respiratory functions and superoxide dismutase activity during liver regeneration, Biochem Intl 28:205-217 (1992).


193

52. P. Schofield, M. Sugden, C. Corstorphine, and V. Zamtnit, Altered interactions between lipogenesis and fatty acid oxidation in regenerating rat liver, Biochem J 241:469-474 (1987A). 53. P. Schofield, T. French, and M. Sugden, Ketone-body metabolism after surgical stress or partial hepatectomy: Evidence for decreased ketogenesis and a site of control distal to carnitine palmitoyltransferase I, Biochem J 241:475-481 (1987B). 54. V. Skulachev, Fatty acid circuit as a physiological mechanism of uncoupling of oxidative phosphorylation, FEBS Lett 294:158-162 (1991). 55. L. Wojtczak, and P. Schönfeld, Effect of fatty acids on energy coupling processes in mitochondria, Biochim Biophys Acta 1183:41-57 (1993). 56. L. Wojtczak, and M. Wieckowski, The mechanisms of fatty acid-induced proton permeability of the inner mitochondrial membrane, J Bioenerg Biomembranes 31:447-455 (1999A). 57. P. Schönfeld, M. Wiêckowski, and L. Wojtczak, Thyroid hormone-induced expression of the ADP/ATP carrier and its effect on fatty acid-induced uncoupling of oxidative phosphorylation, FEBS Lett 416:19-22 (1997A). 58. P. Schönfeld, and R. Bohnensack, Fatty acid-promoted mitochondrial permeability transition by membrane depolarization and binding to the ADP/ATP carrier, FEBS Lett 420:167-170 (1997B). 59. M. Buckle, Regulation of ATP hydrolase activity of the F0-F1 complex of rat-liver mitochrondria during early hepatic regeneration., FEBS Letters 209:197-202 (1986). 60. G. Vendemiale, F. Guerrieri, I. Grattagliano, D. Didonna, L. Muolo, and E. Altomare, Mitochondrial oxidative phosphorylation and intracellular glutathione compartmentation during rat liver regeneration, Hepatology 21:1450-1454 (1995). 61. F. Guerrieri, L. Muolo, T. Cocco, et al., Correlation between rat liver regeneration and mitochondrial energy metabolism, Biochim Biophys Acta 1272:95-100 (1995). 62. K. Ozawa, T. Yamada, M. Ukikusa, et al., Mitochondrial phosphorylative activity and DNA synthesis in regenerating liver of diabetic rats, J Surg Res 31:38-45 (1981). 63. T. Morimoto, Y. Taki, A. Jikko, et al., Changes in oxidative phosphorylation, adenylate energy charge, and respiratory components in chloramphenicol-treated regenerating rat liver, J Lab Clin Med 107:194-198 (1986). 64. T. Inomoto, A. Tanaka, S. Mori, et al., Changes in the distribution of the control of the mitochondrial oxidative phosphorylation in regenerating rabbit liver, Biochim Biophys Acta 1188:311-317 (1994). 65. G. Formiggini, C. Castelluccio, M. Pich, et al., Coenzyme Q depletion in rat plasma after partial hepatectomy., Biochem Molec Biol Intl 39:1135-1140 (1996). 66. P. Forsmark-Andrée, C.-P. Lee, G. Dallner, and L. Ernster, Lipid peroxidation and changes in the ubiquinone content and the respiratory chain enzymes of submitochondrial particles., Free Radical Biol Med 22:391-400 (1997). 67. I. Reynolds, and T. Hastings, Glutamate induces the production of reactive oxygen species in cultured forebrain neurons following NMDA receptor activation, J Neurosci 15:3318-3327 (1995). 68. V. Skulachev, Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants, Quart Rev Biophys 29:169-202 (1996). 69. A. Nègre-Salvayre, C. Hirtz, G. Carrera, et al., A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation, FASEB J 11:809-815 (1997).

194

Chapter 8

70. A. Stout, H. Raphael, B. Kantercwicz, E. Klann, and I. Reynolds, Glutamate-induced neuron death requires mitochondrial calcium uptake, Nature Neurosci 1:366-373 (1998). 71. S. Korshunov, O. Kokina, E. Ruuge, V. Skulachev, and A. Starkov, Fatty acids as natural uncouplers preventing generation of - and by mitochondria in the resting state, FEBS Letts 435:215-218 (1998). 72. K. Decker, and M. Obolenskaya, Cytokines, nitric oxide synthesis and liver regeneration., J Gastroenterol Hepatol 10:S12-S17 (1995). 73. K. Yamamoto, and H. Fahimi, Biogenesis of peroxisomes in regenerating rat liver, I: Sequential changes of catalase and urate oxidase detected by ultrastructural cytochemisty, Eur J Cell Biol 43:293-300 (1987). 74. H. Hayashi, H. Yoshida, F. Hashimoto, and S. Okazeri, Changes in polyamineoxidizing capacity of peroxisomes under various physiological conditions in rats, Biochim Biophys Acta 991:310-316 (1989). 75. G. Lüers, K. Beier, T. Hashimoto, H. Fahimi, and A. Völkl, Biogenesis of peroxisomes: sequential biosynthesis of the membrane and matrix proteins in the course of hepatic regeneration, Eur J Cell Biol 52:175-184 (1990). 76. I. Oikawa, and P. Novikoff, Catalase-negative peroxisomes: transient appearance in rat hepatocytes during liver regeneration after partial hepatectomy, Am J Pathol 146:673-687 (1995). 77. T. Delahunty, and D. Rubinstein, Accumulation and release of triglycerides by rat liver following partial hepatectomy., J Lipid Res 11:536-543 (1970). 78. J. Caruana, C. Sunby, D. Camara, H. Chen, and G. Schneeberger, Rapid alterations in substrate profiles after partial hepatectomy in the rat., Arch Surg 119:437-441 (1984). 79. K. Mohn, T. Laz, A. Melby, and R. Taub, Immediate-early gene expression differs between regenerating liver, insulin-stimulated H-35 cells, and mitogen-stimulated Balb/c 3T3 cells: Liver-specific induction patterns of gene 33, phosphoenolpyruvate carboxykinase, and the jun, fos, and egr families, J Biol Chem 265:21914-21921 (1990). 80. M. Nagino, M. Tanaka, M. Nishikimi, et al., Stimulated rat liver mitochondrial biogenesis after partial hepatectomy, Cancer Res 49:4913-4918 (1989). 81. Z.-Z. Huang, H. Li, J. Cai, J. Kuhlenkamp, N. Kaplowitz, and S. Lu, Changed in glutathione homeostasis during liver regeneration in the rat, Hepatology 27:147-153 (1998). 82. J. Dypbukt, M. Ankarcrona, M. Burkitt, et al., Different prooxidant levels stimulate growth, trigger apoptosis, or produce necrosis of insulin-secreting RINm5F cells: The role of intracellular polyamines., J Biol Chem 269:30553-30560 (1994). 83. V. Nilakantan, B. Spear, and H. Glauert, Liver-specific catalase expression in transgenic mice inhibits NF-kappaB activation and DNA synthesis induced by the peroxisome proliferator ciprofibrate, Carcinogenesis 19:631-637 (1998). 84. T. Finkel, Redox-dependent signal transduction., FEBS Letters 476:52-54 (2000A). 85. S. Nemoto, and T. Finkel, Redox regulation of forkhead proteins through a p66shcdependent signaling pathway, Science 295:2450-2452 (2002). 86. V. Thannickal, R. Day, S. Klinz, M. Bastien, J. Larios, and B. Fanburg, Rasdependent and -independent regulation of reactive oxygen species by mitogenic growth factors and TGF-b1, FASEB J 14:1741-1748 (2000). 87. H. Rabes, R. Wirsching, H.-V. Tuczek, and G. Iseler, Analysis of cell cycle compartments of hepatocytes after partial hepatectomy, Cell Tissue Kinet 9:517-532 (1976).


195

88. N. Barrass, R. Price, B. Lake, and T. Orton, Comparison of the acute and chronic mitogenic effects of the peroxisome proliferators methylclofenapate and clofibiric acid in rat liver., Carcinogenesis 14:1451-1456 (1993). 89. T. Nishihira, J. Tanaka, K. Nishikawa, et al., Biological significance of enhanced mitochondrial membrane potential in regenerating liver, Hepatology 6:220-224 (1986). 90. M. Kataoka, A. Tanaka, Y. Yamaoka, et al., A role of cytoplasmic free adenosine diphosphate in regenerating rabbit liver, J Lab Clin Med 119:354-358 (1992). 91. M. Kataoka, A. Tanaka, I. Ikai, Y. Yamaoka, K. Ozawa, and B. Chance, Kinetic analysis of ATP-synthetase and ATPase in regenerating rabbit liver, Eur Surg Res 25:91-97 (1993). 92. S. Yoshioka, M. Miyazaki, H. Shimizu, et al., Hepatic venous hemoglobin oxygen saturation predicts regenerative status of remnant liver after partial hepatectomy in rats, Hepatol 27:1349-1353 (1998). 93. S. Satoh, A. Tanaka, E. Hatano, et al., Energy metabolism and regeneration in transgenic mouse liver expressing creatine kinase after major hepatectomy, Gastroenterology 110:1166-1174 (1996). 94. M. Selzner, and P. Clavien, Failure of regeneration of the steatotic rat liver: disruption at two different levels in the regeneration pathway, Hepatol 31:35-42 (2000). 95. R. Cornell, B. Liljequist, and K. Bartizal, Depressed liver regeneration after partial hepatectomy of germ-free, athymic and lipopolysaccharide resistant mice., Hepatology 11:916-922 (1990). 96. A. Diehl, M. Yin, J. Fleckenstein, et al., Tumor necrosis factor-alpha induces c-jun during the regenerative response to liver injury., Am J Physiol 267:G552-G561 (1994). 97. Z. Sun, A. S. Klein, S. Radaeva, et al., In vitro interleukin-6 treatment prevents mortality associated with fatty liver transplants in rats, Gastroenterology 125:20215 (2003). 98. N. Selzner, M. Selzner, B. Odermatt, Y. Tian, N. Van Rooijen, and P. A. Clavien, ICAM-1 triggers liver regeneration through leukocyte recruitment and Kupffer celldependent release of TNF-alpha/IL-6 in mice, Gastroenterology 124:692-700 (2003). 99. K. Feingold, M. Soued, and C. Grunfeld, Tumor necrosis factor stimulates DNA synthesis in the liver of intact rats., Biochem Biophys Res Commun 153:576-582 (1988). 100. S. Kuma, M. Inaba, H. Ogata, et al., Effect of human recombinant interleukin-6 on the proliferation of mouse hepatocytes in the primary culture, Immunobiology 180:235-242 (1990). 101. P. Akerman, P. Cote, S. Yang, et al., Antibodies to tumor necrosis factor-alpha inhibit liver regeneration after partial hepatectomy., Am J Physiol 263:G579-G585 (1992). 102. M. Rolfe, N. James, and R. Roberts, Tumour necrosis factor alpha (TNF alpha) suppresses apoptosis and induces DNA synthesis in rodent hepatocytes: A mediator of the hepatocarcinogenicity of peroxisome proliferators?, Carcinogenesis 18:22772280 (1997). 103. D. Cressman, L. Greenbaum, R. Deangelis, et al., Liver failure and defective hepatocyte regeneration in interleukin-6-deficient mice., Science 274:1379-1383 (1996).

196

Chapter 8

104. Y. Yamada, I. Kinillova, J. Reschou, and N. Fausto, Initiation of tumor growth by tumor necrosis factor: deficient liver regeneration in mice lacking type I tumor necrosis factor receptor, Proc Natl Acad Sci USA 94:1441-1446 (1997). 105. B. Beutler, TNF, immunity and inflammatory disease: lessons of the past decade., J Investig Med 43:227-235 (1995). 106. F. Bazzoni, and B. Beutler, The tumor necrosis factor ligand and receptor families., N Engl J Med 334:1717-1725 (1996). 107. K. Schulze-Osthoff, A. Bakker, B. Vanhaesebroeck, R. Beyaert, W. Jacob, and W. Fiers, Cytotoxic activity of tumor necrosis factor is mediated by early damage of mitochondrial functions: Evidence for the involvement of mitochondrial radical generation, J Biol Chem 267:5317-5323 (1992). 108. M. Navasa, K. Feingold, and C. Grunfeld, Effects of endotoxin and cytokines on hepatic lipid metabolism, Prog Liver Dis 15:147-170 (1997). 109. G. Hotamisligil, Mechanisms of TNF-alpha-induced insulin resistance, Exp Clin Endocrinol Diabetes 107:119-125 (1999). 110. G. Hotamisligil, R. Johnson, R. Distel, R. Ellis, V. Papaioannou, and B. Spiegelman, Uncoupling of obesity from insulin resistance through a targeted mutation in aP2, the adipocyte fatty acid binding protein, Science 274:1377-1379 (1996). 111. K. Feingold, I. Staprans, R. Memon, et al., Endotoxin rapidly induces changes in lipid metabolism that produce hypertriglyceridemia: low doses stimulate hepatic triglyceride production while high doses inhibit clearance., J Lipid Res 33:17651776 (1992). 112. R. Memon, K. Feingold, A. Moser, et al., Differential effects of interleukin-1 and tumor necrosis factor on ketogenesis, Am J Physiol 263:E301-E309 (1992). 113. C. Grunfeld, C. Dinarello, and K. Feingold, Tumor necrosis factor-alpha, interleukin-1, and interferon alpha stimulate triglyceride synthesis in HepG2 cells, Metabolism 40:894-898 (1991). 114. E. Vara, J. Arias-Diaz, J. Torres-Melero, C. Garcia, J. Rodriguez, and J. Balibrea, Effect of different sepsis-related cytokines on lipid synthesis by isolated hepatocytes, Hepatology 20:924-931 (1994). 115. R. Memon, N. Bass, A. Moser, et al., Down-regulation of liver and heart specific fatty acid binding proteins by endotoxin and cytokines in vivo, Biochim Biophys Acta 1440:118-126 (1999). 116. S. Mahony, and M. Tisdale, Metabolic effects of tumour necrosis factor alpha in NMRI mice, Br J Cancer 61:514-519 (1990). 117. M. Beylot, H. Vidal, G. Mithieux, M. Odeon, and C. Martin, Inhibition of hepatic ketogenesis by tumor necrosis factor-alpha in rats., Am J Physiol 263:E897-E902 (1992). 118. M. Jin, Y. Shimahara, T. Yamaguchi, et al., The effect of a bolus injection of TNFalpha and IL-1 beta on hepatic energy metabolism in rats, J Surg Res 58:509-515 (1995). 119. R. Memon, J. Fuller, A. Moser, P. Smith, K. Feingold, and C. Grunfeld, In vivo regulation of acyl-CoA synthetase mRNA and activity by endotoxin and cytokines, Am J Physiol 275:E64-72 (1998). 120. K. Beier, A. Völkl, and H. Fahimi, Suppression of peroxisomal lipid beta-oxidation enzymes of TNF-alpha., FEBS Letters 310:273-276 (1992). 121. K. Beier, A. Völkl, and H. Fahimi, TNF-alpha downregulates the peroxisome proliferator activated receptor-alpha and the mRNAs encoding peroxisomal proteins in rat liver., FEBS Letters 12:385-387 (1997).


197

122. S. Ghosh, S. J, and B. R, Lipid biochemistry: functions of glycerolipids and sphingolipids in cellular signaling, FASEB J 11: 45-50 (1997). 123. G. Velasco, D. Gómez, Pulgar, T, D. Carlin, and M. Guzmán, Evidence that the AMPactivated protein kinase stimulates rat liver carnitine palmitoyltransferase I by phosphorylating cytoskeletal components, FEBS Lett 439:317-320 (1998A). 124. G. Velasco, M. Geelen, T. Gomez del Pulgar, and M. Guzmán, Malonyl-CoAindependent acute control of hepatic carnitine palmitoyltransferase I activity: Role of Ca2+/calmodulin-dependent protein kinase II and cytoskeletal components, J Biol Chem 273:21497-21504 (1998B). 125. R. Warren, H. Starnes, Jr, N. Alcock, S. Calvano, and M. Brennan, Hormonal and metabolic response to recombinant human tumor necrosis factor in rat: in vitro and in vivo, Am J Physiol 255:E206-E212 (1988). 126. E. Tredget, Y. Yu, S. Zhong, et al., Role of interleukin 1 and tumor necrosis factor on energy metabolism in rabbits, Am J Physiol 255:E760-E768 (1988). 127. T. Hennet, C. Richter, and E. Peterhans, Tumour necrosis factor-alpha induces superoxide anion generation in mitochondria of L929 cells, Biochem J 289:587-592 (1993B). 128. M. Boermeester, I. Straatsburg, A. Houdijk, et al., Endotoxin and interleukin-1 related hepatic inflammatory response promotes liver failure after partial hepatectomy., Hepatology 22:1499-1506 (1995). 129. P. Martin-Sanz, M. Diaz-Guerra, M. Casado, and L. Bosca, Bacterial lipopolysaccharide antagonizes transforming growth factor beta 1-induced apoptosis in primary cultures of hepatocytes, Hepatology 23:1200-1207 (1996). 130. C. Caulin, C. Ware, T. Magin, and R. Oshima, Keratin-dependent, epithelial resistance to tumor necrosis factor-induced apoptosis., J Cell Biol 149:17-22. (2000). 131. S. Gilbert, A. Loranger, N. Daigle, and N. Marceau, Simple epithelium keratins 8 and 18 provide resistance to Fas-mediated apoptosis: The protection occurs through a receptor-targeting modulation, J Cell Biol 154:763-73 (2001). 132. P. Wang, A. Ayala, Z. Ba, M. Zhou, M. Perrin, and I. Chaudry, Tumor necrosis factor-alpha produces hepatocellular dysfunction despite normal cardiac output and hepatic microcirculation, Am J Physiol 265:G126-G132 (1993). 133. C. Fisch, M. Robin, P. Letteron, et al., Cell-generated nitric oxide inactivates rat hepatocyte mitochondria in vitro but reacts with hemoglobin in vivo., Gastroenterology 110:210-220 (1996). 134. D. Duval, D. Miller, J. Collier, and R. Billings, Characterization of hepatic nitric oxide synthase: identification as the cytokine-inducible form primarily regulated by oxidants., Molec Pharmacol 50:277-284 (1996). 135. H. Kitade, T. Kanemaki, K. Sakitani, et al., Regulation of energy metabolism by interleukin-1 beta, /but not by interleukin-6, is mediated by nitric oxide in primary cultured rat hepatocytes, Biochim Biophys Acta 1311:20-26 (1996). 136. R. Rai, F. Lee, A. Rosen, et al., Impaired liver regeneration in inducible nitric oxide synthase-deficient mice, Proc Natl Acad Sci USA 95:13829-13834 (1998). 137. R. Grimble, and P. Tappia, Modulatory influence of unsaturated fatty acids on the biology of tumour necrosis factor-alpha, Biochem Soc Trans 23:282-287 (1995). 138. P. Sierra, P. Ling, N. Istfan, and B. Bistrian, Fish oil feeding improves muscle glucose uptake in tumor necrosis factor-treated rats, Metabolism 44:1365-1370 (1995).

198

Chapter 8

139. C. Mondon, and H. Starnes, Jr, Differential metabolic responses to tumor necrosis factor with increase in age, Metabolism 41:970-981 (1992). 140. A. Beg, and D. Baltimore, An essential role for NF-kappaB in preventing TNFalpha-induced cell death., Science 274:782-784 (1996). 141. C.-Y. Wang, M. Mayo, and J. Baldwin, AS TNF- and cancer therapy-induced apoptosis: potentiation by inhibition of NF-kappaB, Science 274:784-787 (1996). 142. D. Van Antwerp, S. Martin, T. Kafri, D. Green, and I. Verma, Suppression of TNFalpha-induced apoptosis by NF-kappaB, Science 274:787-789 (1996). 143. O. Ozes, L. Mayo, J. Gustin, S. Pfeffer, L. Pfeffer, and D. Donner, NF-kappaB activation by tumour necrosis factor requires the Akt serine-threonine kinase. Nature 401:82-85 (1999). 144. C. Sen, and L. Packer, Antioxidant and redox regulation of gene transcription, FASEB J 10:709-720 (1996). 145. V. Lakshminarayanan, E. Drab-Weiss, and K. Roebuck, and tumor necrosis factor-alpha induce differential binding of the redox-responsive transcription factors AP-1 and NF-kappaB to the interleukin-8 promoter in endothelial and epithelial cells, J Biol Chem 273:32670-32678 (1998). 146. E. Shaulian, and M. Karin, AP-1 as a regulator of cell life and death, Nat Cell Biol 4:E131-6 (2002). 147. T. Vos, H. Van Goor, L. Tuyt, et al., Expression of inducible nitric oxide synthase in endotoxemic rat hepatocytes is dependent on the cellular glutathione status, Hepatology 29:421-426 (1999). 148. K. Yamamoto, T. Arakawa, N. Ueda, and S. Yamamoto, Transcriptional roles of nuclear factor kappa B and nuclear factor-interleukin-6 in the tumor necrosis factor alpha-dependent induction of cyclooxygenase-2 in MC3T3-E1 cells, J Biol Chem 270:31315-31320 (1995). 149. M. Tamatani, Y. Che, H. Matsuzaki, et al., Tumor necrosis factor induces Bcl-2 and Bcl-x expression through NFkappaB activation in primary hippocampal neurons, J Biol Chem 274:8531-8538 (1999). 150. W. Zong, L. Edelstein, C. Chen, J. Bash, and C. Gélinas, The prosurvival Bcl-2 homolog Bfl-2/A1 is a direct transcriptional target of NF-kappaB that blocks TNFalpha-induced apoptosis, Genes Development 13:382-387 (1999). 151. F. Lee, Y. Li, H. Zhu, et al., Tumor necrosis factor increases mitochondrial oxidant production and induces expression of uncoupling protein-2 in the regenerating mouse liver, Hepatology 29:677-687 (1999). 152. K. Echtay, D. Roussel, J. St-Pierre, et al., Superoxide activates mitochondrial uncoupling proteins., Nature 415:96-99 (2002). 153. F. Bertrand, A. Atfi, A. Cadoret, et al., A role for nuclear factor kappa B in the antiapoptotic function of insulin., J Biol Chem 273:2931-2938 (1998). 154. S. Heck, F. Lezoualc’h, S. Engert, and C. Behl, Insulin-like growth factor-1-mediated neuroprotection against oxidative stress is associated with activation of nuclear factor kappaB, J Biol Chem 274:9828-9835 (1999). 155. K. Pahan, F. Sheikh, M. Khan, A. Namboodiri, and I. Singh, Sphingomyelinase and ceramide stimulate the expression of inducible nitric-oxide synthase in rat primary astrocytes, J Biol Chem 273:2591-2600 (1998). 156. Y. Kim, R. Talanian, and T. Billiar, Nitric oxide inhibits apoptosis by preventing increases in caspase-3-like activity via two distinct mechanisms, J Biol Chem 272:31138-31148 (1997).


199

157. Y. Kim, T. Kim, D. Seol, R. Talanian, and T. Billiar, Nitric oxide suppression of apoptosis occurs in association with an inhibition of Bcl-2 cleavage and cytochrome c release, J Biol Chem 273:31437-31441 (1998). 158. J. Li, and T. Billiar, Nitric Oxide, IV: Determinants of nitric oxide protection and toxicity in liver, Am J Physiol 276:G1069-G1073 (1999). 159. S. Moncada, and J. Erusalimsky, Does nitric oxide modulate mitochondrial energy generation and apoptosis?, Nat Rev Mol Cell Biol 3:214-220 (2002). 160. H. Steinberg, G. Paradisi, G. Hook, K. Crowder, J. Cronin, and A. Baron, Free fatty acid elevation impairs insulin-mediated vasodilation and nitric oxide production, Diabetes 49:1231-1238 (2000). 161. O. Micheau, and J. Tschopp, Induction of TNF receptor I-mediated apoptosis via two sequential signaling complexes, Cell 114:181-90 (2003). 162. N. Fausto, A. Laird, and E. Webber, Liver regeneration, 2: Role of growth factors and cytokines in hepatic regeneration., FASEB J 9:1527-1536 (1995). 163. Y. Li, L. Leung, H. Glauert, and B. Spear, Treatment of rats with the peroxisome proliferator ciprofibrate results in increased liver NF-kappaB activity, Carcinogenesis 17:2305-2309 (1996). 164. M. Menegazzi, P. Carcereri-De, A, H. Suzuki, et al., Liver cell proliferation induced by nafenopin and cyproterone acetate is not associated with increases in activation of transcription factors NF-kappaB and AP-1 or with expression of tumor necrosis factor alpha, Hepatology 25:585-592 (1997). 165. Y. Iimuro, T. Nishiura, C. Hellerbrand, et al., NF kappa B prevents apoptosis and liver dysfunction during liver regeneration, J Clin Invest 101:802-811 (1998). 166. B. Kren, J. Trembley, S. Krajewski, T. Behrens, J. Reed, and C. Steer, Modulation of apoptosis-associated genes bcl-2, bcl-x, and bax during rat liver regeneration, Cell Growth Differentiation 7:1633-1642 (1996). 167. S. Tzung, N. Fausto, and D. Hockenbery, Expression of Bcl-2 family during liver regeneration and identification of Bcl-x as a delayed early response gene, Am J Pathol 150:1985-1995 (1997). 168. N. Zamzami, C. Brenner, I. Marzo, S. Susin, and G. Kroemer, Subcellular and submitochondrial mode of action of Bcl-2-like oncoproteins, Oncogene 16:22652282 (1998). 169. M. L. Chaisson, J. T. Brooling, W. Ladiges, S. Tsai, and N. Fausto, Hepatocytespecific inhibition of NF-kappaB leads to apoptosis after TNF treatment, but not after partial hepatectomy, J Clin Invest 110:193-202 (2002). 170. P. Parker, I. Burr, A. Slonim, F. Ghishan, and H. Greene, Regression of hepatic adenomas in type Ia glycogen storage disease with dietary therapy, Gastroenterology 81:534-536 (1981). 171. R. Ockner, R. Kaikaus, and N. Bass, Fatty acid metabolism and the pathogenesis of hepatocellular carcinoma: review and hypothesis, Hepatology 18:669-676 (1993). 172. D. Hardie, D. Carling, and M. Carlson, The AMP-activated/SNF1 protein kinase subfamily: metabolic sensors of the eukaryotic cell?, Annu Rev Biochem 67:821-855 (1998). 173. S. Sweet, and G. Singh, Accumulation of human promyelocytic leukemic (HL-60) cells at two energetic cell cycle checkpoints, Cancer Res 55:5164-5167 (1995). 174. R. Sheaff, M. Groudine, M. Gordon, J. Roberts, and B. Clurman, Cyclin E-CDK2 is a regulator of p27Kipl, Genes Devel 11:1464-1478 (1997).

200

Chapter 8

175. M. Pagano, S. Tam, A. Theodoras, et al., Role of the ubiquitin-proteasome pathway in regulating abundance of the cyclin-dependent kinase inhibitor p27, Science 269:682-685 (1995). 176. W. Kaufmann, and T. Goldsworthy, Regulation of the hepatocyte cell cycle, Prog Liver Dis 15:31-56 (1997). 177. A. Hall, Rho GTPases and the actin cytoskeleton, Science 279:509-514 (1998). 178. D. Colgan, K. Murthy, C. Prives, and J. Manley, Cell-cycle related regulation of poly(A)polymerase by phosphorylation., Nature 384:282-285 (1996).

Chapter 9 Part II Conclusions

The foregoing analysis supports a model in which regulation of cell growth, injury, and death are linked to cell metabolism. Five principles emerge as central elements of the model: 1. Cell cycle progression and mitosis require dramatic increases in the availability of specific nutrients and in the generation of chemical energy (ATP) to execute obligatory biosynthetic, translocational, regulatory, and signaling processes. 2. Substrate and energetic requirements of aerobic cell proliferation are met predominantly by glycolysis, the pentose phosphate cycle, and the citric acid cycle. These sustain the increases in and thus augmented rates of oxidative phosphorylation (mitochondrial ATP biosynthesis), that are characteristic of most proliferating cells. Glucose also provides carbon atoms and cofactors such as reduced pyridine nucleotides that are required for maintenance of redox balance and for essential biosynthetic reactions, including those for fatty acids, isoprenoids, and sterols. Important exceptions, which accomplish the same overall objectives, include glutamine-derived 2-oxo-glutarate and, under hypoxic conditions, generation of ATP via anaerobic glycolysis. 3. Fatty acids are potentially usable as fuel. However, their increased tissue abundance and mitochondrial oxidation inhibit glucose uptake, glycolysis, pyruvate dehydrogenase, and glycolytic fueling of the citric acid cycle, and predispose to ROS generation and oxidative stress. Long chain fatty acids may also uncouple oxidative phosphorylation, thereby compromising and the efficiency of ATP biosynthesis. Moreover, fatty acids are required in proliferating cells for membrane and eicosanoid biogenesis. Consistent with this, biosynthesis of fatty acids is coordinately upregulated with that of isoprenoids and sterols at both transcriptional and post-transcriptional levels. As a corollary, 201

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extravagances such as augmented fatty acid oxidation in mitosis would seem unlikely to have survived as evolutionarily advantageous, given the historical unpredictability of nutrient availability, the highly efficient energy burst required for rapid transit through the later stages of the mitotic cycle, and the mitosis-associated increases in cellular fatty acid requirement. Instead, mechanisms have evolved that inhibit mitochondrial fatty acid oxidation in dividing cells, thereby both increasing the oxidation of glucose and the efficiency of energy generation, while conserving needed fatty acids. Accordingly, fatty acid utilization in cells undergoing mitosis appears to be principally anabolic, i.e., for biogenesis of eicosanoids and the lipid bilayers that define the very existence of daughter cells and organelles. 4. Consistent with the foregoing principles, factors that promote growth, e.g., peroxisome proliferators, insulin, and proinflammatory cytokines, limit mitochondrial oxidation of fatty acids relative to mitochondrial oxidative capacity while increasing glycolysis and — under normoxic conditions — and oxidative phosphorylation. Growth promotion by fatty acids appears to reflect early generation of eicosanoids, cell membranes, and limited, growth-supporting quantities of ROS. Conversely, factors that arrest growth or induce apoptosis, e.g., butyrate, salicylate, and fatty acids, relatively or absolutely enhance mitochondrial fatty acid oxidation, thereby inhibiting glycolysis, compromising and the efficiency of oxidative phosphorylation, and potentially generating excessive ROS and oxidative stress. The model thus implies a cellular compartmentalization of fatty acids in which their relative or absolute utilization as mitochondrial oxidative substrates is inversely related to cell proliferation. Available evidence suggests that the survival and proliferative arm of this overall regulatory mechanism depends importantly on wingless/Wnt-like PI3K/Akt/PKB signaling and its diverse cellular interactions, including the multicomponent VDAC complex at the contact sites between inner and outer mitochondrial membranes. In the antiproliferative arm, in contrast, the AMP-activated protein kinase appears to play a significant role. As discussed, aspects of this general formulation would not apply to glycolysis-fueled hypoxic proliferation in certain cell populations (in which suppressed fatty acid oxidation is implicit), nor to normal colonic epithelium in which cell metabolism and growth regulation are uniquely influenced by interaction with resident bacterial flora and a short-chain fatty acid-enriched milieu. 5. It is clear that increased availability of substrates and cofactors alone is not sufficient to promote cell proliferation. Thus, when fatty acids are present in excess within the cell — e.g., resulting from the elevated plasma FFA associated with fasting, obesity, insulin resistance,

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diabetes mellitus, fatty liver, HIV infection, or nicotine exposure — they may override the normally limiting effect of malonyl CoA on CPT-I activity. As a result, mitochondrial of fatty acids is accelerated, inhibiting glucose uptake and glycolysis, attenuating and predisposing to ROS generation, thereby potentially fostering genomic instability and carcinogenesis. Acting via differing mechanisms, fatty acids, salicylate, and the “statin” hypolipidemic agents, suppress both cell proliferation and oxidative stress in the implementation of their protective effects. Trans fatty acids less readily undergo in mitochondria, compared with their cis analogs, and therefore are more likely to enter the peroxisomal pathway. The model implies a setting in which ordinary energetic demands, and especially those of the cell cycle, are imposed upon populations of cells that are inevitably heterogeneous in regard to their capacities to generate energy, constrain or neutralize oxidative stress, and adapt to microenvironments. Together, these elements comprise a microcosm in which natural selection would favor cell cycle completion and lineage survival of the“fittest” genotypes and/or epigenetically-regulated gene expression patterns. Under certain conditions, interaction among these elements may contribute importantly to tumorigenesis. The model is supported by a wide variety of experimental observations, including studies of mitochondrial energetics, intermediary metabolism, and gene expression in proliferating, growth arrested, and apoptotic cells. The model also suggests several corollaries that are appropriate issues for further investigation: 1) Failure of the cell to marshal the metabolic and energetic requirements for cell cycle progression will arrest growth, or may induce apoptosis despite (or precipitated by) activation of growth-promoting cell signaling pathways. 2) Conversely, a growth-promoting metabolic profile may be sufficient, per se, to initiate cell cycle progression. This occurs in yeast and other micro-organisms, and may also characterize early embryogenesis and the development of certain neoplasms, e.g., consequent to peroxisome proliferator administration in rodents and in human glycogen storage disease, type I. 3) The model may provide a theoretical basis for development of new methods to characterize the growth regulatory, tumorigenic, and antitumorigenic potential of endogenous and exogenous agents. 4) The model may permit alternative approaches to the understanding and control of cell growth in several clinical settings. These include: a) management of large hepatic resections, fulminant

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hepatic failure, and small-for-recipient-size liver transplantation; b) initial hyperplasia and subsequent programmed death of pancreatic islet in insulin-resistant states; c) elucidation of the relationship of carcinogenesis to fat and calorie intake, obesity, and nicotine; d) pathogenesis, prevention, and treatment of cancer; and e) regulation of hematopoiesis and of immunocyte population kinetics. 5) Integral to the model are selected concepts that may prove applicable to an understanding of other incompletely resolved basic and clinical questions. Thus, whereas transient and/or low grade generation of ROS may activate survival and growth responses, sustained and intensive fatty acid oxidation-induced intramitochondrial oxidative stress in specific cell populations may represent a fundamental element in the pathogenesis of diverse forms of cell and tissue injury. This dichotomous mechanism may be expecially important, e.g., in the regulation of placental, embryonic, and fetal development, the pathogenesis and mitigation of the endothelial lesion of atherosclerosis, nonalcoholic steatohepatitis, ischemia-reperfusion injury, aging, and, as discussed in Part III, neurodegenerative disorders.

PART III: FATTY ACIDS, KETONE BODIES, AND BRAIN METABOLISM: IMPLICATIONS FOR NEURONAL FUNCTION AND ALZHEIMER DISEASE

Chapter 10 Introduction to Part III

In Part III, the influence of local and systemic metabolic factors on neuronal function and injury in brain is examined. Particular emphasis is placed on the critically important interactions between glucose utilization and the consequences of fatty acid oxidation, and between neurons and astrocytes. Energetically and metabolically, neuronal activation bears certain similarities to cell cycle progression in non-post-mitotic cells (see Chapter 3: Nutrient and Energy Metabolism in Cell Proliferation). Thus, neuronal activation and synaptic function require accelerated and highly efficient aerobic glycolysis-fueled mitochondrial ATP generation and wingless/Wnt-like (wg/Wnt-like) signaling. While it has long been recognized that glucose is essential for optimal brain function, it is also clear that full understanding of the role and regulation of glucose utilization in brain in health and disease requires consideration of other potential energy sources, and of the factors that govern their interactions. Particularly important in this regard are the ketone bodies, i.e., acetoacetate and While plasma ketone bodies may substitute for glucose as a source of brain energy during starvation, their utilization has certain unfavorable characteristics, as discussed below. Despite this, ketone bodies are utilized by brain in preference to glucose in both non-fasted and fasted subjects. As a corollary, utilization of glucose in brain is governed not only by neuronal activation and vascular glucose supply, but also by the availability of ketone bodies. The brain also takes up plasma free fatty acids (FFA), a significant fraction of which undergoes mitochondrial Except for those hypothalamic neurons involved in neuroendocrine and metabolic regulation, however, fatty acid oxidation in brain occurs primarily in astrocytes and other glial cells (see Chapter 12: Utilization of Oxidizable Substrates in Brain). Moreover, astrocytes resemble hepatocytes in that their 207

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mitochondrial oxidation of fatty acids may support the generation of ketone bodies, which are exported from the cell. In liver, newly synthesized ketone bodies enter hepatic sinusoidal plasma and the systemic circulation. In brain, in contrast, ketone bodies exported from astrocytes represent an alternative fuel that may be used directly, and in preference to glucose, by nearby neurons. As a consequence, glucose utilization and other aspects of metabolism and function in these neurons would be subject to modulation by seemingly extraneous influences, such as systemic and astrocytic fatty acid metabolism and ketogenesis. Based on these and other considerations to be discussed, it is hypothesized that ketone bodies synthesized both locally (in astrocytes) and systemically (in hepatocytes) may represent an important source of energy that supports brain function, not only under conditions of starvation, but also in other circumstances in which systemic and/or local ketogenesis is increased. Unfortunately, mitochondrial oxidation of ketone bodies is energetically less favorable than that of glucose in that they may exert an uncoupling-like effect on oxidative phosphorylation (ATP biosynthesis), and may predispose to the generation of ROS and therefore oxidative stress. Accordingly, in pre-empting neuronal glucose utiliztion by the neuron, ketone bodies may compromise neuronal energetic efficiency and, together with both endogenously generated and diffusible ROS exported by the fatty acid-oxidizing astrocyte, promote cell injury. This hypothesized phenomenon, reflecting increased fatty acid-driven astrocyte ketogenesis, may be especially significant in systemic disorders that are associated with increased plasma [FFA] such as obesity, insulin resistance, type II diabetes mellitus, and HIV infection. In the following analysis, physiological and clinical implications of neuron-astrocyte metabolic interaction, and other phenomena related to metabolic and energetic aspects of neuronal function, are critically examined. Issues addressed in this analysis include the interactions between systemic and hypothalamic fatty acid metabolism in regulation of feeding behavior, the apparent dissociation between glucose and oxygen utilization during neuronal activation, the antiseizure effect of ketogenic diets, the diverse effects of lithium, and the mechanisms involved in long term potentiation, excitotoxicity, and HIV dementia. Finally, it is hypothesized that in Alzheimer disease, regional suppression of neuronal glucose utilization may in part reflect adverse consequences of competition between glucose and astrocyte-generated ketone bodies. In a hypothetical model of this disorder, neuronal oxidative stress and injury induced by excessive fatty acid oxidation and ketogenesis in astrocytes, are integrated with genetic determinants, amyloid, and phosphorylation in a proposed “vicious cycle” of disease pathogenesis.

Chapter 11 Energetics of Neuronal Activation

11.1. INTRODUCTION - RELEVANT PRINCIPLES OF INTERMEDIARY METABOLISM A central theme throughout this analysis will be the importance of selective utilization of specific substrates and metabolic pathways in the neuronal generation of ATP. The following brief summary highlights aspects of intermediary metabolism that are particularly relevant in this regard. With minor changes appropriate to this neuronal context it recapitulates aspects of the summation provided in Chapter 3.2.: Intermediary Metabolism: General Considerations. Glucose oxidation requires phosphorylation by hexokinase and entry into the glycolytic pathway (Fig. 11.1A). Completion of glycolysis with formation of pyruvate generates a net of 2 molecules of ATP per molecule of glucose. Under certain conditions the process ends here: e.g., in myocytes during sustained excercise, pyruvate may substitute for oxygen as acceptor of the electrons generated by glycolysis, thereby forming lactate which is exported from the cell. This “anaerobic glycolysis” is far less efficient than complete oxidation of glucose (“aerobic glycolysis”), which depends on a stringent, pyruvate dehydrogenase- (PDH)-mediated regulation of the mitochondrial oxidation of pyruvate via the citric acid cycle (Fig. 11.1A and Fig. 3.2, facing p. 21). Aerobic glycolysis yields 30 molecules of ATP per molecule of glucose, almost all of which are generated by ATP synthasemediated oxidative phosphorylation; the yield from complete oxidation of two molecules of lactate is approximately the same. The rate of oxidative phosphorylation is driven by the mitochondrial inner membrane potential which is maintained principally by the export 209

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11. Energetics of Neuronal Activation

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Figure 11.1. Relevant aspects of intermediary metabolism. Pathway arrow thickness approximates relative activity; dotted lines: relatively or absolutely suppressed; enclosed areas represent mitochondria. A. Glycolysis- and citric acid cycle-fueled ATP synthesis. Complete oxidation of glucose via aerobic glycolysis, pyruvate dehydrogenase, and the citric acid cycle, yields 30 molecules of ATP per molecule of glucose. Gluconeogenesis is suppressed. B. Fatty acid oxidation. Mitochondrial oxidation of long chain fatty acids requires CPT-I-mediated formation of acylcarnitine. Fatty acids present in excess may override malonyl CoA inhibition of CPT-I, driving inappropriately augmented mitochondrial oxidation. The theoretical maximum yield of ATP molecules from complete oxidation of one molecule of palmitate (~100) may be diminished to as little as ~20 in a ketogenic cell (astrocyte or hepatocyte) by diversion of acetyl CoA formed through to ketogenesis rather than to the citric acid cycle. C.Gluconeogenesis. The process is normally fueled energetically by the mitochondrial oxidation of fatty acids. Glycolysis is suppressed. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; F26BP: fructose-2,6bisphosphate; NADPH: nucleotide adenine triphosphate, reduced form; PCB: pyruvate carboxylase; PEPCK: phosphoenolpyruvate carboxykinase; PL: glycerophosphatides; TG: triacylglycerols.

of protons across this membrane by complexes I, III, and IV of the electron transport chain (Fig. 3.2, facing p. 21). Fatty acids may also be oxidized in mitochondria to support ATP generation, e.g., in liver, muscle, and astrocytes, but not significantly in most neurons (important exceptions to be discussed; see Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior). In this process (Fig. 11.1B and Fig. 3.2, facing p. 21), a long chain fatty acid (e.g., the 16-carbon palmitate) is converted to its acyl CoA then acylcarnitine derivatives. The latter step, mediated by carnitine palmitoyltransferase-I (CPT-I), regulates and is required for long chain fatty acid entry into the mitochondrial matrix and the spiral; the latter provides electrons to the electron transport chain and generates eight 2-carbon acetyl CoAs. Acetyl CoA may then undergo complete oxidation to and via the citric acid cycle, or (in astrocyte or hepatocyte mitochondria) enter a pathway that leads to formation of 4carbon ketone bodies (acetoacetate and ) that are exported from the cell. A theoretical yield of 106 ATP for each completely oxidized palmitate molecule is thus diminished by ketogenesis, and further by the inefficiency resulting from uncoupling of oxidative phosphorylation (non-ATP synthase-mediated transfer of protons back across the inner membrane) that may be caused by underivatized long chain fatty acids and other agents. In addition, an uncoupling-like effect may be caused by certain substrates that undergo oxidation in the mitochondrial matrix, as will be discussed. Mitochondrial fatty acid oxidation also inhibits several steps in cellular glucose uptake and glycolysis (the “Randle cycle”)1-3, and in astrocytes,

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hepatocytes, and renal tubular epithelial cells fuels gluconeogenesis (Fig. 11.1C). Importantly, control of mitochondrial oxidation of long chain fatty acids is less stringent than that of pyruvate oxidation by PDH. As a result, at high concentrations they may override the CPT-I control mechanism4-8. Moreover, ketone bodies and medium/short chain fatty acids freely enter mitochondrial matrix oxidative pathways independent of CPT-I. Accordingly, higher concentrations of these substrates may drive increased rates of while suppressing pyruvate utilization and glycolysis by inhibiting PDH and glucose uptake. The resulting increase in electron transport chain flux may promote electron “leakage”, formation of reactive oxygen species (ROS), and oxidative stress. Thus, generation of ATP may be primarily glycolytic or primarily dependent on fatty acid oxidation (Fig. 11.1). The latter, less efficient and more likely to generate ROS, suppresses glucose uptake, PDH, and thus glycolysis; in astrocytes and hepatocytes, mitochondrial fatty acid oxidation supports gluconeogenesis and/or ketogenesis. In general, neurons oxidize glucose, lactate, ketone bodies, or citric acid cycle intermediates to generate ATP, but do not oxidize fatty acids (with exceptions to be discussed). Astrocytes are more versatile; not only may they oxidize fatty acids in support of ATP generation, but in addition may generate glucose de novo via gluconeogenesis, and may export lactate and/or ketone bodies. These principles are fundamental to an understanding of brain metabolism as related to the following discussion, and will be developed further as necessary.

11.2.

ENERGY-CONSUMING EVENTS IN NEURONAL ACTIVATION

Depolarization of the neuronal plasma membrane represents the release of potential energy, previously stored through the establishment of transmembrane ion gradients, especially of and This precipitous event initiates the nerve impulse. As the amount of energy made available through depolarization is relatively fixed, an increase in signal intensity is achieved only through an increase in the frequency with which ion gradients can be repeatedly re-established. This frequency not only determines signal amplitude, but may also influence its precision9 and subserve cortical organization of cognitive motor processes10.


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Re-establishment of transmembrane ion gradients requires expenditure of chemical energy, i.e., consumption of ATP, in an amount at least equal to the potential energy that will be consumed in the next depolarization and exceeding it to the extent that the overall process is inefficient, e.g., as the result of transmembrane ion leakage. Clearly, the overall energy cost to the neuron increases with signal amplitude, reflecting requisite acceleration of the depolarization/repolarization cycle. Additional neurons may also be recruited to participate in the event. Increased intensity or duration of the signal, e.g., as would accompany a sustained voluntary muscle contraction or seizure activity, would further heighten each cell’s demand for ATP. That the rate at which ATP is made available may become limiting under conditions of such increased demand is demonstrated by evidence that ATP concentration in rat brain synaptosomes decreases as a result of increased ion pumping, eventually reaching a level at which hexokinase kinetics and thus glycolytic activity become limiting11,12. Decreased cellular ATP concentrations also inhibit exocytosis of neurotransmitters, while augmenting their transmembrane release12. The activation-induced demand for increased neuronal ATP generation thereby creates a demand for increased availability of oxidizable substrate and increased activity of cellular mechanisms for its utilization. These demands are met in part through extraneuronal processes initiated by the depolarization itself. Thus, a depolarizationinduced rise in extracellular possibly associated with a nitric oxide 13,14 (NO) effect , promotes dilation of arterioles in the vicinity, thereby augmenting regional blood flow and glucose extraction, brain uptake of which is flow-dependent. Importantly, the uptake of ketone bodies and FFA is not flow-dependent; thus, their extraction from plasma remains relatively constant and therefore diminished relative to the neuronal activation-associated flow-dependent increase in glucose uptake. The depolarization-induced rise in extracellular also triggers responses in astrocyte function that help support neuronal activation15 (also, see Chapter 13.3.: Amino Acid Metabolism and the Glutamine Shuttle).

11.3.

MITOCHONDRIAL FUNCTION AND THE GENERATION OF ATP

Neuronal activation initiates, and absolutely depends upon, increased cellular generation of ATP, i.e., accelerated conversion of ADP to ATP. Theoretically, this could be accomplished through anaerobic glycolysis, as in exercising muscle in which glucose is converted to lactate; in this

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situation, lactate is exported and subsequently undergoes oxidation or reconversion to glucose via gluconeogenesis in liver. Although yielding a net of only 2 molecules of ATP for each molecule of glucose, anaerobic glycolysis is potentially useful under conditions in which glucose (as glycogen) is abundant and oxygen is limited, e.g., as in the exercising myocyte. In the relatively glycogen-poor neuron, however, the energy demands associated with repeated cycles of depolarization and repolarization, occurring within fractions of a second, require the far more efficient process of oxidative phosphorylation16, through which complete oxidation of one glucose molecule generates 30 molecules of ATP. Mitochondrial generation of ATP through oxidative phosphorylation (Fig. 3.2, facing p. 21) is driven by, and directly proportional to, the electrochemical potential gradient across the inner mitochondrial membrane This gradient is generated and maintained by the proton-extruding activity of complexes I, III, and IV of the electron transport chain. The drives protons back across the inner membrane through the ATP synthase (complex V) where oxidative phosphorylation converts ADP to ATP. Development and maintenance of the heightened necessary to drive ATP biosynthesis at an increased rate requires augmented flow of electrons through the electron transport chain. Increased electron flux, in turn, depends both on availability of oxygen (as electron acceptor) and on an increase in the utilization of oxidizable substrates (as electron donors) to fuel the citric acid cycle17,18. As a corollary, impairment of the processes upon which mitochondrial energetics depend will impair or abrogate neuronal activation, synaptic function, and neurotransmission. In subsequent chapters, evidence will be discussed which supports the concept that the various oxidizable substrates potentially available for neuronal energy generation differ substantially in the efficiency with which their mitochondrial utilization meets these requirements. As a direct corollary, they also differ substantially in their propensity to generate intramitochondrial oxidative stress and to induce neuronal injury.

11.4. REFERENCES 1. P. J. Randle, Regulatory interactions between lipids and carbohydrates: the glucose fatty acid cycle after 35 years, Diabetes Metab Rev 14:263-83 (1998). 2. G. Boden, and G. I. Shulman, Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction, Eur J Clin Invest 32 Suppl 3:14-23 (2002).


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3. J. D. McGarry, Banting lecture 2001: dysregulation of fatty acid metabolism in the etiology of type 2 diabetes, Diabetes 51:7-18 (2002). 4. S. Mills, D. Foster, and J. McGarry, Interaction of malonyl-CoA and related compounds with mitochondria from different rat tissues:Relationship between ligand binding and inhibition of carnitine palmitoyltransferase I, Biochem J 214:83-91 (1983). 5. L. Drynan, P. Quant, and V. Zammit, Flux control exerted by mitochondrial outer membrane carnitine palmitoyltransferase over beta-oxidation, ketogenesis and tricarboxylic acid cycle activity in hepatocytes isolated from rats in different metabolic states, Biochem J 317:791-795 (1996A). 6. L. Drynan, P. Quant, and V. Zammit, The role of changes in the sensitivity of hepatic mitochondrial overt carnitine palmitoyltransferase in determining the onset of the ketosis of starvation in the rat, Biochem J 318:767-770 (1996B). 7. J. Sleboda, K. Risan, O. Spydevold, and J. Bremer, Short-term regulation of carnitine palmitoyltransferase I in cultured rat hepatocytes: spontaneous inactivation and reactivation by fatty acids, Biochim Biophys Acta 1436:541-549 (1999). 8. J. McGarry, and N. Brown, Reconstitution of purified, active and malonyl-CoAsensitive rat liver carnitine palmitoyltransferase I: relationship between membrane environment and malonyl-CoA sensitivity, Biochem J 349:179-187 (2000). 9. J. Lisman, Bursts as a unit of neural information: making unreliable synapses reliable, Trends Neurosci 20:38-43 (1997). 10. A. Riehle, S. Grim, M. Diesmann, and A. Aertsen, Spike synchronization and rate modulation differentially involved in motor cortical function, Science 278:19501953 (1997). 11. M. Erecinska, D. Nelson, J. Deas, and I. Silver, Limitation of glycolysis by hexokinase in rat brain synaptosomes during intense ion pumping, Brain Res 726:153-159 (1996A). 12. M. Erecinska, D. Nelson, and I. Silver, Metabolic and energetic properties of isolated nerve ending particles (synaptosomes), Biochim Biophys Acta 1277:13-34 (1996B). 13. C. Iadecola, D. Pelligrino, M. Moskowitz, and N. Lassen, Nitric oxide synthase inhibition and cerebrovascular regulation, J Cerebral Blood Flow Metab 14:175-192 (1994). 14. G. Bonvento, N. Sibson, and L. Pellerin, Does glutamate image your thoughts?, Trends Neurosci 25:359-64 (2002). 15. E. Kaufman, and B. Driscoll, Evidence for cooperativity between neurons and astroglia in the regulation of CO2 fixation in vitro, Dev Neursci 15:299-305 (1993). 16. O. Kann, S. Schuchmann, K. Buchheim, and U. Heinemann, Coupling of neuronal activity and mitochondrial metabolism as revealed by NAD(P)H fluorescence signals in organotypic hippocampal slice cultures of the rat, Neuroscience 119:87-100 (2003). 17. E. Jonas, J. Buchanan, and L. Kaczmarek, Prolonged activation of mitochondrial conductances during synaptic transmission, Science 286:1347-50 (1999). 18. D. G. Nicholls, Mitochondrial function and dysfunction in the cell: its relevance to aging and aging-related disease, Int J Biochem Cell Biol 34:1372-81 (2002).

Chapter 12 Utilization of Oxidizable Substrates in Brain

12.1. INTRODUCTION As was discussed in Part II, support of cell cycle progression and mitosis, and other time- and energy-critical processes, depends on utilization of specific substrates. Thus, mere abundance of carbon sources is not only insufficient but potentially counterproductive and/or deleterious. In the present chapter, evidence indicating that a similar selectively is required for energization of neuronal activation will also be presented. This selectivity is manifest despite the fact that the major oxidizable substrates present in plasma readily enter brain via more or less well-defined mechanisms. Furthermore, competitive interaction among alternative substrates appears to influence neuronal activity and energetic efficiency in health, and potentially contribute to pathogenesis of neuronal disease. In contrast to most neurons, glial cells, e.g., astrocytes, are able to utilize a broad range of substrates; these differences between astrocytes and neurons, and their equally important functional consequences, are developed in greater detail in the following chapter. Notwithstanding these contrasts, however, available evidence strongly suggests that selected populations of brain neurons, especially those located in hypothalamic centers involved in neuroendocrine and metabolic regulation, exhibit a distinctive profile of substrate utilization, differing substantially from that of their counterparts in other brain regions that are involved in synaptic transmission. Their distinctive metabolic characteristics may be fundamental to their role in regulation of feeding behavior.

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12. 2. GLUCOSE Glucose uptake for utilization in brain 1 is mediated by members of a family of facilitative glucose transporters2. GLUT 1 principally accounts for uptake across the blood brain barrier and into astrocytes, whereas GLUT 3 predominates in neurons. Kinetic characteristics suggest that transport of glucose from interstitial fluid into cells would be more efficient for GLUT 3 (neurons) than GLUT 1 (astrocytes), but anatomical factors and transporter abundance likely influence these rates in vivo. Insulin receptors are widely expressed in brain3-5, but insulin’s effect on glucose uptake has been controversial. Although insulin induces carotid artery vasodilation6, it does not directly regulate overall glucose uptake 7 ; evidence that insulin importantly influences glucose metabolism in brain will be discussed (see Chapter 15.4.: Cerebral Metabolism and Insulin Signaling). Moreover, neuron-specific disruption of the insulin receptor gene caused profound changes in feeding behavior, body fat, and reproductive function 8 , while insulin-like growth factor, which utilizes similar signal transduction pathways, contributes to regulation of glucose metabolism in developing brain 9 . The basis for the dependence of normal brain function on glucose is not intuitively apparent, given the abundance of other potential substrates. Its importance in this context becomes evident, however, upon consideration of the efficiency with which glucose oxidation supports mitochondrial respiration, and oxidative phosphorylation. Two important characteristics contribute to this effectiveness. First, the overall stoichiometry of glucose oxidation is quite favorable. Thus, for each molecule of glucose fully oxidized to and through glycolysis and the citric acid cycle (i.e., aerobic glycolysis), approximately 30 molecules of ATP are generated and 6 oxygen molecules are consumed 10 . As will be seen, this theoretical ratio of ATP molecules generated to oxygen molecules consumed (5.0 for glucose) exceeds that for ketone bodies and fatty acids. In anaerobic glycolysis, in contrast, only 2 molecules of ATP and 2 molecules of lactate are produced for each molecule of glucose utilized. Second, and perhaps even more important, several additional factors render the overall process of aerobic glycolysis especially efficient. Thus, in those cell populations in which high rates of ATP generation are required — e.g., neuronal activation and proliferating cells — hexokinase, which catalyzes the formation of glucose-6-phosphate and thus glucose entry into the glycolytic pathway, is principally associated with mitochondria11,12. This association comprises a component of the

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Figure 12.1. Mitochondrial voltage-dependent anion channel (VDAC) components: relationship to electron transport chain and generation of reactive oxygen species. While detailed structural relationships remain undefined (see Chapter 3.3.1.: Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation), juxtaposition of hexokinase to the adenine nucleotide translocase in neurons and proliferating cells optimizes kinetics of glucose phosphorylation to glucose-6-phosphate and entry into the glycolytic pathway, leading to increased pyruvate formation. Increased mitochondrial fatty acid oxidation inhibits both pyruvate dehydrogenase and hexokinase (Randle cycle). Excess cellular fatty acid abundance may disrupt both VDAC and the contact sites, and overload the electron transport chain. This predisposes to “leakage” from complexes I and III, and ubiquinone, with formation of ROS. Pro-apoptotic members of the Bcl-2 family (e.g., Bax) may be largely cytosolic and associate with the mitochondrial membrane only after activation of the apoptotic cascade. Abbreviations — ADP/ATP: adenosine di-/triphosphate; CYT C: cytochrome c; inner mitochondrial membrane potential; UBIQ: ubiquinone; ROS: reactive oxygen species; VDAC: voltage-dependent anion channel. With permission (see Fig. 3.4.)

mitochondrial voltage-dependent anion channel (VDAC) complex (Fig. 12.1) located at the contact sites between inner and outer mitochondrial membranes. In addition to hexokinase II, the complex includes the adenine nucleotide translocase, porin, creatine kinase, and anti-apoptotic members of the Bcl-2 family12-14. While still incompletely defined15, these structural relationships appear to enhance the rate and efficiency of both hexokinase-mediated glucose phosphorylation and ATP synthasemediated oxidative phosphorylation. Thus, they facilitate access of newly synthesized ATP to the hexokinase, and of the ADP thereby generated to the mitochondrial ATP synthase. In addition, VDAC conductivity may

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be modulated directly by glutamate16 (also, see Chapter 14.5.1.: Energetic Considerations in Synaptic Transmission). Citric acid cycle utilization of the pyruvate produced via glycolysis is highly efficient (Fig. 3.2, facing p. 21). This results from the fact that PDH, the key enzyme in this process, is subject to negative feedback control, resulting in stringent regulation of pyruvate utilization and therefore overall glycolytic rate17. Thus, PDH is subject to kinasemediated inactivation by high ratios of NADH/NAD, acetyl CoA/CoA, and ATP/ADP in the mitochondrial matrix, i.e., those conditions that indicate abundance of reducing equivalents, citric acid cycle substrate, and electron transport chain flux. This kinase-mediated feedback regulation of PDH prevents excessive oxidative decarboxylation of substrate (pyruvate) to acetyl CoA, thereby controlling the flow of electrons into the electron transport chain. In the absence of such stringent control, electrons would be more likely to enter the electron transport chain in excessive amounts and therefore more likely to “leak” from complexes I and III, and ubiquinone. Through one-electron reduction of oxygen to form superoxide anion these electrons contribute to the generation of ROS and oxidative stress. By continuously modulating the rate of glucose (thus, pyruvate) utilization in response to the status of the electron transport chain and intramitochondrial redox potential, PDH operates in effect as the biological equivalent of an automobile’s computerized fuel injection system. As will be discussed in detail, this efficiency stands in marked contrast to the less tightly regulated utilization of ketone bodies and fatty acids in mitochondrial energygenerating pathways. Any inefficiency in energy generation would likely have proved disadvantageous in terms of evolutionary survival: it not only would have jeopardized attainment of rapid and repeated increases in oxidative phosphorylation required for optimal neuronal activation and synaptic function, but also would have been potentially injurious to the cell and thus to the organism through generation of oxidative stress.

12.3. KETONE BODIES The initial demonstration that acetoacetate and could partially substitute for glucose as a major substrate in fasting human brain18 provided important new insight into neuronal energy metabolism. Although starvation increases brain mitochondrial dehydrogenase activity19, it is clear that utilization of ketone bodies in adult brain, principally neuronal20, is not dependent on


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an adaptive process induced by prolonged fasting. Thus, it also occurs in short-term (3.5 days) starvation21,22, as well as in healthy, non-fasting subjects23. Moreover, brain utilization of glucose is suppressed by ketone bodies. This effect is not related to glucose transport kinetics across the blood brain barrier, but rather appears to reflect a change at or beyond the hexokinase-mediated entry of glucose into the glycolytic pathway21,24,25. Given that glucose clearly is the energetically optimal fuel for support of neuronal activation, how is it possible that the less consistently available and (as will be discussed) less efficiently utilized and potentially injurious ketone bodies are immediately accepted as a preferred substrate? The answer to this important question is found in a consideration of the determinants of ketone body transfer from plasma to brain to neuron to mitochondria, and their ultimate utilization in mitochondrial generation of ATP. Ketone body extraction from plasma into brain exhibits regional variation but is directly proportional to plasma concentration22,26. A blood brain barrier transport mechanism shared by other low molecular weight monocarboxylates appears to be a major but relatively constant rate-determinant of the uptake process20,26. Moreover, a carrier-mediated cellular uptake process27,28 operates with sufficiently high efficiency that ketone body concentrations in brain interstitial fluid remain low26. Thus, ketone bodies that have exited plasma and crossed the blood brain barrier rapidly traverse the intercellular space, and enter both neurons and astrocytes, where they undergo mitochondrial oxidation29,30. Relative to glucose, however, the extraction fraction of ketone bodies from cerebral plasma is low26, as a result of which brain uptake is not flow-limited31. Consequently, and in contrast to the increased glucose extraction that occurs as the consequence of neuronal activation-associated changes in regional blood flow, uptake of ketone bodies would be expected to remain relatively constant. The mechanism by which ketone bodies enter the mitochondrial matrix remains incompletely understood, as no specific transporter has been identified32. It has been suggested that in liver mitochondria ketone bodies are transported across the inner membrane by the pyruvate carrier33, but evidence is conflicting34. It has been generally believed, however, that as cytosolic ketone body concentration increases so does the importance of non-ionic diffusion of the protonated acid across the inner mitochondrial membrane33,34. In any case, the overall process is driven by the concentration gradients between plasma, interstitial fluid, cytosol, and mitochondrial matrix. The ability of ketone bodies to suppress and substitute for glucose utilization is readily understandable in terms of their intramitochondrial

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Figure 12.2. Fatty acid and ketone body oxidation in mitochondria suppresses glycolysis. Mitochondrial of fatty acids (astrocytes) and ketone bodies (neurons) generates three moieties which activate physiological kinase-mediated feedback inhibition of PDH, i.e., acetyl CoA, ATP, and NADH. This effect, together with an accompanying suppression of glycolysis, decreases mitochondrial oxidation of pyruvate, and overall utilization of glucose. In peripheral tissues, especially muscle, this effect of mitochondrial fatty acid oxidation contributes to the “Randle cycle” inhibitory effect on glucose utilization and insulin resistance. Dashed lines: inhibition of PDH by acetyl CoA/CoA, ATP/ADP, and NADH/NAD. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; NADPH: nucleotide adenine triphosphate, reduced form; PDH: pyruvate dehydrogenase.

metabolism (Fig. 12.2). Utilization of requires its oxidation to acetoacetate by dehydrogenase, an exclusively mitochondrial enzyme35. Acetoacetate is then converted to its acyl CoA thioester, either directly or via succinyl CoA acyltransferasemediated transfer of CoA from succinyl CoA, an intermediate in the citric acid cycle; it is acetoacetyl CoA that enters the oxidative pathways in the mitochondrial matrix. Two consequences of ketone body entry into the mitochondrial matrix are especially significant in the present context. First, there is no evidence that excessive ketone body oxidation and overload of the electron transport chain are prevented by a regulatory process in a manner analogous to PDH-mediated control of the utilization of pyruvate (and, thus, of glucose). Moreover, ketone bodies may adversely affect the efficiency of oxidative phosphorylation by two distinct mechanisms: a) a minor uncoupling of oxidative phosphorylation resulting from the movement of protonated ketone bodies across the inner mitochondrial membrane; and b) a more significant uncouplinglike effect, similar to that caused by other short chain carboxylic acids36


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(also, see Chapter 6.2.: Butyrate). This uncoupling-like effect results from a futile cycle of synthesis and hydrolysis of acetoacetyl CoA37, consuming ATP and generating AMP within the mitochondrial matrix. The minor protonophoric component is exceeded stoichiometrically by the electrons generated through intramitochondrial oxidation of ketone bodies and short chain fatty acids, which thus increase redox potential and predispose to ROS generation. As a result, the antioxidant effect that is seen with non-metabolized protonophoric uncouplers38-42 is absent. In its place is a substrate oxidation-induced, potentially ROS-generating, increase in electron transport chain flux, as would occur with oxidation of both ketone bodies43 and the short chain fatty acid butyrate44. Second, utilization (mitochondrial oxidation) of acetoacetyl CoA in short-term perfused heart and liver, isolated liver mitochondria45, and tumor cells in culture46, leads to the generation of acetyl CoA, NADH, and ATP. As a result, PDH is inactivated, thereby inhibiting pyruvate utilization, glycolysis, and glucose uptake; such effects are also caused by oxidation of short, medium, and long chain fatty acids17,47,48 (Fig. 12.2). This adverse effect is similar to the Randle cycle in which excessive oxidation of fatty acids inhibits glucose utilization and contributes to insulin resistance in skeletal muscle49-51. Thus, during simultaneous perfusion of rat heart with acetoacetate and glucose, oxidation of glucose was dramatically suppressed52, although cardiac performance was unimpaired52,53. Provision of 5mM acetoacetate alone, however, caused substantial deterioration of cardiac performance relative to that with glucose, and suppressed mitochondrial CoA abundance, citric acid cycle activity, and state 3 respiration54,55. Accordingly, ketone bodies as sole substrate are unable to sustain the energy requirements of the cardiac myocyte, another terminally differentiated cell which in terms of mitochondrial metabolism further resembles the neuron in that it lacks the ability to carry out either gluconeogenesis or ketogenesis45, but which differs importantly from it in its general dependence energetically on mitochondrial fatty acid oxidation (see Chapter 1: Disparate Themes: Origins and Integration). Studies of rat hippocampal slices also indicate important differences between glucose and ketone bodies in their ability to support neuronal viability and synaptic function56. Thus, protected 1457 day embryonic hippocampal cells from , and restored excitatory postsynaptic potentials in glucose-deprived or glycolysisinhibited slices from 15-day postnatal (suckling) rats56. In slices from 120-day (adult) animals, in contrast, was unable to support synaptic function56. Ketone bodies also support lipid synthesis in developing brain58,59, and in proliferating neural cell lines in vitro60. The

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more effective utilization of ketone bodies, as well as lactate, in newborn and suckling animals parallels monocarboxylate carrier expression28. In fetal rabbits, maternal diabetes mellitus and hyperketonemia supressed fetal glucose utilization61. Similarly, in neural tissue from 11- and 12-day rat embryos, suppressed glucose utilization, suggesting a potential pathogenetic role in maternal diabetes mellitusassociated teratogenesis62. Significantly, the apparent shift from glucose to utilization in the latter system did not affect oxygen consumption, a phenomenon that reappears in other informative settings (see below; also, see Chapter 14.2.: Activation-Related Glucose and Oxygen Consumption, and Chapter 15.4.: Cerebral Metabolism and Insulin Signaling). On the basis of these considerations, it is clear that in the developed brain a relative increase in ketone body utilization in the resting neuron may spare glucose but exacts a price nonetheless. Thus, ketone bodyinduced inhibition of the citric acid cycle and uncoupling-like effects on oxidative phosphorylation would compromise and therefore the rate of ATP generation, relative to levels supported by glucose. In addition, the relatively unregulated mitochondrial oxidation of ketone bodies could increase electron flux and reducing potential in the electron transport chain, increasing the likelihood of electron “leakage”, especially at complexes I and III, and ubiquinone (Fig. 12.1). The resulting generation of ROS and oxidative stress would inhibit several components of the electron transport chain, thereby further compromising Together, such inefficiencies in ketone body oxidation would obligate a higher rate of oxygen consumption relative to ATP production than would be required during predominant fueling of the process by the far more efficiently utilized glucose. Therefore, although the theoretical maximum yield of 4.75-4.78 molecules of ATP for each oxygen molecule consumed during complete oxidation of ketone bodies is calculated to be only slightly less than that for glucose (5.0)10, the actual yield would be further diminished by uncoupling-like effects, electron “leakage”, and generation of ROS. These characteristics of ketone bodies (and of short chain monocarboxylic acids in general) provide important perspective in regard to the suggestion that ketone bodies not only are innocuous with respect to neuronal function and viability, but are in fact neuroprotective43,57. The experiments upon which these conclusions were based involved an apparent protective effect of ketone bodies against acutely lethal insults, and/or in embryonal neurons. Consistent with these observations, it has been shown that whereas ketone bodies support viability of neurons from embryonic, post-weaning, and adult animals during glucose deprivation,


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and support synaptic function in neurons from embryonal and suckling animals, they fail to support the critical elements of cognitive function in neurons from post-weaning and adult rats56. Thus, ketone bodies may protect against potentially lethal acute insults at perhaps all stages of neuronal development, but they are unable to support the more exacting metabolic and energetic requirements for synaptic function in the developed brain. These developmental differences may also account for the observation that ketone bodies protected embryonal neurons from 57 acute toxicity induced by amyloid and , and protected rat 43 nigrostriatal dopaminergic neurons against in a model of Parkinson’s disease. As inhibits Complex I of the electron transport chain, sustained support of mitochondrial oxidative phosphorylation and neuronal viability in the presence of this agent would require that electrons be supplied “downstream” of this blockade. Ketone body oxidation accomplishes this by providing electrons to the electron transport chain via complex II (succinate dehydrogenase). While this may maintain neuronal viability under the specified experimental conditions, it would not provide for the tightly-regulated energetic foundation required for support of normal synaptic function in the mature neuron (e.g., see Chapter 14.5.2.1.: Long Term Potentiation). Moreover, it would contribute to the sustained alterations in neuronal substrate utilization, metabolism, redox balance, and energetics that characterize Alzheimer and Parkinson diseases, and which may precede cell death by years or decades. Despite the inefficiencies and other adverse effects associated with ketone body metabolism, and the resulting potential for compromise of neuronal activation, their preferential utilization may be regarded as advantageous under conditions in which glucose availability is limited and ketone bodies abundant, i.e., during starvation. Moreover, to the extent that the physiological and metabolic correlates of neuronal switching between activation and inactivation are also associated with shifts in substrate utilization, the appropriate and timely coordination of these shifts may be a pivotal determinant of neuronal function. As will be discussed, preferential utilization of ketone bodies and the resulting decrease in glucose utilization despite constant oxygen consumption22,62 provides insight into what appears to represent the reverse phenomenon energetically, i.e., neuronal activation. During the latter process oxygen consumption may be relatively constant despite an increase in glucose utilization. Furthermore, these relationships support the concept that utilization of ketone bodies as substrates (and thus, determinants of functional status) may be relatively greater in the non-activated neuron

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(see Chapter Consumption).

14.2.:

Activation-Related

Glucose

and

Oxygen

12.4. FATTY ACID UTILIZATION IN BRAIN AND THE REGULATION OF APPETITE 12.4.1. Uptake of Fatty Acids from Plasma Long chain free (unesterified) fatty acids (FFA) are poorly soluble in aqueous media and in plasma are tightly bound to albumin. For that reason they would be expected to have less ready access to the brain than the water-soluble glucose and ketone bodies. Despite this, brain uptake of plasma FFA is well documented63-65, as is their rapid metabolism in both oxidative and esterification pathways66-68. As in other tissues, uptake is likely directly proportional to the plasma FFA concentration and the fatty acid:albumin molar ratio69. Regional FFA uptake corresponds generally to the activity of oxidative metabolism as reflected by 2-deoxyglucose uptake70. Similar to ketone bodies, FFA extraction from plasma is not flow–dependent and is relatively low compared to that of glucose31,68. As a result, and also true of ketone bodies, FFA uptake would be expected to remain relatively constant despite the fluctuations in regional blood flow and glucose extraction that are associated with neuronal activation. This interpretation is consistent with the demonstration that incorporation of fatty acids into brain lipids is flow-independent71. Although lipoprotein lipase is expressed in cortex and other regions of the brain 72 , its activity is much lower than in spinal cord73 and non-neural tissues; as a result, the contribution of lipoprotein fatty acid esters to overall brain fatty acid uptake would appear to be less than that of FFA74. Because long chain fatty acids are less soluble in aqueous media than shorter chain fatty acids and ketone bodies, they exist both intra- and extracellularly in association with proteins, i.e., albumin in plasma and interstitial fluid, and cytosolic fatty acid binding proteins (FABP) and other moieties within cells75. Fatty acid entry into cells appears to be facilitated by plasma membrane binding proteins and/or transporters69. Within cells, they may enter either esterification pathways, e.g., leading to the biosynthesis of membrane lipids, or oxidative pathways, principally in mitochondria (Fig. 12.3). Fatty acid entry into almost all of these pathways requires initial conversion to the acyl CoA thioester, a reaction catalyzed by an isoform of long chain acyl CoA synthase that is relatively brain-specific and homologous to the liver isoform76. In rat


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Figure 12.3. Pathways of cellular fatty acid metabolism. Both mitochondrial and extramitochondrial FA oxidation pathways are transcriptionally regulated by members of the PPAR family. PPAR may be activated by increased cellular abundance of FA, as well as other factors acting directly or indirectly. Absence of in primary human brain astrocytes and astrocyte-derived lines (see text) suggests that these cells may be compromised in regard to a potentially protective adaptive response to increased fatty acid exposure (see Chapter 12.4.2.: Oxidation of Fatty Acids). Abbreviations — CPT-I: carnitine palmitoyltransferase-I; FA: fatty acid; FA-CoA: fatty acid-coenzyme A thioester; PL: glycerophosphatides; PPAR: peroxisome proliferator-activated receptor; TG: triacylglycerols.

and primate brain, a substantial fraction (i.e., at least 50% of palmitate) 66-68,77 undergoes , indicating that fatty acid oxidation may contribute to energy generation in brain. The potential significance of plasma FFA uptake for brain energy metabolism in humans is suggested by a 9-fold increase in brain FFA uptake during short-term starvation21. Assuming total utilization of these FFA for ketogenesis in astrocytes78, the quantity of ketone bodies generated, e.g., for neuronal utilization, would equal 49.3% of those which entered brain as such from plasma21, i.e., approximately one-third of the combined total. Although the observed increase in FFA uptake did not achieve statistical significance21, such observations are of potentially

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great importance. Calculation of brain FFA uptake has usually been based on arterio-venous differences, and involves the difference between two relatively large, inherently variable numbers. Thus, it is necessarily imprecise, and may underestimate actual FFA uptake, e.g., as demonstrated by other methods such as those involving isotopic techniques as discussed above.

12.4.2. Oxidation of Fatty Acids Mitochondrial of long chain fatty acyl CoA thioester, as noted (see Chapter 11.1.: Introduction - Relevant Principles of Intermediary Metabolism), requires transacylation by the ratedetermining enzyme CPT-I to form the acylcarnitine derivative. Rat and mouse brain express the liver isoform of CPT-I, which exhibits relatively high affinity for carnitine and relatively low susceptibility to inhibition by malonyl CoA78,79. In addition, a novel CPT-I with unusual substrate specificity was recently identified in mouse and human brain, most abundantly in the hippocampus and in hypothalamic nuclei involved in control of feeding behavior80. Acylcarnitines formed by CPT-I are transferred across the inner mitochondrial membrane to enter the spiral (Fig. 12.4B and Fig. 3.2, facing p. 21). Agents that inhibit CPT-I acutely inhibit fatty acid in brain as in other tissues in vivo77,81-83, thereby documenting the mitochondrial location of this process; esterification is increased as a result, largely in the phospholipid fraction. Significantly, feeding behavior is also increased by CPT-I inhibition84-86, an observation of considerable importance that is Figure 12.4. Relevant aspects of intermediary metabolism. Pathway arrow thickness approximates relative activity; dotted lines: relatively or absolutely suppressed; enclosed areas represent mitochondria. A. Glycolysis- and citric acid cycle-fueled ATP synthesis. Complete oxidation of glucose via aerobic glycolysis, pyruvate dehydrogenase, and the citric acid cycle, yields 30 molecules of ATP per molecule of glucose. Gluconeogenesis i s suppressed. B. Fatty acid oxidation. Mitochondrial oxidation of long chain fatty acids requires CPT-I-mediated formation of acylcarnitine. Fatty acids present in excess may override malonyl CoA inhibition of CPT-I, driving inappropriately augmented mitochondrial oxidation. The theoretical maximum yield of ATP from complete oxidation of palmitate (~100) is variably diminished to as little as ~20 in a ketogenic cell (astrocyte or hepatocyte) by diversion of acetyl CoA to ketogenesis rather than to the citric acid cycle. C.Gluconeogenesis. The process is normally fueled energetically by the mitochondrial oxidation of fatty acids. Glycolysis is suppressed. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; F26BP: fructose-2,6bisphosphate; NADPH: nucleotide adenine triphosphate, reduced form; PCB: pyruvate carboxylase; PEPCK: phosphoenolpyruvate carboxykinase; PL: glycerophosphatides; TG: triacylglycerols.


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further discussed below (see this Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior). Products of fatty acid include as well as citric acid cycle intermediates and ketone bodies; a substantial fraction of administered radioactivity is also recovered in aspartate and glutamate87. Acetyl CoA generated by typically enters the citric acid cycle, thus fueling the electron transport chain and oxidative phosphorylation. As in hepatocytes, however, acetyl CoA in astrocytes may be converted to ketone bodies and exported from the cell78,88,89 (Fig. 3.2, facing p. 21). While ketogenesis thus supports the metabolic needs of other cells, ketone body synthesis and export represents a loss to the astrocyte or hepatocyte of potentially oxidizable carbon atoms. Long chain fatty acids also undergo in brain peroxisomes, the activity of which is related to developmental stage90. Although these studies of whole brain do not identify the particular cell population(s) in which plasma-derived long chain fatty acids are metabolized, other evidence suggests that their oxidation occurs primarily in glia, especially astrocytes, and in the subset of neurons involved in neuroendocrine and metabolic regulation (see this Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior). Fatty acids with fewer than 14 carbon atoms may also exert an important influence on brain metabolism under certain circumstances. Thus, short chain fatty acids (SCFA, i.e., 2-4 carbons, e.g., the 4-carbon butyrate) are generated in large quantities by colonic bacteria and are readily absorbed into portal venous plasma. Normally, SCFA are nearly completely extracted during passage through the liver and do not represent a significant potential alternative metabolic fuel for the brain. However, under conditions in which there is substantial portal-systemic shunting as may accompany cirrhosis and other causes of portal hypertension, a fraction of SCFA escapes removal in the hepatic and pulmonary vascular beds and is available to perfuse the brain, potentially contributing to the demonstrated decrease in cerebral glucose utilization in cirrhosis91. As SCFA are far less tightly bound to albumin than are long chain fatty acids, they more readily enter brain64 although in the presence of portocaval shunting their apparent rate of transport across the blood brain barrier may be diminished92. Medium chain fatty acids (MCFA, i.e., 6-12 carbons, e.g., the 8-carbon octanoate) also enter brain Similar to SCFA, MCFA can enter mitochondria from plasma64. independent of CPT-I (Fig. 12.2). While MCFA are not normally produced within the body in substantial quantities, they may become available through their administration as a nutritional supplement or, in the special case of valproate (2-propyl-pentanoate, i.e., an iso-octanoate), as an anticonvulsant. In general, SCFA and MCFA exert effects on


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mitochondrial metabolism similar to those of ketone bodies, i.e., relative lack of control by CPT-I, uncoupling-like effects36, and inhibition of PDH and glycolysis17,47,48 (see Chapter 6.2.: Butyrate, and Fig. 6.2, facing p. 123). Valproate and its derivatives also inhibit enzymes in the spiral itself 93. Most cells that utilize significant quantities of long chain fatty acids express certain genes the products of which serve related regulatory functions (Fig. 12.3). These include one or more members of the nuclear receptor family of peroxisome proliferator-activated receptors (PPARs)94,95, proteins that may be regulated by PPARs such as cytosolic FABP, fatty acid enzymes in peroxisomes and mitochondria, and antioxidant defenses such as glutathione peroxidase and catalase. The FABP can be viewed as protective against the potentially adverse (e.g., surface active) effects of fatty acids themselves, and against the oxidative stress that would be expected if their cellular abundance and mitochondrial or peroxisomal become excessive. In this connection, it is highly significant that human primary astrocytes and a human astrocytoma cell line were found to express (also designated and unlike rat brain astrocytes, however, neither cell type from human brain expressed 96 . The isoform is particularly important in activating those genes that encode FABP and the mitochondrial and peroxisomal enzymes that are important in the adaptive cellular response to increased fatty acid abundance. As in the hepatocyte97, absence of this critical nuclear receptor suggests that astrocytes are relatively compromised in the adaptive response which it activates, and therefore are predisposed in states of fatty acid excess to fatty acid oxidation-induced intramitochondrial oxidative stress and ROS generation98 (also, see Chapter 5.2.1.: and Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). In addition, exported astrocyte-generated diffusible ROS (e.g., would adversely affect neighboring neurons. Consistent with this concept, there is substantial evidence that oxidation of long chain fatty acids may have adverse effects on mitochondrial energetics, maintenance of and the efficiency of ATP generation. Thus, in addition to PDH inhibition, high fatty acid flux rates may override or circumvent the regulatory step at CPT-I99-103, thereby potentially overloading the electron transport chain and predisposing to oxidative stress. While underivatized long chain fatty acids may also weakly uncouple oxidative phosphorylation via a protonophoric effect36, their relatively modest antioxidant effect would be obscured in states of

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fatty acid excess by the pro-oxidant effect of high fatty acid oxidation rates. Given these considerations, it is perhaps not surprising that in most neurons in adult brain, there is little or no mitochondrial oxidation of long chain fatty acids. Limited neuronal capacity for fatty acid oxidation is further supported by expression of the sterol regulatory element binding protein-1 (SREBP-1). SREBPs are key transactivators of genes related to the biosynthesis of isoprenoids (e.g., sterols and farnesyl derivatives) and fatty acids104. Such transcriptional activation of the fatty acid biosynthetic pathway in a cell not evidently active in fatty acid biosynthesis except in axons105 would seem to be wasteful or even counterproductive. However, the increased activity of acetyl CoA carboxylase that results would generate increased quantities of malonyl CoA, thereby suppressing CPT-I and mitochondrial fatty acid oxidation. In this respect, the neuron would further resemble the cardiac myocyte, a cell in which de novo fatty acid biosynthesis is also insignificant, but in which expression of the initial committed step in this pathway, i.e., acetyl CoA carboxylase-mediated malonyl CoA synthesis, contributes to the regulation of mitochondrial fatty acid oxidation through inhibition of CPT-I106,107. In addition, to the extent that cytosolic FABP, catalase, and glutathione peroxidase are expressed in brain, they are largely confined to glial cells, and in particular to astrocytes (see Chapter 13.4.: Fatty Acids, Antioxidant Defense, HIV Dementia, and Cell Proliferation). Thus, as neuronal activation is critically dependendent on a highly efficient, rapidly responsive, and uncompromised augmentation of a mitochondrial increase in the rate of ATP generation, glucose utilization clearly is advantageous as compared to that of the potentially disruptive oxidation of fatty acids and ketone bodies. Somewhat unexpectedly, therefore, both neurons and astrocytes exhibit comparable activity of cytochrome P450 4A-mediated of medium and long chain fatty acids in endoplasmic reticulum108 (Fig. 12.3); the activity of this enzyme in both cell types is equal to or greater than that in hepatocytes. Moreover, a recently described novel isoform, P450 4X1, is expressed predominantly in brain neurons109. As the long chain dicarboxylic acid products of this reaction are potentially toxic and induce peroxisome proliferation in responsive cell types110, the functional significance of this activity in brain is not apparent for either cell type.


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12.4.3. Fatty Acid Oxidation and Regulation of Feeding Behavior In contrast to neurons involved in synaptic function, available evidence suggests strongly that fatty acid oxidation is physiologically important in those neurons involved in neuroendocrine and metabolic regulation. Thus, in rodents and primates, uncoupling protein 2 (UCP2) is expressed in the mitochondria of such neurons, e.g., in the hypothalamus111-114. The uncoupling action of the UCP protein family separates ATP synthesis from flux of protons back across the inner mitochondrial membrane. As a result, the electrochemical energy generated by the electron transport chain and is converted to heat instead of ATP. This accounts for UCP1’s thermogenic effect, e.g., in brown adipose tissue115, and for UCP2’s uncoupling-induced dissipation of intramitochondrial oxidative stress38-42. Because UCP expression and activation are increased by long chain fatty acids115-117, as well as by ROS118, the presence of UCP implies the occurrence of significant mitochondrial fatty acid oxidation in those neurons in which it is expressed. In addition, the hypothalamic regulatory neurons also exhibit a distinctive anatomical and presumably functional relationship to a specific population of astrocytes which abundantly express brain FABP (B-FABP)119, a finding that further suggests a significant relationship to fatty acid metabolism. Thus, regulatory neurons in the hypothalamus differ significantly from their synaptic transmission-related counterparts in other brain regions, in which fatty acid metabolism and antioxidant defenses are minimal. In these latter neurons, fatty acid oxidation and its potential accompaniments — oxidative stress and UCP-mediated uncoupling and thermogenesis120 — would be expected to compromise synaptic function (also, see Chapter 14.5.2.1.: Long-Term Potentiation). The evidence that fatty acids undergo mitochondrial oxidation in hypothalamic neurons involved in metabolic control has important implications for the regulation of feeding behavior. Thus, methyl palmoxirate, an inhibitor of CPT-I-mediated mitochondrial fatty acid oxidation, increases feeding behavior and brain Fos-like immunoreactivity in rats84,85. Similarly, feeding behavior is increased in human subjects by another inhibitor of CPT-I, etomoxir86. In both species, the effect of these agents is most evident when maintenance diets are high in fat84-86, consistent with absence of the effect in short-term human studies121. The reported effect of these agents on feeding behavior has been thought to reflect changes in systemic or hepatic fatty acid metabolism. However, the distinctive characteristics of hypothalamic neurons in regard to fatty acid metabolism would also be consistent with

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an alternative hypothesis: namely, that regulation of mammalian feeding behavior reflects modulation of, and is inversely related to, mitochondrial fatty acid oxidation in specific hypothalamic neuronal populations. The concept that mitochondrial fatty acid oxidation in specific hypothalamic neuronal populations may play a pivotal role in the regulation of feeding behavior would also be consistent with recent evidence that leptin, among its diverse effects122,123, increases systemic energy expenditure and mitochondrial fatty acid oxidation in cardiac and skeletal muscle124-128, adipose tissue129, and endothelial cells130, and augments intramitochondrial ROS generation in the latter. These observations further suggest that the interaction of leptin with hypothalamic neurons, which is critical to leptin’s appetite-suppressing effects, may include a leptin-induced increase in fatty acid oxidation in these cells as in the others cited above. The hypothesis also would serve to integrate hypothalamic with adrenergically-mediated peripheral leptin effects126,131. A functional linkage between leptin signaling and cellular fatty acid metabolism is further evidenced, albeit indirectly, by the demonstration that leptin and fatty acyl CoA thioesters, the latter being obligatory substrates for CPT-I-mediated mitochondrial fatty acid oxidation, activate ATP-sensitive channels: leptin in hypothalamic 132 neurons , and both leptin and acyl CoA in pancreatic islet beta cells133. As discussed in greater detail elsewhere (Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications), increased mitochondrial fatty acid oxidation suppresses cellular insulin signaling and utilization of glucose, and may compromise mitochondrial energetics through generation of ROS. In contrast to this direct relationship between leptin signaling and fatty acid oxidation within the cell, the relationship between their respective concentrations in plasma is inverse; the latter is especially evident during fasting and glucose administration134. This inverse relationship reflects a well-established insulin-induced increase in leptin gene expression and plasma leptin concentration3,135-139. Thus, sustained hyperinsulinemia, under conditions in which glucose concentration is held constant, increases plasma [leptin] and suppresses [FFA]137. Conversely, the fall in plasma [insulin] level that occurs during fasting increases adipocyte lipolysis and plasma [FFA], but suppresses leptin gene expression and plasma concentration. Importantly, changes in plasma [insulin] under these conditions precede those in leptin, which in turn precede significant changes in body weight and adipose tissue mass140. Accordingly, insulin plays a determining role in regulation in leptin formation and secretion. These observations further suggest that augmentation of fatty acid oxidation by leptin124-128 must involve cellular mechanisms that are not


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driven by or dependent upon increases in plasma [FFA], thereby providing the versatility and control required for regulation of feeding behavior. Thus, a leptin-induced, fatty acid oxidation-linked satiety response would need to operate even when plasma [FFA] is normal or low, e.g., during post-prandial hyperinsulinemia134,141,142, while a counterproductive satiety response must be avoided when [FFA] is elevated, e.g., in fasting. This independence from plasma [FFA] appears to depend in part on the sympathetic nervous system126,131. It would be consistent with evidence that leptin induces the oxidation of fatty acids that are intracellular, specifically those derived from cellular triacylglycerols in liver, pancreatic islets, and skeletal muscle143, and with evidence that chronic leptin administration suppresses skeletal muscle expression of the plasma membrane transporter FAT/CD36 and uptake of fatty acids144. Stated simply, then, the present hypothesis holds that feeding behavior (appetite) is at least in part governed by, and inversely related to, leptin-induced mitochondrial oxidation of fatty acids derived from within the cell, in regulatory neurons located primarily in the arcuate nucleus of the hypothalamus. However, the logical extension of this hypothesis would appear to require the existence of a paradoxical relationship between the effects of insulin and leptin in participating hypothalamic neurons. Thus, whereas leptin signaling promotes mitochondrial fatty acid oxidation, insulin suppresses this pathway in diverse tissues124,145. Therefore, participation of insulin in activation of a mitochondrial fatty acid oxidation-dependent satiety response would seem counter-intuitive. Yet, it is conclusively established that insulin, whether administered peripherally, intracerebroventricularly, or into the hypothalamus itself, is a powerful appetite suppressant3,138,139. Moreover, deletion of brain neuronal insulin receptors induces diet-sensitive obesity in mice8, and intracerebroventricular administration of small molecule insulin mimetic agents suppresses appetite146. Thus, insulin’s anorexigenic action is consistently demonstrable despite the fact that it suppresses fatty acid oxidation. Accordingly, if mitochondrial fatty acid oxidation in hypothalamic neurons is to serve as the pivotal determinant of feeding behavior that is hypothesized, opposing effects of insulin and leptin on this pathway must be reconciled. Such reconciliation would of necessity also provide a rational basis for what, as a corollary, must reflect coordinate activation of the satiety response by these dissimilar hormonal agents. At present, information that would definitively provide such rationalization is not available. However, it is significant in this context that the anorexigenic effect of the slowly modulated plasma [leptin] is

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normally manifest following an increase in the more rapidly modulated plasma [insulin], e.g., in association with feeding or weight gain. Conversely, and despite its correlation with body fat stores, leptin’s effects on body weight, and on serum glucose, insulin, and concentrations, are minimal or absent when insulin levels are low, e.g., in fasting and starvation; effects on other aspects of neuro-endocrine signaling are relatively preserved, however147, and may exhibit species differences148. These associations would be consistent with the plausible concept that responsiveness to leptin signaling in the involved regulatory neurons is insulin signaling-dependent. As a corollary, the incompletely understood phenomenon of leptin resisitance may to a large extent reflect resistance to insulin signaling, which commonly accompanies leptinresistance. Further, leptin-induced mitochondrial oxidation of fatty acids, being insulin signaling-dependent, would be subject to inhibition by elevated plasma [FFA], a major determinant of insulin resistance (see Chapter 7: Fatty Acids and Mitochondria, Cell Growth and Injury: Broader Implications). This FFA effect would be evident in insulinresistant states (i.e., plasma [FFA] and [insulin] high), but inapparent or superfluous physiologically, e.g., in starvation (i.e., plasma [FFA] high, plasma [insulin] low). Additional information is clearly needed. Thus, the hypothesis that leptin-induced modulation of neuronal mitochondrial fatty acid oxidation participates in regulation of feeding behavior requires an understanding of how this mechanism might relate functionally to other effectors, including orexigenic and anorexigenic neurons in the arcuate nucleus and in second order nuclei with which they interact. While such an understanding is unavailable at present, the fact that pharmacological inhibition of CPT-I and mitochondrial fatty acid oxidation stimulates feeding behavior in the absence of primary changes in insulin and leptin84-86 suggests that this putative switch may lie downstream of the insulin and leptin receptors and upstream of the regulated release of melanocortin and agouti-related peptide/neuropeptide Y139. Furthermore, additional information is needed concerning leptin receptor signaling, which has been clearly linked both to PI3K124,149,150 (plausibly in part insulin-dependent) and JAK2 pathways150,151, the latter subject to negative Also, regulation by protein tyrosine phosphatase 1B (PTP 1B) 152 . 124-128 , although leptin receptor signaling activates fatty acid oxidation details of this process are incomplete, and a comparable effect has not been directly demonstrated in the arcuate nucleus. The potential for integration of novel enzymes and regulatory molecules pertaining to fatty acid metabolism will represent an ongoing challenge for the hypothesis. For example, a role of the recently


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identified brain CPT-I isoform noted above80 is uncertain. Second, an acetyl CoA carboxylase isoform (ACC2) that regulates fatty acid oxidation in extracranial tissues has been identified106; as its deletion increases feeding behavior rather than decreasing it153 while preventing obesity through increased fatty acid oxidation154, it would not appear to participate in the hypothesized hypothalamic regulatory process. Third, although methods differ, the effects of CPT-I inhibition on feeding behavior have been inconsistent155. Fourth, the potential orexigenic role of the hypothalamic cannabinoid system, and its downregulation by leptin156,157 awaits further elucidation. Finally, a novel fatty acid synthase inhibitor, C75, inhibits feeding behavior and increases cellular malonyl CoA concentration158; the latter should inhibit mitochondrial fatty acid oxidation, implying that feeding behavior and mitochondrial fatty acid oxidation are related directly rather than inversely as hypothesized here. However, as C75 increases mitochondrial fatty acid oxidation peripherally159, and in hypothalamus suppresses neuropeptide Y immunoreactivity (consistent with suppression of feeding behavior) but not that of fatty acid synthase160, C75’s hypothalamic effects and relationship to the present hypothesis await further clarification. The foregoing examples are illustrative of the ongoing examination of important questions and issues that will need to be addressed if the present hypothesis is to be rigorously tested. If validated, however, the hypothesis would represent significant progress toward elucidation of the mechanisms involved in appetite regulation and the possibility of their therapeutic modulation.

12.5. CYTOKINES AND THE BRAIN The interaction of cytokines with brain function has been a subject of intense interest, and it is clear that all brain cell populations — neuronal, glial, and vascular — express various cytokines and/or their receptors161-164. For purposes of this discussion, attention will be focussed on the proinflammatory cytokines, especially tumor necrosis interleukin-1 (IL-1), and IL-6, as these agents have been demonstrated to be intimately involved with, and to profoundly influence, systemic lipid and fatty acid metabolism50,165(also, see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin). It is important to recognize that the net effect of these agents, in brain and elsewhere, depends to a large extent on their interaction with various determinants of oxidative stress, including mitochondrial oxidation of fatty acids and

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regulation of redox-related genes, e.g., inducible nitric oxide synthase (iNOS). expression in adipocytes is increased in obesity, and may contribute to systemic insulin resistance by inhibiting insulin receptor signaling166. Adipocyte expression requires expression of aP2, the adipocyte form of cytosolic FABP167. Moreover, lipopolysaccharide, IL-1, and IL-6 also influence hepatocellular fatty acid metabolism and proliferation165,168,169 (also, see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin). These agents stimulate 165 lipolysis in adipose tissue , thereby increasing plasma [FFA] and as a result tissue uptake of fatty acids, including that by liver. In the hepatocyte, these cytokines also enhance glycolysis and de novo lipogenesis, thereby generating increased amounts of malonyl CoA, the physiological inhibitor of CPT-I and therefore of mitochondrial fatty acid oxidation and ketogenesis165,170. Thus, through their effects on adipose tissue and liver, proinflammatory cytokines augment hepatocellular uptake and de novo biosynthesis of fatty acids which, acting in concert with suppressed fatty acid oxidation, favor diversion of fatty acids into esterification pathways. These actions account for the cytokine-induced increases in TG biosynthesis, very low density lipoprotein (VLDL) TG secretion, and hyperlipidemia that regularly accompany systemic infections, including that by human immunodeficiency virus (HIV), and other inflammatory processes165. The systemic actions of these cytokines also may indirectly affect the brain by increasing plasma concentrations of FFA, brain uptake of which would therefore increase. Moreover, because fatty acids present in excess can override the malonyl CoA/CPT-I control point99-103, they may under some conditions drive augmented mitochondrial fatty acid oxidation, thus generating ROS and oxidative stress. Direct interaction of proinflammatory cytokines with the brain adds complexity to the situation. Thus, IL-1, and IL-6 are produced in and secreted by astrocytes161. It appears likely that their targeted cell populations are within the brain itself, although intracranial secretion of IL-6 may elevate its concentrations in sagittal sinus plasma171. Other glia express receptors for two of these agents, but neurons and astrocytes express receptors for all three161. In addition, systemically produced cytokines appear able to cross the blood brain barrier, and may also influence brain function via peripheral afferent nerve fibers163. Assuming that proinflammatory cytokines produced locally and systemically modulate brain cell function both directly and indirectly, and that their effects on intermediary metabolism generally parallel those on the hepatocyte, the resulting enhancement of glycolysis and inhibition of


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astrocyte mitochondrial fatty acid oxidation could be beneficial in terms of neuronal energetics, under conditions in which fatty acid abundance is not excessive. Such effects would contribute to the demonstrated proinflammatory cytokine enhancement of glycolytic fueling of the astrocyte citric acid cycle172,173, the protective effect of against excitotoxicity and oxidative stress174, and the enhancement of synaptic 175 strength by astrocyte-generated (also, see Chapter 14.5.2.1.: Long-Term Potentiation). Conversely, override of the CPT-I control point by high plasma FFA, as may occur, e.g., in HIV infection, obesity, insulin resistance, diabetes mellitus176, and smoking-related nicotine exposure177-181, would be deleterious. An additional element of considerable importance is the fact that the vascular, glial, and neuronal structures of the brain are richly endowed with constitutive endothelial and neuronal nitric oxide synthases (eNOS, nNOS), as well as iNOS163,182. Astrocytes in culture may release neurotoxic levels of NO in response to IL-1 and other proinflammatory cytokines183 or to the neurotrophic protein184. Furthermore, evidence based on in vivo and in vitro studies suggests that proinflammatory cytokines, e.g., IL-1, may play an important contributory role in the pathogenesis of neuronal dysfunction and death, including that associated with neurodegenerative, inflammatory, and ischemic processes162, as well as major depression185. These neurotoxic cytokine effects may be mediated in part by NO182 and/or by peroxynitrate the product of the reaction of NO with superoxide anion NO and inhibit the mitochondrial electron transport chain, the former at complex IV, i.e., cytochrome c oxidase186,187, and the latter at complexes I-III187. It is clear, however, that proinflammatory cytokines and NO are not invariably deleterious in their effects on neuronal function and survival, and may in fact be salutary depending on other factors that may be operative. Thus, NO suppresses apoptosis in several tissues, an action that appears to be mediated at least in part by NO inhibition of several members of the caspase family188,189; it may depend on the redox status of the target cell190193 and glycolytic substrate availability194. In addition, has recently been shown to be neuroprotective through its activation of glucose-6phosphate dehydrogenase and the pentose phosphate pathway, thus promoting generation of NADPH, a key cofactor in antioxidant defense195. Moreover, although signaling may be disruptive of the mitochondrial electron transport chain196 and may induce ceramidemediated apoptosis197-200, and IL-6 are essential for normal hepatic regeneration201-203, a circumstance in which the energy demands of cell proliferation, like those of neuronal activation, require optimal

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(glycolysis-fueled) mitochondrial energetics (see Chapter 8.4.1.: Effects of Proinflammatory Cytokines: vs. Insulin). Certain of these neuronal anti-apoptotic responses to oxidative stress and proinflammatory cytokines, as in peripheral cells204-206, reflect protective activation of by ROS and oxidative stress207-212. IL-1 and related cytokines also activate in promoting the survival of developing neurons213. The protective effect214,215 is mediated at least in part by transcriptional activation of genes related to redox balance, including inducible nitric oxide synthase (iNOS)216, cyclooxygenase 2 (COX2)217, Bcl2 and Bcl-xL215,218. Importantly, activation involves wg/Wnt-like signaling219, which also enhances cell energetics via phosphatidyl inositol-3-kinase (PI3K) and Akt/protein kinase B (Akt/PKB) (see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable; and Chapter 15.4.: Cerebral Metabolism and Insulin Signaling). Contrary to these interpretations, however, it has been suggested that effect is deleterious, based on the observation that salicylate220-222 induced neuroprotection is associated with suppression of , and 223 the latter’s association with amyloid toxicity . activation indeed As has been implicated in cell injury under certain conditions224. discussed below, however (see Chapter 14.5.2.2.: Excitotoxicity — Origins of Oxidative Stress), these conflicting interpretations can be reconciled by two important considerations: 1) salicylate is a classical protonophoric uncoupler of oxidative phosphorylation225-229; and 2) uncoupling agents decrease intramitochondrial formation of the ROS38-42 upon which activation depends230-232. Accordingly, it is likely that salicylate suppresses activation secondarily and indirectly as a consequence of the primary and beneficial effect of its uncoupling action, i.e., amelioration of intramitochondrial oxidative stress. Thus, proinflammatory cytokines may be viewed as double-edged swords with respect to neuronal function and survival. While they may increase intramitochondrial ROS generation and oxidative stress, this in turn may activate potent neuroprotective effects and determinants of survival, i.e., and wg/Wnt-like signaling. The overall outcome appears to depend to a large extent on other factors such as integrity of the electron transport chain, availability of oxygen, NO generation, fatty acid abundance (see Chapter 8.4.1. Effects of Proinflammatory Cytokines: vs. Insulin), and the effects of other redox determinants. The potential importance of this interplay in the pathogenesis of HIV-associated dementia will be considered briefly below


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(see Chapter 13.4.: Fatty Acids, Antioxidant Defense, HIV Dementia, and Cell Proliferation).

12.6. REFERENCES 1. D. Dwyer, Ed., Glucose metabolism in the brain, International Review of Neurobiology; Series Editors RJ Bradley, RA Harris, and P Jenner 51 (2002). 2. S. Vannucci, F. Maher, and I. Simpson, Glucose transporter proteins in brain: delivery of glucose to neurons and glia, Glia 21:2-21 (1997). 3. M. Schwartz, D. Figlewicz, D. Baskin, S. Woods, and D. Porte, Jr, Insulin in the brain: a hormonal regulator of energy balance, Endocrine Reviews 13:387-414 (1992). 4. P. Freychet, Insulin receptors and insulin actions in the nervous system, Diabetes/Metab Res Rev 16:390-392 (2000). 5. C. Park, Cognitive effects of insulin in the central nervous system, Neurosci Biobehav Rev 25:311-323 (2001). 6. A. Chaudhuri, Y. Kanjwal, P. Mohanty, et al., Insulin-induced vasodilatation of internal carotid artery, Metab Clin Exp 48:1470-1473 (1999). 7. S. Hasselbalch, G. Knudsen, C. Videbaek, et al., No effect of insulin on glucose bloodbrain barrier transport and cerebral metabolism in humans, Diabetes 48:1915-1921 (1999). 8. J. C. Bruning, D. Gautam, D. J. Burks, et al., Role of brain insulin receptor in control of body weight and reproduction, Science 289:2122-5 (2000). 9. C. Cheng, R. Reinhardt, W. Lee, G. Joncas, S. Patel, and C. Bondy, Insulin-like growth factor 1 regulates developing brain glucose metabolism, Proc Natl Acad Sci USA 97:10236-10241 (2000). 10. L. Stryer, Biochemistry, W.H. Freeman and Co, New York (1995). 11. E. McCabe, Microcompartmentation of energy metabolism at the outer mitochondrial membrane: role in diabetes mellitus and other diseases, J Bioenergetics Biomembranes 26:317-325 (1994). 12. G. Beutner, A. Rück, B. Riede, and D. Brdiczka, Complexes between hexokinase, mitochondrial porin and adenylate translocator in brain: regulation of hexokinase, oxidative phosphorylation and permeability transition pore, Biochem Soc Transactions 25:151-157 (1997). 13. N. Zamzami, C. Brenner, I. Marzo, S. Susin, and G. Kroemer, Subcellular and submitochondrial mode of action of Bcl-2-like oncoproteins, Oncogene 16:22652282 (1998). 14. I. Marzo, C. Brenner, N. Zamzami, et al., The permeability transition pore complex: a target for apoptosis regulation by caspases and Bcl-2-related proteins, J Exp Med 187:1261-1271 (1998). 15. V. V. Lemeshko, Model of the outer membrane potential generation by the inner membrane of mitochondria, Biophys J 82:684-92 (2002). 16. D. Gincel, S. Silberberg, and V. Shoshan-Barmatz, Modulation of the voltagedependent anion channel (VDAC) by glutamate, J Bioenerg Biomembranes 32:571583 (2000). 17. R. Behal, D. Buxton, J. Robertson, and M. Olson, Regulation of the pyruvate dehydrogenase multienzyme complex, Annu Rev Nutr 13:497-520 (1993). 18. O. Owen, A. Morgan, H. Kemp, J. Sullivan, M. Herrera, and G. Cahill, Jr, Brain metabolism during fasting, J Clin Invest 46:1589-1595 (1967).

242

Chapter 12

19. A. Smith, H. Satterthwaite, and L. Sokoloff, Induction of brain D(-)-betahydroxybytrate dehydrogenase activity by fasting, Science 163:79-81 (1969). 20. J. W. Pan, R. A. de Graaf, K. F. Petersen, G. I. Shulman, H. P. Hetherington, and D. L. Rothman, [2,4-13 C2 ]-beta-Hydroxybutyrate metabolism in human brain, J Cereb Blood Flow Metab 22:890-8 (2002). 21. S. Hasselbalch, G. Knudsen, J. Jakobsen, L. Hageman, S. Holm, and O. Paulson, Brain metabolism during short-term starvation in humans, J Cerebral Blood Flow Metab 14:125-31 (1994). 22. S. Hasselbalch, G. Knudsen, J. Jakobsen, L. Hageman, S. Holm, and O. Paulson, Blood-brain barrier permeability of glucose and ketone bodies during short-term starvation in humans, Am J Physiol 268:E1161-6 (1995). 23. S. Hasselbalch, P. Madsen, L. Hageman, et al., Changes in cerebral blood flow and carbohydrate metabolism during acute hyperketonemia, Am J Physiol 270:E746-51 (1996). 24. P. Crane, W. Pardridge, L. Braun, and W. Oldendorf, Two-day starvation does not alter the kinetics of blood-brain barrier transport and phosphorylation of glucose in rat brain, J Cerebral Blood Flow Metab 5:40-46 (1985). 25. C. Redies, L. Hoffer, C. Biel, et al., Generalized decrease in brain glucose metabolism during fasting in humans studied by PET, Am J Physiol 256:E805-E810 (1989). 26. G. Blomqvist, M. Alvarsson, V. Grill, et al., Effect of acute hyperketonemia on the cerebral uptake of ketone bodies in nondiabetic subjects and IDDM patients, Am J Physiol Endocrinol Metab 283:E20-8 (2002). 27. T. Moore, A. Lione, M. Sugden, and D. Regen, Beta-hydroxybutyrate transport in rat brain: development and dietary modulations, Am J Physiol 230:619-630 (1976). 28. L. Pellerin, G. Pellegri, J.-L. Martin, and P. Magistretti, Expression of monocarboxylate transporter mRNAs in mouse brain: support for a distinct role of lactate as an energy substrate for the neonatal vs adult brain, Proc Natl Aca Sci USA 95:3990-3995 (1998). 29. J. Tildon, M. McKenna, and J. Stevenson, Jr, Transport of 3-hydroxybutyrate by cultured rat brain astrocytes, Neurochem Res 19:1237-42 (1994). 30. M. Yudkoff, Y. Daikhin, I. Nissim, R. Grunstein, and I. Nissim, Effects of ketone bodies on astrocyte amino acid metabolism, J Neurochem 69:682-92 (1997). 31. G. Wilkinson, Clearance approaches in pharmacology, Pharmacol Rev 39: 1-47 (1987). 32. F. Palmieri, F. Bisaccia, L. Capobianco, et al., Mitochondrial metabolite transporters, Biochim Biophys Acta 1275:127-132 (1996). 33. A. Halestrap, Pyruvate and ketone-body transport across the mitochondrial membrane: Exchange properties, pH-dependence and mechanism of the carrier, Biochem J 172:377-387 (1978). 34. S. Pande, and R. Parvin, Pyruvate and acetoacetate transport in mitochondria: A reappraisal, J Biol Chem 253:1565-1573 (1978). 35. W. Zhang, S. Churchill, and P. Churchill, Developmental regulation of D-betahydroxybutyrate dehydrogenase in rat liver and brain, FEBS Lett 256:71-74 (1989). 36. L. Wojtczak, and P. Schönfeld, Effect of fatty acids on energy coupling processes in mitochondria, Biochim Biophys Acta 1183:41-57 (1993). 37. L. Svensson, S. Kilpeläinen, J. Hiltunen, and S. Alexson, Characterization and isolation of enzymes that hydrolyze short-chain acyl-CoA in rat-liver mitochondria, Eur J Biochem 239:526-31 (1996).


243

38. I. Reynolds, and T. Hastings, Glutamate induces the production of reactive oxygen species in cultured forebrain neurons following NMDA receptor activation, J Neurosci 15:3318-3327 (1995). 39. V. Skulachev, Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants, Quart Rev Biophys 29: 169-202 (1996). 40. A. Nègre-Salvayre, C. Hirtz, G. Carrera, et al., A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation, FASEB J 11:809-815 (1997). 41. A. Stout, H. Raphael, B. Kanterewicz, E. Klann, and I. Reynolds, Glutamate-induced neuron death requires mitochondrial calcium uptake, Nature Neurosci 1:366-373 (1998). 42. S. Korshunov, O. Korkina, E. Ruuge, V. Skulachev, and A. Starkov, Fatty acids as natural uncouplers preventing generation of O2- and H2O2 by mitochondria in the resting state, FEBS Letts 435:215-218 (1998). 43. K. Tieu, C. Perier, C. Caspersen, et al., D-beta-hydroxybutyrate rescues mitochondrial respiration and mitigates features of Parkinson disease., Journal of Clinical Investigation 112:892-901 (2003). 44. R. Ockner, N. Lysenko, N. Wu, and N. Bass, Hepatocyte growth inhibitors modulate mitochondrial and extramitochondrial fatty acid oxidation [Abstract], Hepatology 24:253A (1996). 45. M. Olson, S. Dennis, M. DeBuysere, and A. Padma, The regulation of pyruvate dehydrogenase in the isolated perfused rat heart, J Biol Chem 253:7369-7375 (1978). 46. M. Tisdale, Role of acetoacetyl-CoA synthetase in acetoacetate utilization by tumor cells, Cancer Biochem Biophys 7:101-107 (1984). 47. P. Garland, E. Newsholme, and P. Randle, Effects of fatty acids and ketone bodies and of alloxan-diabetes and starvation on pyruvate metabolism and on lactate/pyruvate and L-glycerol 3-phosphate/dihydroxy acetone phosphate concentration ratios in rat heart and rat diaphragm muscles, Biochem J 93:665-678 (1964). 48. J. Batenburg, and M. Olson, Regulation of pyruvate dehydrogenase by fatty acid in isolated rat liver mitochondria, J Biol Chem 251:1364-1370 (1976). 49. P. J. Randle, Regulatory interactions between lipids and carbohydrates: the glucose fatty acid cycle after 35 years, Diabetes Metab Rev 14:263-83 (1998). 50. G. Boden, and G. I. Shulman, Free fatty acids in obesity and type 2 diabetes: defining their role in the development of insulin resistance and beta-cell dysfunction, Eur J Clin Invest 32 Suppl 3:14-23 (2002). 51. T. Kawaguchi, K. Osatomi, H. Yamashita, T. Kabashima, and K. Uyeda, Mechanism for fatty acid “sparing” effect on glucose-induced transcription: regulation of carbohydrate-responsive element-binding protein by AMP-activated protein kinase, J Biol Chem 277:3829-3835 (2002). 52. R. Russell, 3d, G. Cline, P. Guthrie, G. Goodwin, G. Shulman, and H. Taegtmeyer, Regulation of exogenous and endogenous glucose metabolism by insulin and acetoacetate in the isolated working rat heart: A three tracer study of glycolysis, glycogen metabolism, and glucose oxidation, J Clin Invest 100:2892-2899 (1997). 53. Y. Kashiwaya, M. King, and R. Veech, Substrate signaling by insulin: a ketone bodies ratio mimics insulin action in heart, Am J Cardiol 80:50A-64A (1997). 54. R. Russell, 3d, and H. Taegtmeyer, Changes in citric acid cycle flux and anaplerosis antedate the functional decline in isolated rat hearts utilizing acetoacetate, J Clin Invest 87:384-390 (1991).

244

Chapter 12

55. R. Russell, 3d, and H. Taegtmeyer, Coenzyme A sequestration in rat hearts oxidizing ketone bodies, J Clin Invest 89:968-973 (1992). 56. Y. Izumi, K. Ishii, H. Katsuki, A. Benz, Zorumski, and CF, beta-Hydroxybutyrate fuels synaptic function during development: Histological and physiological evidence in rat hippocampal slices, J Clin Invest 101:1121-1132 (1998). 57. Y. Kashiwaya, T. Takeshima, N. Mori, K. Nakashima, K. Clarke, and R. Veech, Dbeta-hydroxybutyrate protects neurons in models of Alzheimer’s and Parkinson’s disease, Proc Natl Acad Sci USA 97:5440-5444 (2000). 58. Y. Yeh, Biosynthesis of phospholipids and sphingolipids from acetoacetate and glucose in different regions of developing brain in vivo, Journal of Neuroscience Research 11:383-394 (1984). 59. J. Edmond, Energy metabolism in developing brain cells, Can J Physiol Pharmacol 70:S118-29 (1992). 60. L. Roeder, S. Poduslo, and J. Tildon, Utilization of ketone bodies and glucose by established neural cell lines, Journal of Neuroscience Research 8:671-682 (1982). 61. A. Lapidot, and S. Haber, Effect of endogenous beta-hydroxybutyrate on brain glucose metabolism in fetuses of diabetic rabbits, studied by (13)C magnetic resonance spectroscopy, Brain Res Dev Brain Res 135:87-99 (2002). 62. X. Yang, L. Borg, and U. Eriksson, Metabolic alteration in neural tissue of rat embryos exposed to beta-hydroxybutyrate during organogenesis, Life Sci 62:293300 (1998). 63. G. Dhopeshwarkar, Uptake and transport of fatty acids into the breain and the role of the blood-brain barrier system, Adv Lipid Res 11:109-142 (1973). 64. R. Spector, Fatty acid transport through the blood-brain barrier, J Neurochem 50:639-643 (1988). 65. P. Robinson, J. Noronha, J. DeGeroge, L. Freed, T. Nariai, and S. Rapoport, A quantitative method for measuring regional in vivo fatty-acid incorporation into and turnover within brain phospholipids: review and critical analysis, Brain Res Rev 17:187-214 (1992). 66. J. Miller, J. Gnaedinger, and S. Rapoport, Utlization of plasma fatty acid in rat brain: distribution of [14C]palmitate between oxidative and synthetic pathways, J Neurochem 49:1507-1514 (1987). 67. J. Gnaedinger, J. Miller, C. Latker, and S. Rapoport, Cerebral metabolism of plasma [14C]palmitate in awake, adult rat: subcellular localization, Neurochem Res 13:21-29 (1988). 68. S. Rapoport, In vivo fatty acid incorporation into brain phospholipids in relation to signal transduction and membrane remodeling, Neurochemical Research 24:1403-15 (1999A). 69. J. Glatz, J. Luiken, F. van Nieuwenhoven, and G. Van der Vusse, Molecular mechanism of cellular uptake and intracellular translocation of fatty acids, Prostaglandins Leukotrienes and Essential Fatty Acids 57:3-9 (1997). 70. A. Kimes, D. Sweeney, E. London, and S. Rapoport, Palmitate incorporation into different brain regions in the awake rat, Brain Res 274:291-301 (1983). 71. M. Chang, T. Arai, L. Freed, et al., Brain incorporation of [1-11C]arachidonate in normocapnic and hypercapnic monkeys measured with positron emission tomography, Brain Res 755:74-83 (1997B). 72. I. Goldberg, D. Soprano, M. Wyatt, T. Vanni, T. Kirchgessner, and M. Schotz, Localization of lipoprotein lipase mRNA in selected rat tissues, J Lipid Res 30:15691577 (1989).


245

73. D. Bessesen, C. Richards, J. Etienne, J. Goers, and R. Eckel, Spinal cord of the rat contains more lipoprotein lipase than other brain regions, J Lipid Res 34:229-238 (1993). 74. D. Purdon, T. Arai, and S. Rapoport, No evidence for direct incorporation of esterified palmitic acid from plasma into brain lipids of awake adult rat, J Lipid Res 38:526-30 (1997). 75. D. Bernlohr, M. Simpson, A. Hertzel, and L. Banaszak, Intracellular lipid-binding proteins and their genes, Ann Rev Nutr 17:277-303 (1997). 76. T. Fujino, and T. Yamamoto, Cloning and functional expression of a novel longchain acyl-CoA synthetase expressed in brain, J Biochem 111:197-203 (1992). 77. T. Arai, S. Wakabayashi, M. Channing, et al., Incorporation of [1-carbon11]palmitate in monkey brain using PET, J Nuclear Med 36:2261-2267 (1995). 78. C. Blázquez, Sánchez, C, G. Velasco, and M. Guzmán, Role of carnitine palmitoyltransferase I in the control of ketogenesis in primary cultures of rat astrocytes, J Neurochem 71:1597-1606 (1998). 79. N. Brown, J. Hill, V. Esser, et al., Mouse white adipocytes and 3T3-L1 cells display an anomalous pattern of carnitine palmitoyltransferase (CPT) I isoform expression during differentiation: Inter-tissue and inter-species expression of CPT I and CPT II enzymes, Biochem J 327:225-231 (1997). 80. N. Price, F. van der Leij, V. Jackson, et al., A novel brain-expressed protein related to carnitine palmitoyltransferase I, Genomics 80:433-42 (2002). 81. M. Chang, S. Wakabayashi, and J. Bell, The effect of methyl palmoxirate on incorporation of [U-14C]palmitate into rat brain, Neurochem Res 19:1217-1223 (1994). 82. L. Freed, S. Wakabayashi, J. Bell, and S. Rapoport, Effect of inhibition of betaoxidation on incorporation of [U-14C]palmitate and [1-14C]arachidonate into brain lipids, Brain Res 645:41-48 (1994). 83. M. Chang, E. Grange, O. Rabin, and J. Bell, Incorporation of [U-14C]palmitate into rat brain: effect of an inhibitor of beta-oxidation, J Lipid Res 38:295-300 (1997A). 84. C. Horn, and M. Friedman, Methyl palmoxirate increases eating behavior and brain Fos-like immunoreactivity in rats, Brain Res 781:8-14 (1998). 85. M. I. Friedman, R. B. Harris, H. Ji, I. Ramirez, and M. G. Tordoff, Fatty acid oxidation affects food intake by altering hepatic energy status, Am J Physiol 276:R1046-53 (1999). 86. A. Kahler, M. Zimmermann, and W. Langhans, Suppression of hepatic fatty acid oxidation and food intake in men, Nutrition 15:819-28 (1999). 87. N. Kawamura, and Y. Kishimoto, Characterization of water-soluble products of palmitic acid beta-oxidation by a rat brain preparation, J Neurochem 36:1786-1791 (1981). 88. N. Auestad, R. Korsak, J. Morrow, and J. Edmond, Fatty acid oxidation and ketogenesis by astrocytes in primary culture, J Neurochem 56:1376-1386 (1991). 89. C. Blázquez, C. Sánchez, A. Daza, I. Galve-Roperh, and M. Guzmán, The stimulation of ketogenesis by cannabinoids in cultured astrocytes defines carnitine palmitoyltransferase I as a new ceramide-activated enzyme, J Neurochem 72:17591768 (1999A). 90. J. Bourre, and M. Piciotti, Alterations in eighteen-carbon saturated, monounsaturated and plolyunsatruated fatty acid peroxisomal oxidation in mouse brain during development and aging, Biochem Molec Biol Intl 41:461-468 (1997).

246

Chapter 12

91. P. Burra, M. Dam, F. Chierichetti, et al., 18F-fluorodeoxyglucose positron emission tomography study of brain metabolism in cirrhosis: effect of liver transplantation, Transplant Proc 31:418-420 (1999). 92. G. Sarna, M. Bradbury, J. Cremer, J. Lai, and H. Teal, Brain metabolism and specific transport at the blood-brain barrier after portocaval anastomosis in the rat, Brain Res 160:69-83 (1979). 93. S. Ponchaut, and K. Veitch, Valproate and mitochondria, Biochem Pharmacol 46:199-204 (1993). 94. T. Cullingford, K. Bhakoo, S. Peuchen, C. Dolphin, R. Patel, and J. Clark, Distribution of mRNAs encoding the peroxisome proliferator-activated receptor alpha, beta, and gamma and the retinoid X receptor alpha, beta, and gamma in rat central nervous system, J Neurochem 70:1366-1375 (1998). 95. J. Granneman, R. Skoff, and X. Yang, Member of the peroxisome proliferatoractivated receptor family of transcription factors is differentially expressed by oligodendrocytes, J Neurosci Res 51:563-573 (1998). 96. N. Chattopadhyay, D. Singh, O. Heese, et al., Expression of peroxisome proliferatoractivated receptors (PPARs) in human astrocytic cells: PPARgamma agonists as inducers of apoptosis, J Neurosci Res 61:67-74 (2000). 97. M. C. Sugden, K. Bulmer, G. F. Gibbons, B. L. Knight, and M. J. Holness, Peroxisome-proliferator-activated receptor-alpha (PPARalpha) deficiency leads to dysregulation of hepatic lipid and carbohydrate metabolism by fatty acids and insulin, Biochem J 364:361-8 (2002). 98. R. Ockner, Apoptosis and liver diseases: Recent concepts of mechanism and significance, J Gastroenterol Hepatol 16:248-260 (2001). 99. S. Mills, D. Foster, and J. McGarry, Interaction of malonyl-CoA and related compounds with mitochondria from different rat tissues:Relationship between ligand binding and inhibition of carnitine palmitoyltransferase I, Biochem J 214:83-91 (1983). 100. L. Drynan, P. Quant, and V. Zammit, Flux control exerted by mitochondrial outer membrane carnitine palmitoyltransferase over beta-oxidation, ketogenesis and tricarboxylic acid cycle activity in hepatocytes isolated from rats in different metabolic states, Biochem J 317:791-795 (1996A). 101. L. Drynan, P. Quant, and V. Zammit, The role of changes in the sensitivity of hepatic mitochondrial overt carnitine palmitoyltransferase in determining the onset of the ketosis of starvation in the rat, Biochem J 318:767-770 (1996B). 102. J. Sleboda, K. Risan, O. Spydevold, and J. Bremer, Short-term regulation of carnitine palmitoyltransferase I in cultured rat hepatocytes: spontaneous inactivation and reactivation by fatty acids, Biochim Biophys Acta 1436:541-549 (1999). 103. J. McGarry, and N. Brown, Reconstitution of purified, active and malonyl-CoAsensitive rat liver carnitine palmitoyltransferase I: relationship between membrane environment and malonyl-CoA sensitivity, Biochem J 349:179-187 (2000). 104. W. Ong, C. Hu, Y. Soh, T. Lim, P. Pentchev, and S. Patel, Neuronal localization of sterol regulatory element binding protein-1 in the rodent and primate brain: A light and electron microscopic immunocytochemical study, Neurosci 97:143-153 (2000). 105. J. Vance, C. De, EP, R. Campenot, and D. Vance, Role of axons in membrane phospholipid synthesis in rat sympathetic neurons, Neurobiol Aging 16:493-498 (1995). 106. L. Abu-Elheiga, W. R. Brinkley, L. Zhong, S. S. Chirala, G. Woldegiorgis, and S. J. Wakil, The subcellular localization of acetyl-CoA carboxylase 2, Proc Natl Acad Sci U S A 97:1444-9 (2000).


247

107. J. Sakamoto, R. Barr, K. Kavanagh, and G. Lopaschuk, Contribution of malonylCoA decarboxylase to the high fatty acid oxidation rates seen in the diabetic heart, Am J Physiol Heart Circ Physiol 278:H1196-H1204 (2000). 108. J. Alexander, A. Snyder, and J. Tonsgard, Omega-oxidation of monocarboxylic acids in rat brain, Neurochem Res 23:227-233 (1998). 109. J. Bylund, C. Zhang, and D. R. Harder, Identification of a novel cytochrome P450, CYP4X1, with unique localization specific to the brain, Biochem Biophys Res Commun 296:677-84 (2002). 110. R. Kaikaus, W. Chan, N. Lysenko, R. Ray, P. Ortiz de Montellano, and N. Bass, Induction of peroxisomal fatty acid beta-oxidation and liver fatty acid binding protein by peroxisome proliferators: Mediation via the cytochrome P-450 4A1 omegahydroxylase pathway, J Biol Chem 268:9592-9603 (1993). 111. D. Richard, S. Clavel, Q. Huang, D. Sanchis, and D. Ricquier, Uncoupling protein 2 in the brain: distribution and function, Biochem Soc Trans 29:812-817 (2001). 112. T. Horvath, C. Warden, M. Hajos, A. Lombardi, F. Goglia, and S. Diano, Brain uncoupling protein 2: uncoupled neuronal mitochondria predict thermal synapses in homeostatic centers, J Neurosci 19:10417-10427 (1999). 113. S. Diano, H. Urbanski, B. Horvath, et al., Mitochondrial uncoupling protein 2 (UCP2) in the nonhuman primate brain and pituitary, Endocrinol 141:4226-4238 (2000). 114. T. L. Horvath, S. Diano, and C. Barns table, Mitochondrial uncoupling protein 2 in the central nervous system: neuromodulator and neuroprotector, Biochem Pharmacol 65:1917-21 (2003). 115. O. Boss, P. Muzzin, and J.-P. Glacobino, The uncoupling proteins, a review, European Journal of Endocrinology 139:1-9 (1998). 116. K. Chavin, S. Yang, H. Lin, et al., Obesity induces expression of uncoupling protein-2 in hepatocytes and promotes liver ATP depletion, J Biol Chem 26274:5692-5700 (1999). 117. M. Jaburek, M. Varecha, R. Gimeno, et al., Transport function and regulation of mitochondrial uncoupling proteins 2 and 3, J Biol Chem 274:26003-7 (1999). 118. K. Echtay, D. Roussel, J. St-Pierre, et al., Superoxide activates mitochondrial uncoupling proteins, Nature 415:96-99 (2002). 119. J. K. Young, Anatomical relationship between specialized astrocytes and leptinsensitive neurones, J Anat 201:85-90 (2002). 120. D. Yablonskiy, J. Ackerman, and M. Raichle, Coupling between changes in human brain temperature and oxidative metabolism during prolonged visual stimulation, Proc Natl Acad Sci USA 97:7603-7608 (2000). 121. V. B. Hinderling, P. Schrauwen, W. Langhans, and M. S. Westerterp-Plantenga, The effect of etomoxir on 24-h substrate oxidation and satiety in humans, Am J Clin Nutr 76:141-7 (2002). 122. R. S. Ahima, and J. S. Flier, Leptin, Annu Rev Physiol 62:413-37 (2000). 123. A. Z. Zhao, M. M. Shinohara, D. Huang, et al., Leptin induces insulin-like signaling that antagonizes cAMP elevation by glucagon in hepatocytes, J Biol Chem 275:11348-54 (2000). 124. D. M. Muoio, G. L. Dohm, E. B. Tapscott, and R. A. Coleman, Leptin opposes insulin’s effects on fatty acid partitioning in muscles isolated from obese ob/ob mice, Am J Physiol 276:E913-21 (1999). 125. L. L. Atkinson, M. A. Fischer, and G. D. Lopaschuk, Leptin activates cardiac fatty acid oxidation independent of changes in the AMP-activated protein kinase-acetylCoA carboxylase-malonyl-CoA axis, J Biol Chem 277:29424-30 (2002).

248

Chapter 12

126. Y. Minokoshi, Y. Kim, O. Peroni, et al., Leptin stimulates fatty-acid oxidation by activating AMP-activated protein kinase, Nature 415:339-343 (2002). 127. G. Steinberg, A. Bonen, and D. Dyck, Fatty acid oxidation and triacylglycerol hydrolysis are enhanced after chronic leptin treatment in rats, Am J Physiol Endocrinol Metab 282:E593-E600 (2002A). 128. G. R. Steinberg, J. W. Rush, and D. J. Dyck, AMPK expression and phosphorylation are increased in rodent muscle after chronic leptin treatment, Am J Physiol Endocrinol Metab 284:E648-54 (2003). 129. G. Hynes, and P. Jones, Leptin and its role in lipid metabolism, Curr Opin Lipidol 12:321-327 (2001). 130. S. Yamagishi, D. Edelstein, X. Du, Y. Kaneda, M. Guzman, and M. Brownlee, Leptin induces mitochondrial superoxide production and monocyte chemoattractant protein-1 expression in aortic endothelial cells by increasing fatty acid oxidation via protein kinase A, J Biol Chem 276:25096-25100 (2001). 131. R. L. Dobbins, L. S. Szczepaniak, W. Zhang, and J. D. McGarry, Chemical sympathectomy alters regulation of body weight during prolonged ICV leptin infusion, Am J Physiol Endocrinol Metab 284:E778-87 (2003). 132. D. Spanswick, M. A. Smith, V. E. Groppi, S. D. Logan, and M. L. Ashford, Leptin inhibits hypothalamic neurons by activation of ATP-sensitive potassium channels, Nature 390:521-5 (1997). 133. T. J. Kieffer, R. S. Heller, C. A. Leech, G. G. Holz, and J. F. Habener, Leptin suppression of insulin secretion by the activation of ATP-sensitive K+ channels in pancreatic beta-cells, Diabetes 46:1087-93 (1997). 134. G. Sonnenberg, G. Krakower, R. Hoffmann, D. Maas, M. Hennes, and A. Kissebah, Plasma leptin concentrations during extended fasting and graded glucose infusions: relationships with changes in glucose, insulin, and FFA, J Clin Endocrinol Metab 86:4895-4900 (2001). 135. J. W. Kolaczynski, M. R. Nyce, R. V. Considine, et al., Acute and chronic effects of insulin on leptin production in humans: Studies in vivo and in vitro, Diabetes 45:699-701 (1996A). 136. J. W. Kolaczynski, R. V. Considine, J. Ohannesian, et al., Responses of leptin to short-term fasting and refeeding in humans: a link with ketogenesis but not ketones themselves, Diabetes 45:1511-5 (1996B). 137. G. Boden, X. Chen, J. W. Kolaczynski, and M. Polansky, Effects of prolonged hyperinsulinemia on serum leptin in normal human subjects, J Clin Invest 100:110713 (1997). 138. S. C. Woods, M. W. Schwartz, D. G. Baskin, and R. J. Seeley, Food intake and the regulation of body weight, Annu Rev Psychol 51:255-77 (2000). 139. D. E. Cummings, and M. W. Schwartz, Genetics and pathophysiology of human obesity, Annu Rev Med 54:453-71 (2003). 140. G. Boden, X. Chen, M. Mozzoli, and I. Ryan, Effect of fasting on serum leptin in normal human subjects, J Clin Endocrinol Metab 81:3419-23 (1996). 141. R. Saladin, P. De Vos, M. Guerre-Millo, et al., Transient increase in obese gene expression after food intake or insulin administration, Nature 377:527-9 (1995). 142. B. Laferrere, A. Caixas, S. K. Fried, C. Bashore, J. Kim, and F. X. Pi-Sunyer, A pulse of insulin and dexamethasone stimulates serum leptin in fasting human subjects, Eur J Endocrinol 146:839-45 (2002). 143. M. Shimabukuro, K. Koyama, G. Chen, et al., Direct antidiabetic effect of leptin through triglyceride depletion of tissues, Proc Natl Acad Sci U S A 94:4637-41 (1997).


249

144. G. R. Steinberg, D. J. Dyck, J. Calles-Escandon, et al., Chronic leptin administration decreases fatty acid uptake and fatty acid transporters in rat skeletal muscle, J Biol Chem 277:8854-60 (2002B). 145. J. D. McGarry, Banting lecture 2001: dysregulation of fatty acid metabolism in the etiology of type 2 diabetes, Diabetes 51:7-18 (2002). 146. E. L. Air, M. Z. Strowski, S. C. Benoit, et al., Small molecule insulin mimetics reduce food intake and body weight and prevent development of obesity, Nat Med 8:179-83 (2002). 147. R. S. Ahima, D. Prabakaran, C. Mantzoros, et al., Role of leptin in the neuroendocrine response to fasting, Nature 382:250-2 (1996). 148. J. L. Chan, K. Heist, A. M. DePaoli, J. D. Veldhuis, and C. S. Mantzoros, The role of falling leptin levels in the neuroendocrine and metabolic adaptation to short-term starvation in healthy men, J Clin Invest 111:1409-21 (2003). 149. K. D. Niswender, G. J. Morton, W. H. Stearns, C. J. Rhodes, M. G. Myers, Jr., and M. W. Schwartz, Key enzyme in leptin-induced anorexia, Nature 413:794-5 (2001). 150. J. Harvey, and M. L. Ashford, Leptin in the CNS: much more than a satiety signal, Neuropharmacology 44:845-54 (2003). 151. C. Bjorbaek, S. Uotani, B. da Silva, and J. S. Flier, Divergent signaling capacities of the long and short isoforms of the leptin receptor, J Biol Chem 272:32686-95 (1997). 152. J. M. Zabolotny, K. K. Bence-Hanulec, A. Stricker-Krongrad, et al., PTP1B regulates leptin signal transduction in vivo, Dev Cell 2:489-95 (2002). 153. L. Abu-Elheiga, M. M. Matzuk, K. A. Abo-Hashema, and S. J. Wakil, Continuous fatty acid oxidation and reduced fat storage in mice lacking acetyl-CoA carboxylase 2, Science 291:2613-6 (2001). 154. L. Abu-Elheiga, W. Oh, P. Kordari, and S. J. Wakil, Acetyl-CoA carboxylase 2 mutant mice are protected against obesity and diabetes induced by high-fat/highcarbohydrate diets., Proc Natl Aca Sci USA 100:10207-10212 (2003). 155. S. Obici, Z. Feng, A. Arduini, R. Conti, and L. Rossetti, Inhibition of hypothalamic carnitine palmitoyltransferase-1 decreases food intake and glucose production, Nat Med 9:756-61 (2003). 156. V. Di Marzo, S. K. Goparaju, L. Wang, et al., Leptin-regulated endocannabinoids are involved in maintaining food intake, Nature 410:822-5 (2001). 157. D. Cota, G. Marsicano, M. Tschop, et al., The endogenous cannabinoid system affects energy balance via central orexigenic drive and peripheral lipogenesis, J Clin Invest 112:423-31 (2003). 158. M. Kumar, T. Shimokawa, T. Nagy, and M. Lane, Differential effects of a centrally acting fatty acid synthase inhibitor in lean and obese mice, Proc Natl Acad Sci U S A 99:1921-1925 (2002). 159. J. N. Thupari, L. E. Landree, G. V. Ronnett, and F. P. Kuhajda, C75 increases peripheral energy utilization and fatty acid oxidation in diet-induced obesity, Proc Natl Acad Sci U S A 99:9498-502 (2002). 160. E. K. Kim, I. Miller, L. E. Landree, et al., Expression of FAS within hypothalamic neurons: a model for decreased food intake after C75 treatment, Am J Physiol Endocrinol Metab 283:E867-79 (2002). 161. E. Sternberg, Neural-immune interactions in health and disease, J Clin Invest 100:2641-2647 (1997). 162. N. Rothwell, S. Allan, and S. Toulmond, The role of interleukin 1 in acute neurodegeneration and stroke: pathophysiological and therapeutic implications, J Clin Invest 100:2648-2652 (1997).

250

Chapter 12

163. J. Licinio, and M.-L. Wong, Pathways and mechanisms for cytokine signaling of the central nervous system, J Clin Invest 100:2941-2947 (1997). 164. J. Raber, O. Sorg, T. Horn, et al., Inflammatory cytokines: putative regulators of neuronal and neuro-endocrine function, Brain Res Rev 26:320-326 (1998). 165. M. Navasa, K. Feingold, and C. Grunfeld, Effects of endotoxin and cytokines on hepatic lipid metabolism, Prog Liver Dis 15:147-170 (1997). 166. G. Hotamisligil, P. Peraldi, A. Budavari, R. Ellis, M. White, and B. Spiegelman, IRS-1-mediated inhibition of insulin receptor tyrosine kinase activity in TNF-alphaand obesity-induced insulin resistance, Science 271:665-8 (1996A). 167. G. Hotamisligil, R. Johnson, R. Distel, R. Ellis, V. Papaioannou, and B. Spiegelman, Uncoupling of obesity from insulin resistance through a targeted mutation in aP2, the adipocyte fatty acid binding protein, Science 274:1377-1379 (1996B). 168. C. Grunfeld, C. Dinarello, and K. Feingold, Tumor necrosis factor-alpha, interleukin-1, and interferon alpha stimulate triglyceride synthesis in HepG2 cells, Metabolism 40:894-898 (1991). 169. E. Vara, J. Arias-Diaz, J. Torres-Melero, C. Garcia, J. Rodriguez, and J. Balibrea, Effect of different sepsis-related cytokines on lipid synthesis by isolated hepatocytes, Hepatology 20:924-931 (1994). 170. M. Beylot, H. Vidal, G. Mithieux, M. Odeon, and C. Martin, Inhibition of hepatic ketogenesis by tumor necrosis factor-alpha in rats, Am J Physiol 263:E897-E902 (1992). 171. L. Romero, I. Kakucska, R. Lechan, and S. Reichlin, Interleukin-6 (IL-6) is secreted from the brain after intracerebroventricular injection of IL-1 beta in rats, Am J Physiol 270:R518-R524 (1996). 172. N. Yu, J. Martin, N. Stella, and P. Magistretti, Arachidonic acid stimulates glucose uptake in cerebral cortical astrocytes, Proc Natl Acad Sci USA 90:4042-4046 (1993). 173. N. Yu, D. Maciejewski-Lenoir, F. Bloom, and P. Magistretti, Tumor necrosis factoralpha and interleukin-1 alpha enhance glucose utilization by astrocytes: involvement of phospholipase A2, Molec Pharmacol 48:550-558 (1995). 174. B. Cheng, S. Christakos, and M. Mattson, Tumor necrosis factors protect neurons against metabolic-excitotoxic insults and promote maintenance of calcium homeostasis, Neuron 12:139-153 (1994). 175. E. Beattie, D. Stellwagen, W. Morishita, et al., Control of synaptic strength by glial TNFalpha, Science 295:2282-2285 (2002). 176. H. Ginsberg, Insulin resistance and cardiovascular disease, J Clin Invest 106:453458 (2000). 177. P. Cryer, M. Haymond, J. Santiago, and S. Shah, Norepinephrine and epinephrine release and adrenergic mediation of smoking-associated hemodynamic and metabolic events, New England Journal of Medicine 295:573-7 (1976). 178. M. Hellerstein, N. Benowitz, R. Neese, et al., Effects of cigarette smoking and its cessation on lipid metabolism and energy expenditure in heavy smokers, Journal of Clinical Investigation 93:265-72 (1994). 179. K. Fattinger, D. Verotta, and N. Benowitz, Pharmacodynamics of acute tolerance to multiple nicotinic effects in humans, J Pharmacol Exp Ther 281:1238-46 (1997). 180. J. Rincón, A. Krook, D. Galuska, H. Wallberg-Henriksson, and J. Zierath, Altered skeletal muscle glucose transport and blood lipid levels in habitual cigarette smokers, Clin Physiol 19:135-142 (1999).


251

181. J. Manson, U. Ajani, S. Liu, D. Nathan, and C. Hennekens, A prospective study of cigarette smoking and the incidence of diabetes mellitus among US male physicians, Am J Med 109:538-542 (2000). 182. K. Christopherson, and D. Bredt, Nitric oxide in excitable tissues: physiological roles and disease, J Clin Invest 100:2424-2429 (1997). 183. C. Chao, S. Hu, W. Sheng, D. Bu, M. Bukrinsky, and P. Peterson, Cytokinestimulated astrocytes damage human neurons via a nitric oxide mechanism, Glia 16:276-284 (1996). 184. J. Hu, A. Ferreira, and L. Van Eldik, S100beta induces neuronal cell death through nitric oxide release from astrocytes, J Neurochem 69:2294-2301 (1997). 185. M. Maes, and R. Smith, Fatty acids, cytokines, and major depression, Biol Psych 43:313-314 (1998). 186. V. Borutaité, and G. Brown, Rapid reduction of nitric oxide by mitochondria and reversible inhibition of mitochondrial respiration by nitric oxide, Biochem J 315:295-299 (1996). 187. I. Lizasoain, M. Moro, R. Knowles, V. Darley-Usmar, and S. Moncada, Nitric oxide and peroxynitrite exert distinct effects on mitochondrial respiration which are differentially blocked by glutathione or glucose, Biochem J 314:877-880 (1996). 188. J. Li, T. Billiar, R. Talanian, and Y. Kim, Nitric oxide reversibly inhibits seven members of the caspase family via S-nitrosylation, Biochem Biophys Res Commun 240:419-424 (1997). 189. Y. Kim, R. Talanian, and T. Billiar, Nitric oxide inhibits apoptosis by preventing increases in caspase-3-like activity via two distinct mechanisms, J Biol Chem 272:31138-31148 (1997). 190. S. Lipton, Y. Choi, Z. Pan, et al., A redox-based mechanism for the neuroprotective and neurodestructive effects of nitric oxide and related nitroso-compounds, Nature 364:626-632 (1993). 191. D. Wink, I. Hanbauer, M. Krishna, et al., Nitric oxide protects against cellular damage and cytotoxicity from reactive oxygen species, Proc Natl Acad Sci USA 90:9813-9817 (1993). 192. J. Bolaños, A. Almeida, E. Fernández, et al., Potential mechanisms for nitric oxidemediated impairment of brain mitochondrial energy metabolism, Biochem Soc Transact 25:944-949 (1997A). 193. J. Bolaños, A. Almeida, V. Stewart, et al., Nitric oxide-mediated mitochondrial damage in the brain: mechanisms and implications for neurodegenerative diseases, J Neurochem 68:2227-2240 (1997B). 194. B. Beltrán, A. Mathur, M. Duchen, J. Erusalimsky, and S. Moncada, The effect of nitric oxide on cell respiration: A key to understanding its role in cell survival or death, Proc Natl Acad Sci USA 97:14602-14607 (2000). 195. P. García-Nogales, A. Almeida, and J. Bolaños, Peroxynitrite protects neurons against nitric oxide-mediated apoptosis, A key role for glucose-6-phosphate dehydrogenase in neuroprotection., Journal of Biological Chemistry 278:864-874 (2003). 196. K. Schulze-Osthoff, A. Bakker, B. Vanhaesebroeck, R. Beyaert, W. Jacob, and W. Fiers, Cytotoxic activity of tumor necrosis factor is mediated by early damage of mitochondrial functions: Evidence for the involvement of mitochondrial radical generation, J Biol Chem 267:5317-5323 (1992). 197. L. Obeid, C. Linardic, L. Karolak, and Y. Hannun, Programmed cell death induced by ceramide, Science 259:1769-1771 (1993).

252

Chapter 12

198. R. Kolesnick, and M. Krönke, Regulation of ceramide production and apoptosis, Annu Rev Physiol 60:643-665 (1998). 199. M. Burow, C. Weldon, B. Collins-Burow, et al., Cross-talk between phosphatidylinositol 3-kinase and sphingomyelinase pathways as a mechanism for cell survival/death decisions, J Biol Chem 275:9628-9635 (2000). 200. T. Lin, L. Genestier, M. Pinkoski, et al., Role of acidic sphingomyelinase in Fas/CD95-mediated cell death, J Biol Chem 275:8657-8663 (2000). 201. P. Akerman, P. Cote, S. Yang, et al., Antibodies to tumor necrosis factor-alpha inhibit liver regeneration after partial hepatectomy, Am J Physiol 263:G579-G585 (1992). 202. D. Cressman, L. Greenbaum, R. DeAngelis, et al., Liver failure and defective hepatocyte regeneration in interleukin-6-deficient mice, Science 274:1379-1383 (1996). 203. Y. Yamada, I. Kinillova, J. Reschou, and N. Fausto, Initiation of tumor growth by tumor necrosis factor: deficient liver regeneration in mice lacking type I tumor necrosis factor receptor, PNAS 94:1441-1446 (1997). 204. A. Beg, and D. Baltimore, An essential role for NK-kappaB in preventing TNFalpha-induced cell death, Science 274:782-784 (1996). 205. D. Van Antwerp, S. Martin, T. Kafri, D. Green, and I. Verma, Suppression of TNFalpha-induced apoptosis by NF-kappaB, Science 274:787-789 (1996). 206. C.-Y. Wang, M. Mayo, and A. Baldwin, Jr, TNF- and cancer therapy-induced apoptosis: potentiation by inhibition of NF-kappaB, Science 274:784-787 (1996). 207. S. Barger, D. Hörsier, K. Furukawa, Y. Goodman, J. Krieglstein, and M. Mattson, Tumor necrosis factors alpha and beta protect neurons against amyloid beta-peptide toxicity: evidence for involvement of a kappa B-binding factor and attenuation of peroxide and Ca2+ accumulation, Proc Natl Acad Sci USA 92:9328-9332 (1995). 208. C. Kaltschmidt, B. Kaltschmidt, and P. Baeuerle, Stimulation of ionotropic glutamate receptors activates transcription factor NF-kappa B in primary neurons, Proc Natl Acad Sci USA 92:9618-9822 (1995). 209. B. Kaltschmidt, M. Uherek, B. Volk, P. Baeuerle, and C. Kaltschmidt, Transcription factor NF-kappa B is activated in primary neurons by amyloid beta peptides and in neurons surrounding early plaques from patients with Alzheimer disease, Proc Natl Acad Sci USA 94:2642-2647 (1997). 210. M. Mattson, Y. Goodman, H. Luo, W. Fu, Furukawa, and K, Activation of NF-kappa B protects hippocampal neurons against oxidative stress-induced apoptosis: evidence for induction of manganese superoxide dismutase and suppression of peroxynitrite production and protein tyrosine nitration, J Neurosci Res 49:681-697 (1997). 211. A. Migheli, R. Piva, C. Atzori, D. Troost, and D. Schiffer, c-Jun, JNK/SAPK kinases and transcription factor NF-kappa B are selectively activated in astrocytes, but not motor neurons in amyotrophic lateral sclerosis, J Neuropathol Exp Neurol 56:13141322 (1997). 212. S. Lipton, Janus faces of NF-kappa B: neurodestruction versus neuroprotection, Nature Med 3:20-22 (1997). 213. G. Middleton, M. Hamanoue, Y. Enokido, et al., Cytokine-induced nuclear factor kappa B activation promotes the survival of developing neurons, J Cell Biol 148:325-332 (2000). 214. P. García-Nogales, A. Almeida, E. Fernández, J. Medina, and J. Bolaños, Induction of glucose-6-phosphate dehydrogenase by lipopolysaccharide contributes to preventing nitric oxide-mediated glutathione depletion in cultured rat astrocytes, J Neurochem 72:1750-8 (1999).


253

215. J. Tan, T. Town, A. Placzek, A. Kundtz, H. Yu, and M. Mullan, Bcl-X(L) inhibits apoptosis and necrosis produced by Alzheimer’s beta-amyloid1-40 peptide in PC 12 cells, Neuroscience Letters 272:5-8 (1999). 216. T. Vos, H. Van Goor, L. Tuyt, et al., Expression of inducible nitric oxide synthase in endotoxemic rat hepatocytes is dependent on the cellular glutathione status, Hepatology 29:421-442 (1999). 217. K. Yamamoto, T. Arakawa, N. Ueda, and S. Yamamoto, Transcriptional roles of nuclear factor kappa B and nuclear factor-interleukin-6 in the tumor necrosis factor alpha-dependent induction of cyclooxygenase-2 in MC3T3-E1 cells, J Biol Chem 270:31315-31320 (1995). 218. M. Tamatani, Y. Che, H. Matsuzaki, et al., Tumor necrosis factor induces Bcl-2 and Bcl-x expression through NFkappaB activation in primary hippocampal neurons, J Biol Chem 274:8531-8538 (1999). 219. O. Ozes, L. Mayo, J. Gustin, S. Pfeffer, L. Pfeffer, and D. Donner, NF-kappaB activation by tumour necrosis factor requires the Akt serine-threonine kinase, Nature 401:82-85 (1999). 220. M. Grilli, M. Pizzi, M. Memo, Spano, and P, Neuroprotection by aspirin and sodium salicylate through blockade of NF-kappaB activation, Science 274:1383-1385 (1996). 221. H. Ko, K. Park, H. Kim, et al., Ca2+-mediated activation of c-Jun N-terminal kinase and nuclear factor kappa B by NMDA in cortical cell cultures, J Neurochem 71:13901395 (1998). 222. M. Grilli, and M. Memo, Possible role of NF-kappaB and p53 in the glutamateinduced pro-apoptotic nuonal pathway, Cell Death Differen 6:22-27 (1999). 223. K. Bales, Y. Du, R. Dodel, G. Yan, E. Hamilton-Byrd, and S. Paul, The NFkappaB/Rel family of proteins mediates A beta-induced neurotoxicity and glial activation, Molec Brain Res 57:63-72 (1998). 224. N. Perkins, The Rel/NF-kappa B family: friend and foe, Trends Biochem Sci 25:434440 (2000). 225. M. Whitehouse, Uncoupling of oxidative phosphorylation in a connective tissue (cartilage) and liver mitochondria by salicylate analogues: Relationship of structure to activity, Biochem Pharmacol 13:319-336 (1964). 226. M. Mehlman, R. Tobin, and E. Sporn, Oxidative phosphorylation and respiration by rat liver mitochondria from aspirin-treated rats, Biochem Pharmacol 21:32793285 (1972). 227. R. Haas, W. Parker, Jr., D. Stumpf, and L. Eguren, Salicylate-induced loose coupling: protonmotive force measurements, Biochem Pharmacol 34:900-902 (1985). 228. S. Somasundaram, H. Hayllar, S. Rafi, J. Wrigglesworth, A. Macpherson, and I. Bjarnason, The biochemical basis of non-steroidal anti-inflammatory drug-induced damage to the gastrointestinal tract: a review and a hypothesis, Scand J Gastroenterol 30:289-299 (1995). 229. T. Mahmud, S. Rafi, D. Scott, J. Wrigglesworth, and I. Bjarnason, Nonsteroidal antiinflammatory drugs and uncoupling of mitochondrial oxidative phosphorylation, Arthritis Rheum 39:1998-2003 (1996). 230. C. Sen, and L. Packer, Antioxidant and redox regulation of gene transcription, FASEB J 10:709-720 (1996). 231. V. Lakshminarayanan, E. Drab-Weiss, and K. Roebuck, H2O2 and tumor necrosis factor-alpha induce differential binding of the redox-responsive transcription factors

254

Chapter 12

AP-1 and NF-kappaB to the interleukin-8 promoter in endothelial and epithelial cells, J Biol Chem 273:32670-32678 (1998). 232. E. Shaulian, and M. Karin. AP-l as a regulator of cell life and death, Nat Cell Biol 4:E131-6 (2002).

Chapter 13 Astrocyte Metabolism and Astrocyte-Neuron Interaction

13.1.

INTRODUCTION

A critically important dimension in the characterization of cerebral metabolism is the now well documented interaction between neurons and astrocytes. This metabolic interplay is the subject of this chapter, with emphasis on carbohydrates, amino acids, and lipids. The interaction may involve diverse processes, e.g., astrocytic support of neuronal metabolism and energetics, and astrocytic export of agents that may compromise normal neuronal function. Accordingly, astrocyte-neuron metabolic interaction is fundamental to an understanding of both normal synaptic function as well as neuronal injury and death. The initial portion of the present chapter emphasizes those aspects that can be viewed as normal. Certain phenomena that adversely affect cerebral function are addressed in the latter portion of this chapter, and in subsequent chapters.

13.2.

CARBOHYDRATE METABOLISM

Metabolically, the astrocyte is uniquely versatile among brain cells1. It has the capacity to take up glucose from the interstitial fluid and to direct it along any of several pathways including glycolysis, the pentose phosphate pathway (hexose monophosphate shunt), and glycogen synthesis, the latter stimulated by both insulin and glutamate2. Moreover, the astrocyte expresses enzymes required for gluconeogenesis, utilizing precursors derived from interstitial fluid such as lactate and alanine (Fig. 13.1C). Significantly, despite the astrocyte’s ability to generate glucose255

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6-phosphate via either glycogenolysis or gluconeogenesis, and its limited expression of glucose-6-phosphatase, it does not export glucose3. However, the astrocyte may export lactate4,5, derived from glucose obtained from extracellular sources, glycogenolysis, or gluconeogenesis. As carrier-mediated neuronal lactate uptake6,7 and its cellular metabolism8 are established, it has been suggested that astrocyte release of lactate is important in the support of neuronal metabolic and energy requirements1,4,9-11. With regard to the energetics and efficiency of ATP generation, lactate oxidation is similar to glucose in that its complete mitochondrial oxidation provides a theoretical yield of 5.0 ATP molecules per molecule consumed. Also, its efficiency in fueling the citric acid cycle and the electron transport chain is advantageous because its initial conversion to pyruvate provides for regulation at the level of PDH (Fig. 3.2, facing p. 21, and Fig. 13.2). Lactate has been found to support synaptic function in rat hippocampal slices12,13; while this may reflect in part the formation of glutamate by a reaction sequence initiated by pyruvate carboxylase-mediated conversion of pyruvate to oxaloacetate14, the quantitative significance of this reaction in neurons has been questioned15,16. Moreover, recent evidence indicates that lactate alone is insufficient to maintain normal neuronal oxidative metabolism17, or normal synaptic potential and and that lactate derived from plasma exchanges with a brain lactate pool but does not replace glucose as an energy source19. In addition, uptake of lactate by the hepatocyte is inhibited by ketone bodies20,21, a mechanism which presumably would operate in neurons. Thus, the importance of astrocyte lactate export in the fueling of neuronal metabolism during activation remains uncertain; Figure 13.1. Relevant aspects of intermediary metabolism. Pathway arrow thickness approximates relative activity; dotted lines: relatively or absolutely suppressed; enclosed areas represent mitochondria. A. Glycolysis- and citric acid cycle-fueled ATP synthesis. Complete oxidation of glucose via aerobic glycolysis, pyruvate dehydrogenase, and the citric acid cycle, yields approximately 30 molecules of ATP per molecule of glucose. Gluconeogenesis is suppressed. B. Fatty acid oxidation. Mitochondrial oxidation of long chain fatty acids requires CPT-I-mediated formation of acylcarnitine. Fatty acids present in excess may override malonyl CoA inhibition of CPT-I, driving inappropriately augmented mitochondrial oxidation. The theoretical maximum yield of ATP from complete oxidation of palmitate (~100) is variably diminished to as little as ~20 in a ketogenic cell (astrocyte or hepatocyte) by diversion of acetyl CoA to ketogenesis rather than to the citric acid cycle. C.Gluconeogenesis. The process is normally fueled energetically by the mitochondrial oxidation of fatty acids. Glycolysis is suppressed. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; F26BP: fructose-2,6-bisphosphate; NADPH: nucleotide adenine triphosphate, reduced form; PCB: pyruvate carboxylase; PEPCK: phosphoenolpyruvate carboxykinase; PL: glycerophosphatides; TG: triacylglycerols.

13. Astrocyte Metabolism and Astrocyte-Neuron Interaction

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Figure 13.2. Fatty acid and ketone body oxidation in mitochondria suppresses glycolysis. Mitochondrial of fatty acids (astrocytes) and ketone bodies (neurons) generates three moieties that normally activate kinase-mediated feedback inhibition of PDH, i.e., acetyl CoA, ATP, and NADH. This effect, together with an accompanying suppression of glycolysis, decreases mitochondrial oxidation of pyruvate, and overall utilization of glucose. In peripheral tissues, especially muscle, this effect of mitochondrial fatty acid oxidation contributes to the “Randle cycle” inhibition of glucose utilization, and insulin resistance. Dashed lines: inhibition of PDH by acetyl CoA/CoA, ATP/ADP, and NADH/NAD. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; NADPH: nucleotide adenine triphosphate, reduced form; PDH: pyruvate dehydrogenase.

this question is discussed further below (see this Chapter 13.3.: Amino Acid Metabolism and the Glutamine Shuttle). Astrocyte expression of pyruvate carboxylase22 is of major importance in brain metabolism, and is an essential component of the gluconeogenic pathway (Fig. 13.1C). Pyruvate carboxylase-catalyzed conversion of pyruvate to oxaloacetate also contributes to the formation of citric acid cycle intermediates, thereby supporting oxidative energetics and other important metabolic functions. Among these, formation of NADPH, a key requirement for astrocyte biosynthesis of fatty acids, sterols, and glutathione23,24, is catalyzed by malic enzyme-mediated oxidative decarboxylation of malate to pyruvate25. Although neuronal expression of pyruvate carboxylase is of uncertain significance15,16, generation of citric acid cycle intermediates via this pathway in astrocytes appears to be of considerable importance in the exchange of neurotransmitter amino acids between astrocytes and neurons.


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13.3. AMINO ACID METABOLISM AND THE GLUTAMINE SHUTTLE Regulated neuronal synaptic export of the amino acids glutamate and (GABA) plays an essential role in neurotransmission. These mediators can be synthesized de novo from 2-oxo-glutarate an intermediate in the citric acid cycle. However, substantial utilization of this pathway without replenishment of the citric acid cycle intermediates thus consumed could compromise the supply of electrons to the electron transport chain and ATP generation. As pyruvate carboxylase-mediated neuronal compensation for this circumstance appears at best uncertain as noted15,16, the requirement during neuronal activation for near simultaneous increases in both ATP generation and neurotransmitter amino acid export would impose conflicting demands on the citric acid cycle. This dilemma appears to be addressed at least in part by the glutamine shuttle, a neuron-astrocyte interaction that depends on astrocyte glutamine synthase expression5,26-30. In this process, neuronal depolarization induces a rise in extracellular which leads to activation of pyruvate carboxylase-mediated formation of oxaloacetate in the astrocyte31. The resulting increase in citric acid cycle intermediates supports augmented astrocyte glutamine synthesis via transamination of 2-oxo-glutarate to glutamate, followed by glutamine synthase-mediated amidation of glutamate. The glutamine so produced is exported from the astrocyte and is readily taken up by neighboring neurons, which convert it to glutamate (via glutaminase) or GABA (via glutamate decarboxylase). These neurotransmitters are returned to the interstitial compartment during synaptic release; glutamate that is thereby reacquired by astrocytes is reconverted in part to glutamine. Thus, the glutamine shuttle plays an important role in brain metabolism. Although it appears to represent a net loss of 2-oxoglutarate from the astrocyte citric acid cycle, the loss may be compensated for by pyruvate carboxylase-mediated conversion of pyruvate to oxaloacetate. Unfortunately, this compensatory pathway consumes pyruvate; and, as pyruvate is also the major precursor of lactate, availability of lactate for export by the astrocyte in support of neuronal utilization during activation could be limited as a result. To some extent, this limitation may be ameliorated by neuronal export of alanine which is readily utilized by astrocytes, providing both for glutamine resynthesis and pyruvate32. In any case, neuronal uptake of glutamine exported from astrocytes would diminish the need for neuronal diversion of citric acid cycle intermediates toward neurotransmitter amino acid

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synthesis. In turn, this would permit maximal commitment of citric acid cycle intermediates to ATP generation, thereby augmenting the energy available for neuronal activation. Apart from its recognized relationship to neurotransmitter generation, then, the potentially favorable influence of the glutamine shuttle on neuronal energetics should not be underestimated28 (Fig. 3.2, facing p. 21). Thus, glutamine alone is able to support ATP generation and rapid growth in several normal and neoplastic cell types33,34, including hepatoma35 and fibrosarcoma cells36, at rates equal to those attainable with glucose as sole substrate; moreover, glutamine is the principal nutrient of enterocytes in vivo33. In the special case of neuronal activation, however, it appears likely that the relative impact of glutamine on each of the two obligatory yet potentially conflicting neuronal priorities (i.e., the requirement for near simultaneous neurotransmitter release and increased energy generation) will depend on the functional state of both the astrocyte and the neuron, and on the availability of alternative substrates. Recent studies have further addressed the metabolic and energetic interactions between the neuron and the astrocyte, as concerns their respective roles in cerebral glucose utilization in relation to glutamatergic 37 , it was shown neuron activation. In studies employing that rat cerebral glucose oxidation was coupled stoichiometrically at close to 1:1 with glutamate-neurotransmitter cycling, as reflected by the rate of glutamine synthesis. The findings were interpreted as showing that, above a certain basal level required by ongoing metabolic activity in the nonactivated neuron, incremental cortical energy production was linearly related to the support of glutamatergic synaptic activity. The results of these important observations were regarded as consistent with earlier evidence10 which suggested a striking and novel compartmentalization of glucose metabolism between astrocyte and neuron38. In these earlier co-culture studies10, it was demonstrated that astrocyte uptake of glutamate stimulated glycolysis and release of lactate, which was taken up by neurons and oxidized via the citric acid cycle. Thus, anaerobic glycolysis would provide the astrocyte with 2 ATP from each molecule of glucose. While export of the lactate so produced could provide the neuron with an additional approximately 30 ATP (from oxidation of two molecules of lactate), this potential energy supply would be diminished to the extent that pyruvate is consumed in neuronal generation of alanine for export to the astrocyte, as described above32. In view of the energy costs associated with glutamate uptake and glutamine synthesis (1 ATP for each step), these observations were interpreted37 as consistent with earlier evidence of lactate production by astrocytes and uptake by neurons4,6,9, as well as with the more recent evidence discussed


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above suggesting a 1:1 stoichiometry between glucose utilization and glutamate cycling. Support for this hypothesis was also provided by the similarities in glial localization of and glutamate transporter39, and by analogous compartmentalization of glucose utilization in other systems, including the glial/photoreceptor cells of the retina and the Sertoli cells/spermatids of the testis; in both cases, the latter cell population utilizes lactate generated via glycolysis in the former10. However, in the above experiments glutamate-stimulated lactate production appeared to be higher than would be expected, exceeding the 10 . apparent uptake of glucose as estimated with Furthermore, in cultured astrocytes only a small fraction of exogenous glutamate is converted to glutamine, the remainder being utilized in the citric acid cycle and other pathways40. Although this would permit diversion of glucose to the generation and export of lactate, glutamate itself would be progressively consumed. Thus, the model38 would appear to require that the products of anaerobic glycolysis (ATP and lactate) be dedicated entirely to the generation of intermediates that would be unavailable for astrocyte needs: ATP would be consumed in glutamate uptake and glutamine synthesis/export41, while lactate would be exported28. In the absence of some other energy source, this would seem to be incompatible with astrocyte viability, given other ongoing energy needs of this metabolically active cell. Moreover, the glutamine so supplied to the neuron may be utilized in pathways other than as a source of glutamate42. Similarly, as some fraction of the lactate taken up by neurons may be utilized in the generation of glutamate14,43 and alanine32, not all of it would be available for support of oxidative phsophorylation. Furthermore, as discussed above (see this Chapter 13.2.: Carbohydrate Metabolism), lactate may not support synaptic function as effectively as glucose17,18,44, while plasma-derived lactate does not appear to contribute to energy generation19. Possibly, metabolic demands on the astrocyte would be diminished to the extent that interstitial glucose is utilized directly by neurons45, given that these cells express plasma membrane glucose transporters the kinetics of which are relatively favorable46. The proposed model has been questioned on kinetic and other grounds as well47. Thus, while it retains attractive features11,38,48-50, and given current uncertainties in interpretation of isotope exchange data51, it is important to consider the possibility that in vivo availability of fuels other than glucose and glutamine, e.g., fatty acids, could contribute to the support of metabolic and energy requirements of the astrocyte. In addition, fatty acid oxidation in astrocyte mitochondria would suppress astrocyte PDH, thereby limiting oxidation of glucose and possibly glutamate. This would permit maximal diversion to the neuron of two substrates that are

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uniquely important in neuronal activation and synaptic transmission, i.e., plasma-derived glucose and astrocyte-generated glutamine.

13.4. FATTY ACIDS, ANTIOXIDANT DEFENSE, HIV DEMENTIA, AND CELL PROLIFERATION Substantial evidence indicates that, in contrast to the neuron, both neonatal and adult astrocytes are broadly equipped to oxidize long chain fatty acids52-55, and to defend themselves against the formation of reactive oxygen species that may emanate from that and other sources. In addition to the requisite components for mitochondrial fatty acid including the liver isoform of CPT-I55,56, the astrocyte expresses three or four distinct FABP isoforms, i.e., brain (B-FABP), heart (HFABP), and keratinocyte (K-FABP)57,58, and in adult rat, the liver isoform (L-FABP)59. Expression of B-FABP60,61 is most abundant in glial cells57,58,62, is regulated developmentally in response to a signal originating in neurons63, and is expressed in GFAP-expressing malignant glioma cells64. While its normal expression is region-specific62,65, it is the most abundant isoform in mature brain, exceeding H-FABP in abundance in mouse brain by an order of magnitude66. Its binding affinities for most long chain fatty acids are similar to those of other mammalian FABP. However, human B-FABP preferentially binds oleic acid as well as omega-3 fatty acids67, which may be important in their neuroprotective effects68,69. In contrast, the affinity of the murine protein for arachidonate is the highest reported for any FABP-ligand 70 interaction . B-FABP likely represents the active principle in a partially purified brain FABP fraction shown earlier to stimulate synaptosomal uptake of aspartate, glutamate, and GABA, through its binding and sequestration of long chain fatty acids59,71; this observation also demonstrates the vulnerability of this critical transport mechanism to inhibition by fatty acids. A cDNA for human B-FABP has been isolated and expressed72. It exhibits 81-86% nucleotide sequence identities and 87-91% amino acid sequence identities with rat brain, mouse brain, and chick retina FABPs. Recent studies demonstrate that binds long chain fatty acids and exhibits certain sequence homologies with FABP, suggesting a relationship to the FABP family73, and possibly contributing to redox-dependent adduct formation in Parkinson disease74.


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Figure 13.3. Plasma FFA-driven astrocyte ketogenesis: hypothesized neuronal oxidative stress and impaired glycolysis. Astrocyte fatty acid uptake, mitochondrial and generation of ketone bodies and ROS, are driven by the plasma concentration of FFA ([FFA]) and the [FFA]/[albumin] molar ratio; high [FFA] may override the CPT-I carnitine palmitoyltransferase-I control point. Ketone bodies and diffusible ROS (e.g., are exported from the astrocyte. Ketone bodies derived from astrocyte and/or plasma are utilized in preference to glucose by nearby neurons, in which their largely unregulated entry into mitochondria induces feedback inhibition of both PDH and glucose utilization (see Fig. 13.2), while their uncoupling-like effect may compromise mitochondrial energetics. Ketone bodies may also generate neuronal ROS, as the result of their relatively uncontrolled fueling of the citric acid cycle and electron transport chain. Thus, in addition to the inefficiencies of ketone body utilization as fuel, the neuron may be subjected to oxidative stress, reflecting ROS generated in both astrocyte and neuron. Dashed lines: inhibition of PDH by increased ratios of acetyl CoA/CoA, ATP/ADP, and NADH/NAD. Abbreviations — CPT-I: carnitine palmitoyltransferase-I; FFA: free fatty acid; PDH: pyruvate dehydrogenase; ROS: reactive oxygen species.

In view of the fact that long chain fatty acids are not utilized as an energy source by most neurons and do not accumulate in the interstitial compartment, fatty acid uptake by astrocytes and other glial elements appears to be directly related to brain uptake of plasma FFA. Similar to the hepatocyte, astrocyte fatty acids readily undergo mitochondrial and thus may contribute to ATP generation via the citric acid cycle. As noted, however, the astrocyte also is capable of diverting fatty acid-derived acetyl CoA units to the synthesis of ketone bodies (Fig. 3.2, facing p. 21, and Fig. 13.3). This process is more efficient than that in liver, in that as the process of nears completion in the astrocyte, the terminal four carbon remnant is converted directly to acetoacetate, bypassing the final thiolase step and the methyl-glutaryl CoA cycle segment of the ketogenesis pathway54.

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Ketogenesis in astrocytes is enhanced by tetrahydrocannabinol75,76, and may also be supported by leucine metabolism77. Moreover, receptormediated utilization of anandamide is associated with lipid peroxidation, indicating oxidative stress78. In hepatocytes, fatty acids79, as well as hypoxia and increased [AMP]/[ATP], activate AMP-activated protein kinase (AMPK); in both hepatocytes and astrocytes, AMPK augments fatty acid oxidation and ketogenesis80. After export into the interstitial fluid, ketone bodies are readily taken up by neurons as the preferred oxidizable substrate52-54,77 as described above (see Chapter 12.3.: Ketone Bodies).

13.4.1. Astrocyte-Neuron Interaction and Oxidative Stress This aspect of astrocyte fatty acid metabolism may prove to be critically important in astrocyte-neuron interactions in health and disease. For example, it appears likely that mitochondrial of fatty acids would constitute an important energy source for the astrocyte during or immediately following neuronal activation, as this is the period during which astrocyte citric acid cycle intermediates may be consumed in the biosynthesis and export of glutamine and lactate. In this context, therefore, availability of plasma-derived long chain fatty acids to the astrocyte would help to support neuronal function, in part by suppressing astrocyte consumption of glucose (and that of both pyruvate and lactate) via inhibition of PDH. Conversely, under conditions predisposing to sustained elevation of plasma FFA, such as HIV infection81, and obesity, insulin resistance, or diabetes mellitus82, increased brain fatty acid uptake may drive excessive astrocyte fatty acid oxidation and ketogenesis (as may also occur in liver), with adverse consequences for both astrocyte and neuron. Thus, in the astrocyte, excessive fatty acid oxidation would contribute to ROS generation and oxidative stress as a result of “leakage” of electrons from the electron transport chain in mitochondria (Fig. 13.4) and/or fatty acid in peroxisomes 83 , a potentially (Fig. 13.5). As human astrocytes do not express ameliorating adaptive response to these ROS-generating phenomena is unavailable (see Chapter 12.4.2.: Oxidation of Fatty Acids). Such adverse effects, and a protective effect of fatty acid binding by albumin, are demonstrable in vitro84. For the neuron, sustained increases in plasma FFA may compromise this cell’s function indirectly in several ways. Thus, augmented astrocyte ketogenesis would drive increased neuronal utilization of ketone bodies in preference to either glucose or lactate (Figs. 13.2 and 13.3), impairing efficiency of


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Figure 13.4. Mitochondrial voltage-dependent anion channel (VDAC) components: relationship to electron transport chain and generation of reactive oxygen species. While detailed structural relationships remain undefined (see Chapter 3.3.1. Aerobic Glycolysis, Mitochondrial Respiration, and Oxidative Phosphorylation), juxtaposition of hexokinase to the adenine nucleotide translocase in proliferating cells optimizes kinetics of glucose phosphorylation to glucose-6-phosphate and entry into the glycolytic pathway, leading to increased pyruvate formation. Increased mitochondrial fatty acid oxidation inhibits both pyruvate dehydrogenase and hexokinase (Randle cycle). Excess cellular fatty acid abundance may disrupt both VDAC and the contact sites, and overload the electron transport chain. This predisposes to “leakage” from complexes I and III, and ubiquinone, with formation of ROS. Pro-apoptotic members of the Bcl-2 family (e.g., Bax) may be largely cytosolic and associate with the mitochondrial membrane only after activation of the apoptotic cascade. Abbreviations — ADP/ATP: adenosine di/triphosphate; CYT C: cytochrome c; inner mitochondrial membrane potential; UBIQ: ubiquinone; ROS: reactive oxygen species; VDAC: voltage-dependent anion channel. With permission (see Fig. 3.4.)

activation and promoting oxidative stress. Moreover, oxidative stressinduced impairment of neuronal function and synaptic transmission may also result from generation in astrocytes of longer-lived diffusible oxidants such as which decrease synaptosomal ATP concentration and inhibit activation-induced, release of glutamate85-89. Finally, generation of ROS and oxidative stress within the astrocyte inhibits glutamate uptake90, potentially contributing to excitotoxic neuronal injury (see Chapter 14.5.2.2.: Excitotoxicity — Origins of Oxidative Stress). Unfortunately, as has been emphasized24,91, neurons (except for those expressing UCP-2 and involved in neuroendocrine and metabolic

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Figure 13.5. Pathways of cellular fatty acid metabolism. Both mitochondrial and extramitochondrial FA oxidation pathways are transcriptionally regulated by members of the PPAR family. PPAR may be activated by increased cellular abundance of FA, as well as other factors acting directly or indirectly. Absence of in primary human brain astrocytes and astrocyte-derived lines (see text) suggests that these cells may be compromised in regard to a potentially protective adaptive response to increased fatty acid exposure (see Chapter 12: Utilization of Oxidizable Substrates in Brain). Abbreviations — CPT-I: carnitine palmitoyltransferase-I; FA: fatty acid; FA-CoA: fatty acid-coenzyme A thioester; PL: glycerophosphatides; PPAR: peroxisome proliferatoractivated receptor; TG: triacylglycerols.

regulation) are less well equipped than astrocytes to mitigate oxidative stress, and therefore are more susceptible to its consequences92. Thus, glial cells in general93 and astrocytes in particular reportedly contain higher concentrations of glutathione than do neurons92,94,95, although exceptions have been noted96. Mitochondrial manganese superoxide dismutase is strongly expressed in astrocytes97, but variably in neurons98, although neurons contain higher concentrations of ascorbate93. Astrocyte defense against is mediated by both catalase and glutathione 95,99 , which indirectly may also protect neurons100,101; astrocytes peroxidase also protect neurons against nitric oxide toxicity 102 . In addition, astrocytes export glutathione, thereby providing thiols to neurons95,103, and rapidly replenish their own normally high glutathione concentrations95,104. Catalase associated with peroxisome-like structures105-107, and glutathione reductase108, are present throughout brain.


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Two critically important implications of this metabolic specialization of neurons and astrocytes deserve particular emphasis. First, neurons appear to be far more susceptible than glial cells to the adverse effects of exposure to fatty acids and their oxidation products. Second, except as noted above (see Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior), oxidative metabolism of fatty acids in brain under normal conditions occurs chiefly in astrocytes or other glia. This compartmentalization affords some degree of protection to the neurons with respect to the oxidative stress and inefficiency that may result from exposure to, and oxidation of, excessive quantities of long chain fatty acids. It also establishes a mechanism by which fatty acid oxidation can be employed advantageously both as an energy source for the astrocyte and/or the in situ generation of important alternative neuronal fuels, i.e., ketone bodies. To the extent that ketone bodies are needed to support neuronal metabolism, they are thus immediately available via ketogenesis in adjacent astrocytes. However, ketone body oxidation in neuronal mitochondria represents a potentially abundant endogenous cause of mitochondrial energetic inefficiency and source of ROS and oxidative stress. Thus, excessive brain exposure to FFA (with resulting neuronal exposure to ketone bodies) could over time contribute significantly to oxidant-induced neuronal injury and the pathogenesis of neurodegenerative disorders. Two important and poorly understood causes of progressive dementing illness may exemplify the consequences of this mechanism. The first, Alzheimer disease, is dealt with in detail below (see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). The second is HIV-associated dementia, alluded to briefly above (see Chapter 12.5.: Cytokines and the Brain) and in the following paragraph. In this condition, progressive dementia occurs despite the absence of neuronal HIV infection and the lack of any consistent relationship to viral burden or HIV encephalitis extent and location109-111. The latter process appears to involve primarily both astrocytes and systemic macrophages recruited to take up residence in brain.

13.4.2. HIV Dementia HIV-associated dementia appears to bear certain similarities in its pathogenesis to the evolution of the endothelial lesion of atherosclerosis (see Chapter 7.4.: Consequences of Intramitochondrial Oxidative Stress: Atherosclerosis and Beyond). In the latter, cellular lipid overload resulting from sustained increases in uptake of oxidized low density lipoproteins (LDL) and/or plasma FFA may drive excessive mitochondrial

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fatty acid oxidation in both endothelial cell and arterial wall macrophage, in turn generating oxidative stress and thus contributing importantly to monocyte/macrophage recruitment and the generation of proinflammatory cytokines112-119. As noted above, the proinflammatory cytokine response that is a hallmark of advanced HIV infection increases adipocyte lipolysis, plasma [FFA], and as a result tissue fatty acid uptake120. The resulting increases in tissue fatty acid abundance would be expected to drive increased mitochondrial fatty acid oxidation in both astrocytes and macrophages. Although macrophages express human brain astrocytes do not83 (see above, and Chapter 12.4.2.: Oxidation of Fatty Acids), and therefore may not adapt sufficiently to ameliorate fatty acid oxidation-induced intramitochondrial oxidative stress121. Excessive astrocytic generation of both diffusible ROS (e.g., and ketone bodies would also adversely affect nearby neurons, in which antioxidant defenses are less well developed. The resulting oxidative stress-induced neuronal injury would be sustained and progressive, while ketone body utilization as fuel would compromise neuronal mitochondrial energetics. Available evidence suggests that this hypothetical scenario could account for other indications of an altered astrocyte-neuron interaction in HIV122, and/or neurochemical changes observed in a murine model of virally-induced immunodeficiency and While such interactions would be expected to encephalopathy123. contribute significantly to the development and evolution of HIVassociated dementia, direct experimental testing of this hypothesis is required (see Fig. 13.3, and Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors).

13.4.3. Cell Proliferation and Tumorigenesis Neuron-astrocyte metabolic interactions may have other significant consequences. Thus, it is possible that increased astrocyte fatty acid oxidation and ketogenesis, and the resulting excessive neuronal oxidation of ketone bodies, would suppress glucose utilization by both cell types. As a result, glucose availability to a proliferative neural progenitor cell compartment may be increased124-127. These cells, which may be either glial or neuronal128-130, are potentially susceptible to the proliferative stimulus that abundance of glutamate131, glucose, and fatty acids may represent (see Chapter 4.4.: Fatty Acids, Modulation of Cell Growth, and the AMP-Activated Protein Kinase, and Chapter 8: Metabolism and Gene Expression in Liver Regeneration). Glioma cells, in contrast, appear to release glutamate, enhancing their own growth by excitotoxic elimination Furthermore, proinflammatory of potentially competing neurons132.


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cytokines are mitogenic for astrocytes133,134, as they are for hepatocytes after partial hepatectomy. In both circumstances, cell proliferation is supported by wg/Wnt-like signaling135(also, see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable), suppressed mitochondrial fatty acid oxidation, augmented glycolytic fueling of the citric acid cycle, and availability of fatty acids for membrane and eicosanoid biogenesis (see Chapter 4.4.: Fatty Acids, Modulation of Cell Growth, and the AMP-Activated Protein Kinase). These wg/Wnt-like signaling features are consistent with inhibition of neural progenitor cell growth by the tumor suppressor PTEN gene product136, a lipid phosphatase which inhibits PI3K/Akt/PKB signaling137. Clearly, the metabolic characteristics of the astrocyte and astrocyte-neuronal interaction are of overriding significance; among their other effects, they have the potential to influence cell proliferation and tumorigenesis in brain.

13.5. REFERENCES 1. H. Wiesinger, B. Hamprecht, and R. Dringen, Metabolic pathways for glucose in astrocytes, Glia 21:22-34 (1997). 2. M. Hamai, Y. Minokoshi, and T. Shimazu, L-Glutamate and insulin enhance glycogen synthesis in cultured astrocytes from the rat brain through different intracellular mechanisms, J Neurochem 73:400-407 (1999). 3. J. Gotoh, Y. Itoh, T. Kuang, M. Cook, M. Law, and L. Sokoloff, Negligible glucose-6phosphatase activity in cultured astroglia, J Neurochem 74:1400-1408 (2000). 4. R. Dringen, R. Gebhardt, and B. Hamprecht, Glycogen in astrocytes: possible function as lactate supply for neighboring cells, Brain Res 623:208-214 (1993A). 5. A. Schousboe, N. Westergaard, H. Waagepetersen, O. Larsson, I. Bakken, and U. Sonnewald, Trafficking between glia and neurons of TCA cycle intermediates and related metabolites, Glia 21:99-105 (1997). 6. R. Dringen, H. Wiesinger, and B. Hamprecht, Uptake of L-lactate by cultured rat brain neurons, Neurosci Letters 163:5-7 (1993D). 7. L. Pellerin, G. Pellegri, J.-L. Martin, and P. Magistretti, Expression of monocarboxylate transporter mRNAs in mouse brain: support for a distinct role of lactate as an energy substrate for the neonatal vs adult brain, Proc Natl Aca Sci USA 95:3990-3995 (1998). 8. M. McKenna, J. Tildon, J. Stevenson, and I. Hopkins, Energy metabolism in cortical synaptic terminals from weanling and mature rat brain: evidence for multiple compartments of tricarboxylic acid cycle activity, Dev Neurosci 16:291-300 (1994). 9. R. Dringen, and B. Hamprecht, Differences in glycogen metabolism in astroglia-rich primary cultures and sorbitol-selected astroglial cultures derived from mouse brain, Glia 8:143-149 (1993B). 10. L. Pellerin, and P. Magistretti, Glutamate uptake into astrocytes stimulates aerobic glycolysis: a mechanism coupling neuronal activity to glucose utilization, Proc Natl Acad Sci USA 91:10625-10629 (1994).

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11. L. Pellerin, Lactate as a pivotal element in neuron-glia metabolic cooperation, Neurochem Int 43:331-8 (2003). 12. A. Schurr, C. West, and B. Rigor, Lactate-supported synaptic function in the rat hippocampal slice preparation, Science 240:1326-1328 (1988). 13. Y. Izumi, A. Benz, C. Zorumski, and J. Olney, Effects of lactate and pyruvate on glucose deprivation in rat hippocampal slices, Neuroreport 5:617-620 (1994). 14. B. Hassel, and A. Bråthe, Neuronal pyruvate carboxylation supports formation of transmitter glutamate, J Neurosci 20:1342-1347 (2000B). 15. B. Hassel, Pyruvate carboxylation in neurons, J Neurosci Res 66:755-762 (2001). 16. H. Waagepetersen, H. Qu, A. Schousboe, and U. Sonnewald, Elucidation of the quantitative significance of pyruvate carboxylation in cultured cerebellar neurons and astrocytes, J Neurosci Res 66:763-770 (2001). 17. A. Gjedde, and S. Marrett, Glycolysis in neurons, not astrocytes, delays oxidative metabolism of human visual cortex during sustained checkerboard stimulation in vivo, J Cereb Blood Flow Metab 21:1384-1392 (2001). 18. T. Takata, T. Sakurai, B. Yang, K. Yokono, and Y. Okada, Effect of lactate on the synaptic potential, energy metabolism, calcium homeostasis and extracellular glutamate concentration in the dentate gyrus of the hippocampus from guinea-pig, Neuroscience 104:371-378 (2001). 19. G. Leegsma-Vogt, K. Venema, and J. Korf, Evidence for a lactate pool in the rat brain that is not used as an energy supply under normoglycemic conditions, J Cereb Blood Flow Metab 23:933-41 (2003). 20. P. Fafournoux, C. Demigé, and C. Rémésy, Carrier-mediated uptake of lactate in rat hepatocytes: Effects of pH and possible mechanisms for L-lactate transport, J Biol Chem 260:292-299 (1985). 21. R. Poole, and A. Halestrap, Transport of lactate and other monocarboxylates across mammalian plasma membranes, Am J Physiol 264:C761-C782 (1993). 22. M. Cesar, and B. Hamprecht, Immunocytochemical examination of neural rat and mouse primary cultures using monoclonal antibodies raised against pyruvate carboxylase, J Neurochem 64:2312-2318 (1995). 23. M. McKenna, J. Tildon, J. Stevenson, X. Huang, and K. Kingwell, Regulation of mitochondrial and cytosolic malic enzymes from cultured rat brain astrocytes, Neurochem Res 20:1491-1501 (1995). 24. J. Wilson, Antioxidant defense of the brain: a role for astrocytes, Can J Physiol Pharmacol 75:1149-1163 (1997). 25. P. Alves, R. Nunes, C. Zhang, et al., Metabolism of 3-(13)C-malate in primary cultures of mouse astrocytes, Devel Neurosci 22:456-462 (2000). 26. W. Gamberino, D. Berkich, C. Lynch, B. Xu, and K. LaNoue, Role of pyruvate carboxylase in facilitation of synthesis of glutamate and glutamine in cultured astrocytes, J Neurochem 69:2312-2325 (1997). 27. U. Sonnewald, N. Westergaard, and A. Schousboe, Glutamate transport and metabolism in astrocytes, Glia 21:56-63 (1997). 28. L. Hertz, R. Dringen, A. Schousboe, and S. Robinson, Astrocytes: glutamate producers for neurons, J Neurosci Res 57:417-428 (1999). 29. L. Hertz, A. Yu, G. Kala, and A. Schousboe, Neuronal-astrocytic and cytosolicmitochondrial metabolite trafficking during brain activation, hyperammonemia and energy deprivation, Neurochem Intl 37:83-102 (2000). 30. M. McKenna, J. Stevenson, X. Huang, and I. Hopkins, Differential distribution of the enzymes glutamate dehydrogenase and aspartate aminotransferase in cortical synaptic


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mitochondria contributes to metabolic compartmentation in cortical synaptic terminals, Neurochem Intl 37:229-241 (2000). 31. E. Kaufman, and B. Driscoll, Evidence for cooperativity between neurons and astroglia in the regulation of CO2 fixation in vitro, Dev Neursci 15:299-305 (1993). 32. H. Waagepetersen, U. Sonnewald, O. Larsson, and A. Schousboe, A possible role of alanine for ammonia transfer between astrocytes and glutamatergic neurons, J Neurochem 75:471-479 (2000). 33. Z. Kovacevic, and J. McGivan, Mitochondrial metabolism of glutamine and glutamate and its physiological significance, Physiolog Rev 63:547-605 (1983). 34 M. Board, S. Humm, and E. Newsholme, Maximum activities of key enzymes of glycolysis, glutaminolysis, pentose phosphate pathway and tricarboxylic acid cycle in normal, neoplastic and suppressed cells, Biochem J 265:503-509 (1990). 35. R. Nakashima, M. Paggi, and P. Pedersen, Contributions of glycolysis and oxidative phosphorylation to adenosine production in AS-30D hepatoma cells, Cancer Res 44:5702-5706 (1984). 36. V. Goossens, J. Grooten, and W. Fiers, The oxidative metabolism of glutamine: A modulator of reactive oxygen intermediate-mediated cytotoxicity of tumor necrosis factor in L929 fibrosarcoma cells, J Biol Chem 271:192-196 (1996). 37. N. Sibson, A. Dhankhar, G. Mason, D. Rothman, K. Behar, and R. Shulman, Stoichiometric coupling of brain glucose metabolism and glutamatergic neuronal activity, Proc Natl Acad Sci USA 95:316-321 (1998). 38. P. Magistretti, L. Pellerin, D. Rothman, and R. Shulman, Energy on demand, Science 283:496-497 (1999). 39. N. Cholet, L. Pellerin, P. Magistretti, and E. Hamel, Similar Perisynaptic Glial Localization for the Na(+),K(+)-ATPase alpha(2) Subunit and the Glutamate Transporters GLAST and GLT-1 in the Rat Somatosensory Cortex, Cereb Cortex 12:515-525 (2002). 40. H. Qu, E. Faerø, P. Jørgensen, et al., Decreased glutamate metabolism in cultured astrocytes in the presence of thiopental, Biochem Pharmacol 58:1075-1080 (1999). 41. V. Lebon, K. Petersen, G. Cline, et al., Astroglial contribution to brain energy metabolism in humans revealed by 13C nuclear magnetic resonance spectroscopy: elucidation of the dominant pathway for neurotransmitter glutamate repletion and measurement of astrocytic oxidative metabolism, J Neurosci 22:1523-1531 (2002). 42. U. Sonnewald, and M. McKenna, Metabolic compartmentation in cortical synaptosomes: influence of glucose and preferential incorporation of endogenous glutamate into GABA, Neurochem Res 27:43-50 (2002). 43. B. Hassel, and A. Bråthe, Cerebral metabolism of lactate in vivo: Evidence for neuronal pyruvate carboxylation, J Cerebral Blood Flow Metab 20:327-336 (2000A). 44. G. Dienel, and L. Hertz, Glucose and lactate metabolism during brain activation, J Neurosci Res 66:824-838 (2001). 45. E. McNay, T. Fries, and P. Gold, Decreases in rat extracellular hippocampal glucose concentration associated with cognitive demand during a spatial task, Proc Natl Acad Sci USA 97:2881-2885 (2000). 46. S. Vannucci, F. Maher, and I. Simpson, Glucose transporter proteins in brain: delivery of glucose to neurons and glia, Glia 21:2-21 (1997). 47. C. Chih, and E. Roberts, Jr, Energy substrates for neurons during neural activity: a critical review of the astrocyte-neuron lactate shuttle hypothesis., J. Cerebral Blood Flow & Metabolism 23:1263-1281 (2003). 48. R. Shulman, F. Hyder, and D. Rothman, Lactate efflux and the neuroenergetic basis of brain function, NMR Biomed 14:389-396 (2001B).

272

Chapter 13

49. Y. Itoh, T. Esaki, K. Shimoji, et al., Dichloroacetate effects on glucose and lactate oxidation by neurons and astroglia in vitro and on glucose utilization by brain in vivo, Proc Natl Acad Sci U S A 100:4879-84 (2003). 50. A. Bouzier-Sore, P. Voisin, P. Canioni, P. Magistretti, and L. Pellerin, Lactate is a preferential oxidative energy substrate over glucose for neurons in culture., J. Cerebral Blood Flow & Metabolism 23:1298-1306 (2003). 51. G. A. Kimmich, J. A. Roussie, and J. Randies, Aspartate aminotransferase isotope exchange reactions: implications for glutamate/glutamine shuttle hypothesis, Am J Physiol Cell Physiol 282:C1404-13 (2002). 52. J. Edmond, R. Robbins, J. Bergstrom, R. Cole, and J. de Vellis, Capacity for substrate utilization in oxidative metabolism by neurons, astrocytes, and oligodendrocytes from developing brain in primary culture, J Neurosci Res 18:551561 (1987). 53. J. Edmond, Energy metabolism in developing brain cells, Can J Physiol Pharmacol 70:S118-29 (1992). 54. N. Auestad, R. Korsak, J. Morrow, and J. Edmond, Fatty acid oxidation and ketogenesis by astrocytes in primary culture, J Neurochem 56:1376-1386 (1991). 55. C. Blázquez, Sánchez, C, G. Velasco, and M. Guzmán, Role of carnitine palmitoyltransferase I in the control of ketogenesis in primary cultures of rat astrocytes, J Neurochem 71:1597-1606 (1998). 56. N. Brown, J. Hill, V. Esser, et al., Mouse white adipocytes and 3T3-L1 cells display an anomalous pattern of carnitine palmitoyltransferase (CPT) I isoform expression during differentiation: Inter-tissue and inter-species expression of CPT I and CPT II enzymes, Biochem J 327:225-231 (1997). 57. Y. Owada, T. Yoshimoto, and H. Kondo, Increased expression of the mRNA for brainand skin-type but not heart-type fatty acid binding proteins following kainic acid systemic administration in the hippocampal glia of adult rats, Brain Res Molec Brain Res 42:156-160 (1996A). 58. Y. Owada, T. Yoshimoto, and H. Kondo, Spatio-temporally differential expression of genes for three members of fatty acid binding proteins in developing and mature rat brains, J Chem Neuroanat 12:113-122 (1996B). 59. N. Bass, E. Raghupathy, D. Rhoads, J. Manning, and R. Ockner, Partial purification of molecular wieght 12,000 fatty acid binding proteins from rat brain and their effect on synaptosomal NA+-dependent amino acid uptake, Biochem 23:6539-6544 (1984). 60. E. Bennett, K. Stenvers, P. Lund, and B. Popko, Cloning and characterization of a cDNA encoding a novel fatty acid binding protein from rat brain, J Neurochem 63:1616-1624 (1994). 61. L. Feng, M. Hatten, and N. Heintz, Brain lipid-binding protein (BLBP): a novel signaling system in the developing mammalian CNS, Neuron 12:895-908 (1994). 62. A. Kurtz, A. Zimmer, F. Schnütgen, G. Brüning, F. Spener, and T. Müller, The expression pattern of a novel gene encoding brain-fatty acid binding protein correlates with neuronal and glial cell development, Development 120:2637-2649 (1994). 63. L. Feng, and N. Heintz, Differentiating neurons activate transcription of the brain lipid-binding protein gene in radial glia through a novel regulatory element, Development 121:1719-1730 (1995). 64. R. Godbout, D. Bisgrove, D. Shkolny, and R. Day, 3rd, Correlation of B-FABP and GFAP expression in malignant glioma, Oncogene 16:1955-1962 (1998).


273

65. F. Schnütgen, Börchers, T, T. Müller, and F. Spener, Heterologous expression and characterisation of mouse brain fatty acid binding protein, Biol Chem Hoppe-Seyler 377:211-215 (1996). 66. S. Myers-Payne, T. Hubbell, L. Pu, et al., Isolation and characterization of two fatty acid binding proteins from mouse brain, J Neurochem 66:1648-1656 (1996). 67. G. Balendiran, F. Schnütgen, G. Scapin, et al., Crystal structure and thermodynamic analysis of human brain fatty acid-binding protein, J Biol Chem 275:27045-27054 (2000). 68. Y. Ge, X. Wang, Z. Chen, N. Landman, E. H. Lo, and J. X. Kang, Gene transfer of the Caenorhabditis elegans n-3 fatty acid desaturase inhibits neuronal apoptosis, J Neurochem 82:1360-6 (2002). 69. D. S. Martin, P. E. Lonergan, B. Boland, et al., Apoptotic changes in the aged brain are triggered by interleukin-1 beta-induced activation of p38 and reversed by treatment with eicosapentaenoic acid, J Biol Chem 277:34239-46 (2002). 70. L. Xu, R. Sánchez, A. Sali, and N. Heintz, Ligand specificity of brain lipid-binding protein, J Biol Chem 271:24711-24719 (1996). 71. D. Rhoads, R. Ockner, N. Peterson, and E. Raghupathy, Modulation of membrane transport by free fatty acids: inhibition of synaptosomal sodium-dependent amino acid uptake, Biochem 22:1965-1970 (1983). 72. F. Shimizu, T. Watanabe, H. Shinomiya, Y. Nakamura, and T. Fujiwara, Isolation and expression of a cDNA for human brain fatty acid-binding protein (B-FABP), Biochim Biophys Acta 1354:24-28 (1997). 73. R. Sharon, M. Goldberg, I. Bar-Josef, R. Betensky, J. Shen, and D. Selkoe, alphaSynuclein occurs in lipid-rich high molecular weight complexes, binds fatty acids, and shows homology to the fatty acid-binding proteins, Proc Natl Acad Sci U S A 98:9110-9115 (2001). 74. K. Conway, J. Rochet, R. Bieganski, and P. Lansbury, Jr, Kinetic stabilization of the alpha-synuclein protofibril by a dopamine-alpha-synuclein adduct, Science 294:1346-1349 (2001). 75. C. Bláquez, C. Sánchez, A. Daza, I. Galve-Roperh, and M. Guzmán, The stimulation of ketogenesis by cannabinoids in cultured astrocytes defines carnitine palmitoyltransferase I as a new ceramide-activated enzyme, J Neurochem 72:17591768 (1999A). 76. M. Guzmán, and C. Sánchez, Effects of cannabinoids on energy metabolism, Life Sci 65:657-664 (1999). 77. M. Bixel, and B. Hamprecht, Generation of ketone bodies from leucine by cultured astroglial cells, J Neurochem 65:2450-61 (1995). 78. B. Costa, and M. Colleoni, Changes in rat brain energetic metabolism after exposure to anandamide or delta9-tetrahydrocannabinol, Eur J Pharmacol 395:1-7 (2000). 79. T. Kawaguchi, K. Osatomi, H. Yamashita, T. Kabashima, and K. Uyeda, Mechanism for fatty acid “sparing” effect on glucose-induced transcription: regulation of carbohydrate-responsive element-binding protein by AMP-activated protein kinase, J Biol Chem 277:3829-3835 (2002). 80. C. Blázquez, A. Woods, C. de, ML, D. Carling, and M. Guzmán, The AMP-activated protein kinase is involved in the regulation of ketone body production by astrocytes, Journal of Neurochemistry 73:1674-82 (1999B). 81. M. Navasa, K. Feingold, and C. Grunfeld, Effects of endotoxin and cytokines on hepatic lipid metabolism, Prog Liver Dis 15:147-170 (1997). 82. H. Ginsberg, Insulin resistance and cardiovascular disease, J Clin Invest 106:453458 (2000).

274

Chapter 13

83. N. Chattopadhyay, D. Singh, O. Heese, et al., Expression of peroxisome proliferator-

activated receptors (PPARs) in human astrocytic cells: PPARgamma agonists as inducers of apoptosis, J Neurosci Res 61:67-74 (2000). 84. A. Tabernero, A. Medina, L. Sánchez-Abarca, E. Lavado, and J. Medina, The effect of albumin on astrocyte energy metabolism is not brought about through the control of cytosolic Ca2+ concentrations but by free-fatty acid sequestration, Glia 25:1-9 (1999). 85. J. Sanchez-Prieto, T. Sihra, and D. Nicholls, Characterization of the exocytotic release of glutamate from guinea-pig cerebral cortical synaptosomes, J Neurochem 49:58-64 (1987). 86. S. Gilman, M. Bonner, and T. Pellmar, Peroxide effects on [3H]L-glutamate release by synaptosomes isolated from the cerebral cortex, Neurosci Lett 140:157-160 (1992). 87. F. Zoccarato, M. Valente, and A. Alexandre, Hydrogen peroxide induces a longlasting inhibition of the Ca(2+)-dependent glutamate release in cerebrocortical synaptosomes without interfering with cytosolic Ca2+, J Neurochem 64:2552-2558 (1995). 88. L. Tretter, C. Chinopoulos, and V. Adam-Vizi, Enhanced depolarization-evoked calcium signal and reduced [ATP]/[ADP] ratio are unrelated events induced by oxidative stress in synaptosomes, J Neurochem 69:2529-2537 (1997). 89. L. Tretter, and V. Adam-Vizi, Inhibition of Krebs cycle enzymes by hydrogen peroxide: A key role of [alpha]-ketoglutarate dehydrogenase in limiting NADH production under oxidative stress, J Neurosci 20:8972-8979 (2000). 90. Y. Chen, W. Ying, V. Simma, et al., Overexpression of Cu,Zn superoxide dismutase attenuates oxidative inhibition of astrocyte glutamate uptake, J Neurochem 75:939945 (2000). 91. S. Peuchen, J. Bolaños, S. Heales, A. Almeida, M. Duchen, and J. Clark, Interrelationships between astrocyte function, oxidative stress and antioxidant status within the central nervous system, Prog Neurobiol 52:261-281 (1997). 92. J. Bolaños, S. Heales, J. Land, and J. Clark, Effect of peroxynitrite on the mitochondrial respiratory chain: differential susceptibility of neurones and astrocytes in primary culture, J Neurochem 64:1965-1972 (1995). 93. M. Rice, and I. Russo-Menna, Differential compartmentalization of brain ascorbate and glutathione between neurons and glia, Neurosci 82:1213-1223 (1998). 94. S. Raps, J. Lai, L. Hertz, and A. Cooper, Glutathione is present in high concentrations in cultured astrocytes but not in cultured neurons, Brain Res 493:398401 (1989). 95. R. Dringen, Metabolism and functions of glutathione in brain, Prog Neurobiol 62:649-671 (2000). 96. J. Huang, and M. Philbert, Distribution of glutathione and glutathione-related enzyme systems in mitochondria and cytosol of cultured cerebellar astrocytes and granule cells, Brain Res 680:16-22 (1995). 97. E. Pinteaux, M. Perraut, and G. Tholey, Distribution of mitochondrial manganese superoxide dismutase among rat glial cells in culture, Glia 22:408-414 (1998). 98. L. Medina, G. Figueredo-Cardenas, and A. Reiner, Differential abundance of superoxide dismutase in interneurons versus projection neurons and in matrix versus striosome neurons in monkey striatum, Brain Research 708:59-70 (1996). 99. R. Dringen, and B. Hamprecht, Involvement of glutathione peroxidase and catalase in the disposal of exogenous hydrogen peroxide by cultured astroglial cells, Brain Res 759:67-75 (1997).


275

100. S. Desagher, J. Glowinski, and J. Premont, Astrocytes protect neurons from hydrogen peroxide toxicity, J Neurosci 16:2553-2562 (1996). 101. E. Blanc, A. Bruce-Keller, and M. Mattson, Astrocytic gap junctional communication decreases neuronal vulnerability to oxidative stress-induced disruption of Ca2+ homeostasis and cell death, J Neurochem 70:958-970 (1998). 102. Y. Chen, N. Vartiainen, W. Ying, P. Chan, J. Koistinaho, and R. Swanson, Astrocytes protect neurons from nitric oxide toxicity by a glutathione-dependent mechanism, J Neurochem 77:1601-1610 (2001). 103. X. Wang, and M. Cynader, Astrocytes provide cysteine to neurons by releasing glutathione, J Neurochem 74:1434-1442 (2000). 104. R. Dringen, and B. Hamprecht, Glutathione restoration as indicator for cellular metabolism of astroglial cells, Develop Neurosci 20:401-407 (1998). 105. G. Gaunt, and D. de, C, Subcellular distribution of d-amino acid oxidase and catalase in rat brain, J Neurochem 26:749-759 (1976). 106. O. McKenna, G. Arnold, and E. Holtzman, Microperoxisome distribution in the central nervous system of the rat, Brain Res 117:181-194 (1976). 107. T. Brannan, H. Maker, and I. Raes, Regional distribution of catalase in the adult rat brain, J Neurochem 36:307-309 (1981). 108. T. Brannan, H. Maker, I. Raes, and C. Weiss, Regional distribution of glutathione reductase in the adult rat brain, Brain Res 200:474-477 (1980). 109. H. Smits, L. Boven, C. Pereira, J. Verhoef, and H. Nottet, Role of macrophage activation in the pathogenesis of Alzheimer’s disease and human immunodeficiency virus type I-associated dementia, Euro J Clin Invest 30:526-535 (2000). 110. H. Fox, M. Weed, S. Huitron-Resendiz, et al., Antiviral treatment normalizes neurophysiological but not movement abnormalities in simian immunodeficiency virus-infected monkeys, J Clin Invest 106:37-45 (2000). 111. M. Kaul, G. Garden, and S. Lipton, Pathways to neuronal injury and apoptosis in HIV-associated dementia, Nature 410:988-994 (2001). 112. G. Chinetti, S. Griglio, M. Antonucci, et al., Activation of proliferator-activated receptors alpha and gamma induces apoptosis of human monocyte-derived macropages, J Biol Chem 273:25573-25580 (1998). 113. P. Delerive, K. De Bosscher, S. Besnard, et al., Peroxisome proliferated-activated receptor alpha negatively regulates the vascular inflammatory gene response by negative cross-talk with transcription factors NF-kappaB and AP-1, J Biol Chem 274:32048-32054 (1999). 114. L, Michalik, and W. Wahli, Peroxisome proliferator-activated receptors: three isotypes for a multitude of functions, Curr Opin Biotech 10:564-570 (1999). 115. N. Marx, G. Sukhova, T. Collins, P. Libby, and J. Plutzky, PPARalpha activators inhibit cytokine-induced vascular cell adhesion molecule-1 expression in human endothelial cells, Circulation 99:3125-3131 (1999). 116. I. Rusyn, C. Bradham, L. Cohn, et al., Corn oil rapidly activates nuclear factorkappaB in hepatic Kupffer cells by oxidant-dependent mechanisms, Carcinogenesis 20:2095-2100 (1999). 117. R. De Caterina, J. Liao, and P. Libby, Fatty acid modulation of endothelial activation, Am J Clin Nutr 71:213S-223S (2000). 118. P. Delerive, C. Furman, E. Teissier, J.-C. Fruchart, P. Duriez, and B. Staels, Oxidized phosopholipids activate PPARalpha in a phospholipase A2-dependent manner, FEBS Letters 471:34-38 (2000). 119. P. Libby, P. M. Ridker, and A. Maseri, Inflammation and atherosclerosis, Circulation 105:1135-43 (2002).

276

Chapter 13

120. D. N. Reeds, B. Mittendorfer, B. W. Patterson, W. G. Powderly, K. E. Yarasheski, and S. Klein, Alterations in lipid kinetics in men with HIV-dyslipidemia, Am J Physiol Endocrinol Metab 285:E490-7 (2003). 121. M. C. Sugden, K. Bulmer, G. F. Gibbons, B. L. Knight, and M. J. Holness, Peroxisome-proliferator-activated receptor-alpha (PPARalpha) deficiency leads to dysregulation of hepatic lipid and carbohydrate metabolism by fatty acids and insulin, Biochem J 364:361-8 (2002). 122. F. Sabri, K. Titanji, A. De Milito, and F. Chiodi, Astrocyte activation and apoptosis: their roles in the neuropathology of HIV infection, Brain Pathol 13:84-94 (2003). 123. Y. Sei, Y. Kustova, Y. Li, H. Morse, 3rd, P. Skolnick, and A. Basile, The encephalopathy associated with murine acquired immunodeficiency syndrome, Ann New York Acad Sci 840:822-834 (1998). 124. P. Eriksson, E. Perfilieva, T. Björk-Eriksson, et al., Neurogenesis in the adult human hippocampus, Nature Med 4:1313-1317 (1998). 125. J. Frisén, C. Johansson, C. Lothian, and U. Lendahl, Central nervous system stem cells in the embryo and adult, Cell Molec Life Sci 54:935-945 (1998). 126. E. Gould, and P. Tanapat, Stress and hippocampal neurogenesis, Biol Psych 46:1472-1479 (1999). 127. J. Parent, and D. Lowenstein, Mossy fiber reorganization in the epileptic hippocampus, Curr Opin Neurol 10:103-109 (1997). 128. A. Alvarez-Buylla, J. M. Garcia-Verdugo, and A. D. Tramontin, A unified hypothesis on the lineage of neural stem cells, Nat Rev Neurosci 2:287-93 (2001). 129. D. Kornack, and P. Rakic, Cell proliferation without neurogenesis in adult primate neocortex, Science 294:2127-2130 (2001). 130. H. van Praag, A. Schinder, B. Christie, N. Toni, T. Palmer, and F. Gage, Functional neurogenesis in the adult hippocampus, Nature 415:1030-1034 (2002). 131. W. Rzeski, L. Turski, and C. Ikonomidou, Glutamate antagonists limit tumor growth, Proc Natl Acad Sci U S A 98:6372-6377 (2001). 132. T. Takano, J. Lin, G. Arcuino, Q. Gao, J. Yang, and M. Nedergaard, Glutamate release promotes growth of malignant gliomas, Nat Med 7:1010-1015 (2001). 133. K. Selmaj, M. Farooq, W. Norton, C. Raine, and C. Brosnan, Proliferation of astrocytes in vitro in response to cytokines: A primary role for tumor necrosis factor, J Immunol 144:129-135 (1990). 134. D. Giulian, J. Woodward, D. Young, J. Krebs, and L. Lachman, Interleukin-1 injected into mammalian brain stimulates astrogliosis and neovascularization, J Neurosci 8:2485-2490 (1988). 135. A. Chenn, and C. A. Walsh, Regulation of cerebral cortical size by control of cell cycle exit in neural precursors, Science 297:365-9 (2002). 136. M. Groszer, R. Erickson, D. Scripture-Adams, et al., Negative regulation of neural stem/progenitor cell proliferation by the Pten tumor suppressor gene in vivo, Science 294:2186-2189 (2001). 137. L. Cantley, and B. Neel, New insights into tumor suppression: PTEN suppresses tumor formation by restraining the phosphoinositide 3-kinase/AKT pathway, Proc Natl Acad Sci U S A 96(8):4240-5 (1999).

Chapter 14 Neuronal Energy Metabolism in Brain: Astrocyte as both Metabolic “Buffer” and Mediator of Neuronal Injury

14.1.

INTRODUCTION

The metabolic interactions discussed above suggest that astrocytes support neuronal function by providing substrates required for generation of both neurotransmitter amino acids and ATP, as well as through their neuroprotective antioxidant function. Astrocytes thereby buffer neuronal dependence on, and vulnerability to, concentrations of essential substrates in plasma, which could be compromised by factors affecting systemic metabolism and/or local delivery. For example, availability of ketone bodies within the interstitial compartment reflects not only uptake from plasma (largely determined by plasma ketone body concentration) but also in situ generation by astrocytes (largely determined by plasma FFA concentrations, plasma FFA:albumin molar ratio, and activity of astrocyte fatty acid and ketogenesis pathways). Unfortunately, this interaction may also be deleterious. The potential importance of these interactions is illustrated by consideration of the following physiological and pathological phenomena, and is developed more extensively in the subsequent discussion of Alzheimer Disease (Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors).

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14.2. ACTIVATION-RELATED GLUCOSE AND OXYGEN CONSUMPTION Regional increases in blood flow induced by neuronal activation result in increased flow-dependent glucose extraction and consumption, whereas uptake of FFA and of ketone bodies, being relatively flowindependent, is therefore more constant. In terms of neuronal substrate supply, these contrasting responses to changes in perfusion imply that the ratio of glucose availability to that of systemically or locally produced ketone bodies is lower for resting, non-activated neurons and higher for activated neurons. Activation-induced increases in the ratio of glucose utilization (and of glutamine-derived 2-oxo-glutarate) to that of ketone bodies by the neuron would augment and thus the rate and efficiency of neuronal ATP generation. Several studies have suggested that, despite the activation-induced increase in neuronal glucose utilization, oxygen consumption (i.e., cerebral metabolic rate of oxygen — remains relatively constant during this process1-3; this results in an activation-induced dissociation between utilization of glucose and that of oxygen. Other reports have questioned this interpretation, suggesting that: 1) the apparent dissociation of glucose and oxygen consumption is transient, not accounted for by lactate, and is compensated for after stimulation (data based on cerebral arteriovenous concentration differences and immediate in situ brain freezing)4; or 2) that oxygen consumption increases even before cerebral blood flow (CBF) (data based on direct measurement of activity-dependent changes in oxygen tension)5. Although this question remains controversial6-9, several recent studies employing functional MRI10-12 or PET13 demonstrate that the increase in induced by metabolic activity is indeed proportionately less than the increase in CBF and, therefore, in glucose uptake. Although some uncertainty persists, such a dissociation would not be unexpected, given that the yield of ATP per molecule of consumed is greater with glucose and/or glutamine than with ketone bodies, as discussed above (see Chapter12.3.: Ketone Bodies). Indeed, the findings are consistent with the concept that activation-related, glucose-supported regional rates of oxygen consumption are lower than would be required for comparably increased ATP generation through oxidation of ketone bodies. The “discrepancy”, then, would reflect not a shift to anaerobic glycolysis or delayed oxidation of glycolysis-derived intermediates, but rather the increase in efficiency of aerobic metabolism (thus, relatively constant oxygen consumption) that would be expected of a shift to more efficiently utilized substrate that results from increased glucose

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utilization. The converse is of equal importance. Thus, oxygen consumption is relatively constant despite decreased glucose utilization, e.g., as is documented to occur in Alzheimer disease (see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors). This circumstance would be analogous to the cessation of neuronal activation with return to basal levels of regional blood flow and glucose uptake. It suggests that the non-activated neuron preferentially utilizes the energetically less efficient ketone bodies derived from astrocytes (see Chapter 12.3.: Ketone Bodies). An alternative hypothesis is that much of the increase in glucose utilization is directed toward the relatively oxygen-independent de novo synthesis of glutamate14,15; these interpretations would not appear to be mutually exclusive.

14.3. INCREASES IN PLASMA CONCENTRATIONS OF KETONE BODIES AND FFA In starvation, FFA mobilization from adipose tissue and hepatic ketogenesis are augmented; plasma concentrations of ketone bodies and FFA are increased, while that of glucose is decreased. As a result, brain extraction of ketone bodies and FFA, being directly proportional to their plasma concentrations, also increases, both absolutely and relative to that of glucose. Increased extraction of ketone bodies and their preferential neuronal utilization would result in a corresponding pyruvate dehydrogenase-mediated suppression of glucose utilization (Figs. 14.1 and 14.2). FFA, in contrast, are utilized primarily by astrocytes, which convert them at least in part to ketone bodies. The latter would be exported and rapidly utilized preferentially by neurons along with plasma-derived ketone bodies. Increased utilization of ketone bodies, from either or both souces as substrate would render generation of neuronal metabolic energy less efficient, implying that neuronal activation under these conditions would be suboptimal. Elevation of plasma FFA may also be more sustained, as occurs in association with HIV infection, obesity, insulin resistance, and diabetes mellitus16-18. As in starvation, brain uptake of FFA, astrocyte fatty acid oxidation and ketogenesis, and the supply of ketone bodies and diffusible oxidants (e.g., to neurons would also increase. Fatty acid and ketone body oxidation-induced, PDH-mediated suppression of brain glucose utilization (Figs. 14.1 and 14.2) would represent a local equivalent of “insulin resistance”, while neuronal utilization of ketone bodies as fuel would compromise mitochondrial energetics and efficiency. Chronic exposure to such conditions may promote ROS

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Figure 14.1. Fatty acid and ketone body oxidation in mitochondria suppresses glycolysis. Mitochondrial of fatty acids (astrocytes) and ketone bodies (neurons) generates three moieties which activate kinase-mediated feedback inhibition of PDH, i.e., acetyl CoA, ATP, and NADH. This effect, together with an accompanying suppression of glycolysis, decreases mitochondrial oxidation of pyruvate, and overall utilization of glucose. In peripheral tissues, especially muscle, this effect of mitochondrial fatty acid oxidation contributes to the “Randle cycle” inhibitory effect on glucose utilization and insulin resistance. Dashed lines: inhibition of PDH by acetyl CoA/CoA, ATP/ADP, and NADH/NAD. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; NADPH: nucleotide adenine triphosphate, reduced form; PDH: pyruvate dehydrogenase.

generation, impaired neuronal activation, oxidative stress-induced neuronal injury, and contribute to the pathogenesis of neurodegenerative processes such as HIV dementia and Alzheimer disease.

14.4.

KETOGENIC DIET AND CONTROL OF SEIZURE ACTIVITY

Seizure activity involves excessive, repetitive, and synchronous neuronal activation. These repeated cycles of membrane depolarization and repolarization obligate substantial increases in neuronal substrate utilization, and in the rates at which ATP is generated and consumed. Regional blood flow increases, as does utilization of glucose and, in this extreme circumstance, oxygen. Experimentally, it can be shown that


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cerebral ATP concentrations and ATP/ADP ratios fall, but these changes are greatly ameliorated if cerebral arterial oxygenation is maintained19, implying that oxygen and oxidative phosphorylation rather than substrate availability are limiting under these conditions. Thus, the seizure places an extreme demand on neuronal mitochondrial oxidative pathways and the electron transport chain. As the rate of ATP generation depends on it follows that initiation and perpetuation of seizure activity is necessarily linked to this key determinant of oxidative phosphorylation. For reasons discussed above, reliable support of generally requires provision of glycolysis-derived pyruvate to the citric acid cycle, reflecting its efficient generation of ATP and tightly regulated PDHmediated entry of acetyl CoA into the citric acid cycle (Fig. 3.2, facing p. 21). The resulting control of electron transport chain flux would tend to

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minimize “leakage”, ROS formation, and oxidative stress. Conversely, increased neuronal oxidation of ketone bodies, generated either in astrocytes20 or liver, would inhibit PDH, thereby suppressing glycolytic fueling of the electron transport chain (Fig. 14.1; also, see Chapter 12.3.: Ketone Bodies). High rates of ketone body oxidation may also exert uncoupling-like effects and promote oxidative stress. Both of these mitochondrial effects compromise and ATP generation, would be expected to diminish the likelihood of seizure initiation and to limit its duration, and may account for adverse cognitive effects of ketogenic weight-reducing21 and seizure-suppressing22 diets. Although this concept awaits experimental verification, it could largely account for seizure control by the ketogenic diet, the mechanism of which has remained unknown 23 . Ketone bodies also increase the abundance of the inhibitory neurotransmitter GABA in rat brain synaptosomes, an effect which if manifest in vivo would also limit seizure activity24. An alternative interpretation of the metabolic effects of ketone bodies has been proposed25.

14.5.

SYNAPTIC TRANSMISSION, LONG-TERM POTENTIATION, AND EXCITOTOXICITY

14.5.1. Energetic Considerations in Synaptic Transmission Most excitatory synaptic transmission in brain is mediated by glutamatergic neurons. In this process, activation-induced release of glutamate from the presynaptic nerve terminal increases its concentration in the synaptic cleft, resulting in the activation of three classes of glutamate receptor ion channels in the postsynaptic neuron26,27. These are named for the synthetic, physiologically irrelevant glutamate analogs that activate them experimentally, i.e., N-methyl-D-aspartate (NMDA), amino-3-hydroxy-5-methyl-4-isoxazolepropionate (AMPA), and kainate. Upon activation, channel opening permits movement of and/or down their electrochemical gradients into the postsynaptic neuron, which is thereby depolarized. Expenditure of metabolic energy is obligated in the subsequent restoration of neuronal membrane polarization, through increased activity of ATP-coupled membrane ion transporters. As discussed, generation of this ATP requires corresponding increases in citric acid cycle activity and oxidative phosphorylation. Precision in synaptic transmission is achieved in part by the brief duration and rapid repetition of each impulse28,29, thus permitting


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continuous modulation of neuronal activation and signal amplitude as may be dictated by changing circumstances. In contrast to the cholinesterase-mediated hydrolysis of acetyl choline in cholinergic neurotransmision, these characteristics are conferred in the case of glutamatergic transmission by rapid removal of glutamate from the synaptic cleft by adjacent astrocytes and neurons. The uptake process is mediated by specific plasma membrane glutamate transporters, largely and thus dependent on both glutamate concentration and transmembrane electrochemical gradient. Astrocytes do not undergo depolarization and would therefore be expected to take up glutamate at sustained high rates. Neuronal uptake, in contrast, would be expected to vary as influenced by activation-related changes in membrane potential. Rapid reduction of glutamate concentration in the synaptic cleft promotes ion channel closure, permitting postsynaptic neuronal repolarization and repeated subsequent depolarization events as dictated by the frequency of presynaptic neuronal re-activation and neurotransmitter release. Although depolarization of postsynaptic neurons results from the opening of glutamate-activated and channels, there is evidence that can be interpreted as suggesting that glutamate’s contribution to the overall depolarization/repolarization cycle is not limited to this initiating event. Thus, not only astrocytes but also postsynaptic neurons possess glutamate transporters26,30-40. Inhibitors of glutamate transporters induce neuronal death in vivo41-44, suggesting that these agents predominantly affect glutamate uptake by astrocytes. However, glutamate transporters differ in distribution and sensitivity to and hypoxia-ischemia46, thus pharmacological inhibition34,44,45 complicating interpretation of in vivo experiments with regard to the inhibitors’ primary site(s) of action. Carrier-mediated glutamate entry into the cytosol of the postsynaptic neuron is presumably driven by the resultant of the transmembrane electrochemical gradient and synaptic cleft glutamate concentration40. Although morphological data suggested that the specifically neuronal transporter EAAC1 was principally localized in non-synaptic regions of the postsynaptic neuron 32 , other evidence indicates a primarily dendrosomatic localization37,38,40. Indeed, postsynaptic neuronal uptake may contribute significantly to removal of glutamate from the synaptic cleft31,34, and thereby to ion channel closure. Neuronal uptake of glutamate may also be mediated by a glutamate-cystine exchanger, with potentially significant effects on neuronal energy and redox balance47 (also, see this Chapter 14.5.2.1.: Long-Term Potentiation, and 14.5.2.2.: Excitotoxicity — Origins of Oxidative Stress). Furthermore, it is of particular interest and importance that glutamate uptake may subserve an

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additional function within the postsynaptic neuron itself. This function, involving a direct relationship to the energetics of neuronal repolarization, potentially provides insight into both the mechanism of long-term potentiation48 and the pathogenesis of “excitotoxicity”, i.e., glutamate-induced neuronal injury49,50. Neuronal utilization of glutamate largely reflects either its glutamate decarboxylase-mediated biotransformation to GABA (perhaps unlikely in the postsynaptic nerve terminal) or its conversion to the citric acid cycle intermediate 2-oxo-glutarate by glutamate dehydrogenase or 51 aminotransferases . In astrocytes, glutamate metabolism is largely partitioned between either glutamine synthesis or entry (as 2-oxoglutarate) into the citric acid cycle, the latter increasing in proportion to extracellular glutamate concentration52,53; presumably pathways of metabolism in neurons would also exhibit such concentration dependence. Glutamate may also modulate VDAC conductivity54. To the extent that glutamate-derived 2-oxo-glutarate utilization increases neuronal citric acid cycle activity39, electron transport chain flux and oxidative phosphorylation would also increase. Thus, glutamate in the synaptic cleft effects ion channel-mediated depolarization of the postsynaptic neuron, while glutamate uptake into postsynaptic neuronal mitochondria would potentially contribute to generation of the ATP required for neuronal repolarization. Because of glutamate-induced increases in activity of the citric acid cycle and electron transport chain, PDH would be relatively inactivated, effecting compensatory decreases in glucose and pyruvate utilization. This would serve to optimize efficiency of the system by controlling substrate utilization and thus the flow of electrons into the electron transport chain, thereby minimizing electron “leakage” that would otherwise result in ROS generation and oxidative stress. Under normal conditions, therefore, a smoothly operating reciprocal modulation of glucose and glutamate utilization could be envisioned, fueling ATP generation in the postsynaptic neuron while responsive to the activity of synaptic transmission, and minimizing the generation of ROS and oxidative stress. The reported regional correlation between glucose utilization and glutamate abundance55 would be consistent with such an interaction. It is of considerable interest in this context that, unlike the excitatory neurotransmitter glutamate, the inhibitory neurotransmitter GABA is also transported into the neuron where it not only diminishes utilization of glutamate in energy generation56 but is itself poorly utilized in the citric acid cycle24. The fact that the effects of these “excitatory” and “inhibitory” neurotransmitters on postsynaptic neuronal activity appear to parallel their support of postsynaptic neuronal energy-


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generating metabolic pathways suggests that their consequences may also be causally linked. That is, that “excitatory” and “inhibitory” neurotransmission may at least in part result from direct interaction of the neurotransmitter with, and utilization in, postsynaptic neuronal energy generation. If validated, this concept may contribute to an understanding of two phenomena in which the postsynaptic neuron is exposed to greater than usual amounts of glutamate, i.e., long term potentiation (LTP) and experimental excitotoxicity.

14.5.2. Augmented Glutamate Exposure: Long Term Potentiation and Excitotoxicity Despite their strikingly divergent effects on neuronal function, current evidence is consistent with the possibility that both of these phenomena reflect postsynaptic neuronal energetic responses to increased glutamate exposure, with linkage in both cases to wg/Wnt-like signaling, but with critically important differences in the intensity, rate, and duration of the exposure. Thus, LTP results from more frequently repeated physiological presynaptic glutamate discharges over time, whereas excitotoxicity is induced by acute and potentially lethal exposure to unphysiologically high glutamate concentrations. In LTP, timedependent postsynaptic neuronal adaptive processes may be activated in response to metabolic changes; in acute excitotoxicity such adaptative responses are precluded. 14.5.2.1. Long-Term Potentiation.

Repeated physiological synaptic activity would increase the frequency with which glutamate concentration is elevated in the synaptic cleft. As discussed above, this would increase the frequency of physiological glutamate entry into the postsynaptic neuron, providing increased amounts of 2-oxo-glutarate to the citric acid cycle. To the extent that the resulting increase in citric acid cycle activity exceeds cell energy demands, high and accumulation of ATP would be reflected in a decreased availability of ADP, constraining the rate at which electrons in the electron transport chain could be consumed in oxidative phosphorylation, and thereby promoting electron “leakage” and generation of ROS57-59 (also, see Chapter 7.4.2.2.: Paradoxical Benefits of Adversity). Glutamate inhibition of neuronal cystine uptake, and therefore of glutathione synthesis47, would aggravate the adverse consequences of this potentially ROS- and oxidative stress-inducing effect. Such circumstances are dealt with in other cell types through a

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variety of protective mechanisms, which may involve activation of antioxidant defenses such as glutathione synthesis and superoxide dismutase expression. In the poorly antioxidant-defended neuron, however, these measures are less readily available (see Chapter 13.4.: Fatty Acids, Antioxidant Defense, HIV Dementia, and Cell Proliferation). Other less direct responses may appear to be energetically wasteful but also decrease the abundance of electrons in the electron transport chain. For example, in several cell types including the brown adipocyte, myocyte, and hepatocyte, mitochondrial uncoupling proteins (UCP) may be expressed. As noted, UCP2 expression has also been demonstrated in hypothalamic neurons involved in neuroendocrine and metabolic regulation60-63 (see Chapter 12.4.3.: Fatty Acid Oxidation and Regulation of Feeding Behavior). These, and uncouplers of oxidative phosphorylation in general, decrease ROS formation by accelerating the removal of electrons from the electron transport chain, permitting their reaction directly with protons and oxygen to form As this process bypasses ATP synthase, the electrochemical energy that would otherwise be utilized to synthesize ATP is diverted to thermal energy instead64-68. Similar to UCP, Enrichment of the mitochondrial membrane with fatty acids also increases its permeability to protons69 (also, see Chapter 4.3.: Omega-3 Fatty Acids), conferring neuroprotection70,71 and preventing impairment of LTP, e.g., by radiation-induced ROS generation72. For most brain neurons under normal conditions, however, antioxidant defenses are limited, and neither the rise in temperature61,63,73 nor the compromised that necessarily result from uncoupling of oxidative phosphorylation would seem compatible with the stringent physicochemical requirements for optimal control of neruonal activation and synaptic function. Thus, an alternative mechanism, in which increased expenditure of ATP were utilized to support a physiologically beneficial activity, would seem preferable. Increased expression of the glutamate-sensitive AMPA receptor(AMPAR)-activated channel, a characteristic of long term 48,74,75 , would accomplish this objective. Relative to potentiation postsynaptic neurons expressing NMDA receptors only, synaptic depolarization of postsynaptic neurons which in addition express AMPAR would increase transmembrane ion flux (excitatory postsynaptic current - EPSC). As a corollary, their repolarization would obligate a correspondingly greater expenditure of energy as ATP, thereby accelerating flux through an otherwise relatively stagnant electron transport chain. This concept is consistent with other evidence of the importance of mitochondrial energetic integrity in the process76. If the concept is correct, such neurons should be capable of attaining maximal


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rates of ATP generation that exceed those of non-adapted neurons. Moreover, NMDA-induced entry would serve to activate not only the protein kinase II (CaMKII)48,77 and wg/Wnt-like PI3K/Akt/PKB signaling78, but also citric acid cycle-associated dehydrogenases, thereby accelerating NADH formation, electron transport chain flux and ATP generation79,80. This hypothetical formulation, in which the phenomenon of long term potentiation is addressed in terms of the metabolic and energetic activity and demands of the postsynaptic neuron, appears to be consistent with current understandings of LTP, is compatible with the recently recognized importance of protein kinase A-mediated phosphorylation of AMPA subunits in synaptogenesis81, and would satisfy LTP requirements for CaMKII, and increased AMPAR-mediated EPSC48,74,82. Moreover, it is consistent with recent evidence that astrocyte-released is important in AMPAR expression and maintenance of synaptic strength83; the latter may be related to the energetic effects of TNFR1induced activation of wg/Wnt-like signaling84 (also, see Chapter 15.4.2.: Wingless/Wnt-Like Signal Transduction and Effects of Lithium, and Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). Further, these effects are consistent in the aggregate with recent evidence that wg/Wnt-like PI3K/Akt/PKB signaling is required in synaptic strength85 and LTP86,87, involving its direct interaction with AMPAR in the latter’s synaptic membranous insertion88. Given these observations, it may seem difficult to reconcile the overall importance of insulin, wg/Wnt-like signaling, and glucose utilization in cognitive function (see Chapter 15: Pathogenesis of Alzheimer Disease: Metabolic Factors), with evidence that insulin induces endocytotic removal of membrane AMPAR89-92. Perhaps this apparent contradiction may be reconciled if insulin and other activators of wg/Wnt-like signaling such as integrins93,94 serve energetically to support sustained cycling of AMPAR, rather than either its membranous insertion or removal alone. Overall, the present formulation is compatible with several recognized characteristics of LTP. Thus: 1) LTP is associated with an NMDA receptor-dependent increase in translocation of the EAAC1 glutamate transporter from cytosol to plasma membrane as well as in glutamate uptake40; 2) AMPAR turnover is rapid and involves multiple compartments89,95,96; 3) phosphorylation of glutamate receptor-gated ion channels enhances ESPC97; 4) NMDA receptor subtype populations differentially influence the process98 and may be subject to negative regulation by H-Ras99; and 5) ROS, e.g., as would be generated by the above hypothesized glutamate-induced increase in electron transport chain flux, both contribute to LTP100 and activate wg/Wnt-like

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PI3K/Akt/PKB signaling, whereas increased fatty acid abundance, e.g., in obesity101 may disrupt this pathway (see Chapters 3.3.2.: Wingless/WntLike Signaling: Convergence of Antecedents and the Unpredictable; 3.3.2.3.: Reactive Oxygen Species; and 7.4.2.2.: Paradoxical Benefits of Adversity). The formulation should be amenable to experimental testing. 14.5.2.2. Excitotoxicity – Origins of Oxidative Stress Excitotoxicity, in contrast to long term potentiation, results from acute neuronal exposure to concentrations of glutamate that are unphysiologically high, as a result of which adaptation is necessarily limited or precluded. It is characterized by apoptotic or necrotic neuronal death caused by altered mitochondrial function and oxidative stress, accompanied by changes in neuronal metabolism102-104, and 105 efflux of glutathione . The initiating exposure to glutamate or NMDA opens the NMDA-activated glutamate receptor-associated channel, and excitotoxicity is prevented or ameliorated experimentally under conditions in which extracellular is low or the NMDA receptor is 64,102,106 blocked . The pathogenesis of this complex process, however, remains incompletely understood. Thus, neuronal sensitivity to glutamate toxicity is modulated by other substances in serum107. Further, the commonly employed NMDA receptor antagonist MK801 also inhibits monoamine transporters108, implying that one or more other actions may complicate interpretation of its protective effect. Moreover, accumulating evidence indicates that factors other than influx are involved26,107,109,111. For example, changes in bulk cytoplasmic free appear to depend on interaction with intracellular stores, especially mitochondrial, do not necessarily directly reflect entry via glutamate-activated ion 112-115 , and do not necessarily correlate with the severity of channels injury113,115,116. In addition, cell death and extracellular have been dissociated in a model of thapsigargin-induced apoptosis in the neuronal GT1-7 cell line117, NMDA receptor-mediated channel activation has been specifically excluded in a model of glutamate toxicity in PC12 cells110, and increased was observed to follow rather than precede glutathione depletion induced by diminished cystine uptake and oxidative stress47. To the extent that does contribute to this process, its effects may be related in part to enhanced activity of citric acid cycle-associated dehydrogenases, which promote NADH generation and electron transport chain flux as discussed above79,80 (also, see this Chapter 14.5.2.1.: LongTerm Potentiation). This would be consistent with evidence that pyruvate


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and malate, which provide regulated entry of fuel into the citric acid cycle, protect against glutamate excitotoxicity and decrease by 118,119 . A role for oxidative stress increasing its mitochondrial sequestration in excitotoxicity is supported by the results of numerous studies26,47,102,110,120; it is not apparently attributable to phospholipase A2mediated release of arachidonic acid, nor to formation of eicosanoid arachidonate metabolites64,106. Information concerning a contributory role of NOS is conflicting50,64, These diverse observations may be reconciled if the origin of oxidative stress and the pathogenesis of excitotoxicity were to reflect at least in part the consequences of excessive increases in citric acid cycle and electron transport chain activity. Such increases could be induced by the effects of glutamate, acting both as inhibitor of cystine uptake — and thus of both glutathione formation and antioxidant defense47 — and as citric acid cycle substrate (i.e., as 2-oxo-glutarate), combined with those of acting as mitochondrial matrix dehydrogenase activator79,80. Together, their augmentation of electron transport chain flux would promote ROS generation. Oxidative stress, moreover, would be augmented by glutamate-induced depletion of glutathione in both mitochondria and cell47,105,121. Oxidative stress appears to increase while increases in both ROS generation and abundance would contribute to permeability transition pore opening, collapse of and cell death122,123. In light of these considerations, uncertainties concerning this complex pathogenesis may in part relect findings in studies of cellular responses to non-physiological NMDA-receptor ligands (e.g., NMDA itself), the impact of which on mitochondrial energetics would differ substantially from that of the pathophysiologically relevant glutamate. Although extramitochondrial processes could also generate ROS in this setting124, an intramitochondrial origin is supported by several studies64,106,125. This would also provide a mechanistic basis for the presumably NFKB-mediated126-128 sustained expression of COX2 in Direct dendritic arborizations of excitatory cortical neurons129. mitochondrial involvement in excitotoxicity is further supported by evidence that the relative resistance to NMDA excitotoxicity of cultured neurons from immature rats is unrelated to peak but instead depends on mitochondrial maintenance of ensuing homeostasis130. Finally, a mitochondrial origin is most consistent with the observation that the oxidative phosphorylation uncoupler FCCP prevented oxidative stress64. As discussed in the preceding section addressing Long-Term Potentiation, and below, the uncoupler in this setting would both decrease entry of entry into the

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mitochondrial matrix and accelerate removal of electrons through the 2electron reduction of oxygen to Together, these effects would decrease electron abundance at more proximal sites in the electron transport chain — complexes I and III, and ubiquinone — at which electron “leakage” with ROS formation is most prominent. Uncouplers of oxidative phosphorylation diminish mitochondrial ROS generation in diverse settings64-68, and this mechanism would also account for the neuroprotective effect of salicylate, a recognized uncoupler of oxidative phosphorylation131-133. Salicylate may also act as a trap for hydroxyl radicals134. It has been suggested that salicylate neuroprotection reflects suppression of an adverse effect of the 135 transcriptional regulator . However, it appears more likely that suppression of activation is not protective per se, but rather is a consequence of the suppressive effect of the uncoupling agent on the mitochondrial ROS generation, upon which activation may depend126-128. Depolarization of mitochondria with rotenone/oligomycin125 also diminishes electron “leakage” and superoxide generation, in this case by blocking electron entry into the electron transport chain at complex I. Conversely, and consistent with this general formulation, inhibition of the electron transport chain at complex III by antimycin A115,136 augments ROS generation through electron “leakage” from the “upstream” complex I and ubiquinone. Thus, while mitochondrial polarization (i.e., the maintenance of a significant is viewed as an indicator of increased likelihood of excitotoxic death, this may simply reflect:1) the necessarily higher electron transport chain flux upon which polarization depends, the propensity of which to generate ROS is inherently greater; and 2) the increased propensity of maximally energized mitochondria, in which ADP availability is diminished, to “leak” electrons and generate ROS57-59(also, see Chapter 7.4.2.2.: Paradoxical Benefits of Adversity). The relationship between glutamate-induced excitotoxicity and mitochondrial energetics proposed here is consistent with evidence that mitochondrial energetics are intimately linked to the wg/Wnt-like PI3K/Akt/PKB pathway in a concentration-dependent or graded manner, as discussed in connection with LTP (this Chapter 14.5.2.1.: Long-Term Potentiation; also, see Chapter 15.4.2.1.: Wingless/Wnt-Like Signaling). Thus, glutamate excitotoxicity may actually be suppressed by PI3Kmediated signaling initiated by NMDA137 and hepatocyte growth factor138, as well as by lithium137,139,140 which augments PI3K/Akt/PKB signaling through inhibition of As suggested (see this Chapter 14.5.1.: Energetic Considerations in Synaptic Transmission), utilization of glucose and glutamate may be


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normally balanced such that the potential for oxidative stress is minimized. This balance could be perturbed under various conditions in which altered rates of substrate utilization and/or other factors promote electron “leakage” from the electron transport chain, generating ROS. For example, excessive glutamate entry into the postsynaptic neuron may be driven by abnormally high concentrations in the synaptic cleft as the result of insufficient glutamate uptake by astrocytes141, a process which is inhibited by ROS142-144, by amyloid (25-35)145, and under certain conditions by Excess influx of glutamate may exceed neuronal energy needs and thus its capacity to utilize substrate for ATP generation. As noted above, the implied limitation in availability of ADP for oxidative phosphorylation would suppress electron transport chain flux, an effect which may also result from hypoxia or chemical inhibitors such as cyanide110. Glutamate may also directly modulate VDAC activity and thus ADP-ATP exchange54. The concept that excitotoxicity may result from disruption of mitochondrial energetics and electron transport chain-related ROS generation is further supported by evidence that inhibition of the complex II enzyme succinate dehydrogenase (and as a result the citric acid cycle) is sufficient to initiate excitotoxicity as well as to increase sensitivity to glutamate147-150. Further, the concept is consistent with available information concerning the complex interactions among several key areas and issues. These include: 1) mitochondrial energetics and ROS generation; 2) ATP abundance, and extra- and intracellular sources 110,115,151,152 of cytosolic ; 3) the fact that glutamate excitotoxicity may occur independent of neuronal depolarization153; and 4) the temporally distinctive clinical expressions of the process in acute vs. chronic neurodegenerative disorders49,50. Moreover, antioxidants are protective in this setting152,154, including an apparently antioxidant effect of the activator-ligand troglitazone 155,156. In the foregoing examples, oxidative stress is depicted as the consequence of an increased “leakage” of electrons from an overloaded electron transport chain. The overload reflects a discrepancy between the rate at which electrons are supplied, primarily by the citric acid cycle, and the rate at which they are consumed in the cytochrome oxidase-mediated 2-electron reduction of oxygen. The discrepancy may be caused by either or both of two general perturbations, i.e., excessive electron input (e.g., derived from excessive glutamate fueling of the citric acid cycle as 2-oxo-glutarate), or impaired electron removal through the normal 2electron reduction of oxygen (e.g., as the consequence of hypoxia, ADP depletion, or other factors which inhibit the electron transport chain or oxidative phosphorylation).

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Importantly, overload of the electron transport chain could theoretically be caused as well by excessive utilization of any substrate, entry of which into the citric acid cycle is not subject to control of the kind that is exerted upon pyruvate utilization (and thus upon glucose or lactate utilization) by PDH. In addition to glutamate and its congeners, ketone bodies are potentially such substrates. As discussed above, mitochondrial oxidation of ketone bodies, and thus their provision of electrons to the electron transport chain, are relatively unregulated. Although ketone bodies may be derived from plasma, it is likely that astrocyte ketogenesis, fueled by plasma-derived FFA, is more directly relevant to neuronal oxidative stress, and could reflect sustained increases in plasma FFA, e.g., in obesity, insulin resistance, diabetes mellitus, or HIV infection (see Chapter 13.4.2.: HIV Dementia). Consistent with such a mechanism, increased expression of 2 FABP isoforms in hippocampal glial cells following systemic kainate administration157 imply activation of a cellular response to increased fatty acid exposure. Moreover, sustained exposure to excessive amounts of ketone bodies would be expected to act in concert with excess glutamate in contributing to chronic oxidative stress and the evolution of chronic neurodegenerative disease. Furthermore, as FFA inhibit glutamate transport by both 158,159 , impaired removal of glutamate from synaptosomes and astrocytes the synaptic cleft under these conditions may disproportionately affect the postsynaptic neuron. In addition to these factors, and as discussed above (see Chapter 13.4.: Fatty Acids, Antioxidant Defense, HIV Dementia, and Cell Proliferation), sustained increases in plasma [FFA] may generate oxidative stress in astrocytes, impairing glutamate uptake from the synaptic cleft142-144 and predisposing to excitotoxicity. Finally, in view of the above-noted cannabinoid receptor-mediated stimulation of astrocyte ketogenesis and oxidative stress160-162 (also, see Chapter 13.4.: Fatty Acids, Antioxidant Defense, HIV Dementia, and Cell Proliferation), it is of potentially major significance that astrocytes also produce anandamide and related endocannabinoids163, and that blockade of the receptor is protective against NMDA-induced excitotoxicity164. In contrast, activation of the same receptor by the xenobiotic cannabinoid tetrahydrocannabinol acts via PI3K/Akt/PKB and possibly ERK signaling to protect astrocytes from ceramide-induced apoptosis165. It seems likely that the diverse cellular responses elicited by various cannabinoids162 may be to some extent cell-specific. However, they also may reflect differences in biotransformation dictated by the fatty acid amide hydrolase and hydrolyzable fatty acid amide moiety that are important in the regulation of endocannabinoid signaling166; the potential for oxidation of the fatty


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acid moiety167,168 that is present in endocannabinoids but not in tetrahydrocannabinol may also be significant in this regard.

14.6. REFERENCES 1. P. Fox, M. Raichle, M. Mintun, and C. Dence, Nonoxidative glucose consumption during focal physiologic neural activity, Science 241:462-464 (1988). 2. P. Madsen, S. Hasselbalch, L. Hagemann, et al., Persistent resetting of the cerebral oxygen/glucose uptake ratio by brain activation: evidence obtained with the KetySchmidt technique, J Cerebral Blood Flow Metab 15:485-491 (1995). 3. F. Hyder, D. Rothman, G. Mason, A. Rangarajan, K. Behar, and R. Shulman, Oxidative glucose metabolism in rat brain during single forepaw stimulation: a spatially localized 1H13C nuclear magnetic resonance study, J Cerebral Blood Flow Metab 17:1040-1047 (1997). 4. P. Madsen, N. Cruz, L. Sokoloff, and G. Dienel, Cerebral oxygen/glucose ratio is low during sensory stimulation and rises above normal during recovery: excess glucose consumption during stimulation is not accounted for by lactate efflux from or accumulation in brain tissue, J Cerebral Blood Flow Metab 19:393-400 (1999). 5. I. Vanzetta, and A. Grinvald, Increased cortical oxidative metabolism due to sensory stimulation: implications for functional brain imaging, Science 286:1555-8 (1999). 6. R. Buxton, The elusive initial dip, Neuroimage 13:953-958 (2001). 7. M. Mintun, B. Lundstrom, A. Snyder, A. Vlassenko, G. Shulman, and M. Raichle, Blood flow and oxygen delivery to human brain during functional activity: theoretical modeling and experimental data, Proc Natl Acad Sci U S A 98:6859-6864 (2001). 8. R. Shulman, F. Hyder, and D. Rothman, Cerebral energetics and the glycogen shunt: neurochemical basis of functional imaging, Proc Natl Acad Sci U S A 98:6417-6422 (2001A). 9. R. Shulman, F. Hyder, and D. Rothman, Lactate efflux and the neuroenergetic basis of brain function, NMR Biomed 14:389-396 (2001B). 10. R. Hoge, J. Atkinson, B. Gill, G. Crelier, S. Marrett, and G. Pike, Linear coupling between cerebral blood flow and oxygen consumption in activated human cortex, Proc Natl Acad Sci USA 96:9403-9408 (1999A). 11. R. Hoge, J. Atkinson, B. Gill, G. Crelier, S. Marrett, and G. Pike, Investigation of BOLD signal dependence on cerebral blood flow and oxygen consumption: the deoxyhemoglobin dilution model, Magnetic Resonance Med 42:849-863 (1999B). 12. J. Oja, J. Gillen, R. Kauppinen, M. Kraut, and P. van Zijl, Determination of oxygen extraction ratios by magnetic resonance imaging, J Cerebral Blood Flow Metab 19:1289-1295 (1999). 13. M. Vafaee, and A. Gjedde, Model of blood-brain transfer of oxygen explains nonlinear flow-metabolism coupling during stimulation of visual cortex, J Cerebral Blood Flow Metab 20:747-754 (2000). 14. L. Hertz, R. Dringen, A. Schousboe, and S. Robinson, Astrocytes: glutamate producers for neurons, J Neurosci Res 57:417-428 (1999). 15. L. Hertz, A. Yu, G. Kala, and A. Schousboe, Neuronal-astrocytic and cytosolicmitochondrial metabolite trafficking during brain activation, hyperammonemia and energy deprivation, Neurochem Intl 37:83-102 (2000).

294

Chapter 14

16. E. Fraze, C. Donner, A. Swislocki, Y. Chiou, Y. Chen, and G. Reaven, Ambient plasma free fatty acid concentrations in noninsulin-dependent diabetes mellitus: evidence for insulin resistance, J Clin Endocrinol Metab 61:807-811 (1985). 17. Y. Chen, A. Golay, A. Swislocki, and G. Reaven, Resistance to insulin suppression of plasma free fatty acid concentrations and insulin stimulation of glucose uptake in noninsulin-dependent diabetes mellitus, J Clin Endocrinol Metab 64:17-21 (1987). 18. A. Golay, A. Swislocki, Y. Chen, and G. Reaven, Relationships between plasma-free fatty acid concentration, endogenous glucose production, and fasting hyperglycemia in normal and non-insulin-dependent diabetic individuals, Metab Clin Exp 36:692696 (1987). 19. R. Young, O. Petroff, B. Chen, J. Gore, and W. Aquila, Brain energy state and lactate metabolism during status epilepticus in the neonatal dog: in vivo 31P and 1H nuclear magnetic resonance study, Ped Res 29:191-195 (1991). 20. T. E. Cullingford, D. A. Eagles, and H. Sato, The ketogenic diet upregulates expression of the gene encoding the key ketogenic enzyme mitochondrial 3-hydroxy3-methylglutaryl-CoA synthase in rat brain, Epilepsy Res 49:99-107 (2002). 21. R. Wing, J. Vazquez, and C. Ryan, Cognitive effects of ketogenic weight-reducing diets, Intl J Obesity Rel Metab Disord 19:811-816 (1995). 22. S. Su, M. Cilio, Y. Sogawa, D. Silveira, G. Holmes, and C. Stafstrom, Timing of ketogenic diet initiation in an experimental epilepsy model, Devel Brain Res 29125:131-138 (2000). 23. P. Schwartzkroin, Mechanisms underlying the anti-epileptic efficacy of the ketogenic diet, Epilepsy Research 37:171-180 (1999). 24. M. Erecinska, D. Nelson, Y. Daikhin, and M. Yudkoff, Regulation of GABA level in rat brain synaptosomes: fluxes through enzymes of the GABA shunt and effects of glutamate, calcium, and ketone bodies, J Neurochem 67:2325-2334 (1996C). 25. M. Yudkoff, Y. Daikhin, I. Nissim, A. Lazarow, and I. Nissim, Ketogenic diet, amino acid metabolism, and seizure control, J Neurosci Res 66:931-940 (2001). 26. S. Lipton, and P. Rosenberg, Excitatory amino acids as a final common pathway for neurologic disorders, New Engl J Med 330:613-622 (1994). 27. S. Ozawa, H. Kamiya, and K. Tsuzuki, Glutamate receptors in the mammalian central nervous system, Prog Neurobiol 54:581-618 (1998). 28. J. Lisman, Bursts as a unit of neural information: making unreliable synapses reliable, Trends Neurosci 20:38-43 (1997). 29. A. Riehle, S. Grün, M. Diesmann, and A. Aertsen, Spike synchronization and rate modulation differentially involved in motor cortical function, Science 278:19501953 (1997). 30. Y. Kanai, and M. Hediger, Primary structure and functional characterization of a highaffinity glutamate transporter, Nature 360:467-471 (1992). 31. M. Takahashi, M. Sarantis, and D. Attwell, Postsynaptic glutamate uptake in rat cerebellar Purkinje cells, J Physiol 497:523-530 (1996). 32. S. Coco, C. Verderio, D. Trotti, J. Rothstein, A. Volterra, and M. Matteoli, Nonsynaptic localization of the glutamate transporter EAAC1 in cultured hippocampal neurons, Eur J Neurosci 9:1902-1910 (1997). 33. A. Furuta, L. Martin, C. Lin, M. Dykes-Hoberg, and J. Rothstein, Cellular and synaptic localization of the neuronal glutamate transporters excitatory amino acid transporter 3 and 4, Neurosci 81:1031-1042 (1997). 34. T. Otis, M. Kavanaugh, and C. Jahr, Postsynaptic glutamate transport at the climbing fiber-Purkinje cell synapse, Science 277:1515-1518 (1997).


295

35. S. Eliasof, J, Arriza, B. Leighton, M. Kavanaugh, and S. Amara, Excitatory amino acid transporters of the salamander retina: identification, localization, and function, J Neurosci 18:698-712 (1998). 36. L. Gaal, B. Roska, S. Picaud, S. Wu, R. Marc, and F. Werblin, Postsynaptic response kinetics are controlled by a glutamate transporter at cone photoreceptors, J Neurophysiol 79:190-196 (1998). 37. G. Gegelashvili, and A. Schousboe, Cellular distribution and kinetic properties of high-affinity glutamate transporters, Brain Res Bull 45:233-238 (1998). 38. W. Fairman, and S. Amara, Functional diversity of excitatory amino acid transporters: ion channel and transport modes, American Journal of Physiology 277:F481-6 (1999). 39. K. Sims, and M. Robinson, Expression patterns and regulation of glutamate transporters in the developing and adult nervous system, Crit Rev Neurobiol 13:169197 (1999). 40. J. Levenson, E. Weeber, J. Selcher, L. Kategaya, J. Sweatt, and A. Eskin, Long-term potentiation and contextual fear conditioning increase neuronal glutamate uptake, Nat Neurosci 5:155-161 (2002). 41. C. Arias, I. Arrieta, L. Massieu, and R. Tapia, Neuronal damage and MAP2 changes induced by the glutamate transport inhibitor dihydrokainate and by kainate in rat hippocampus in vivo, Exp Brain Res 116:467-476 (1997). 42. A. Hirata, R. Nakamura, S. Kwak, N. Nagata, and K. Kamakura, AMPA receptormediated slow neuronal death in the rat spinal cord induced by long-term blockade of glutamate transporters with THA, Brain Res 771:37-44 (1997). 43. J. Liévens, M. Dutertre, C. Forni, P. Salin, and L. Kerkerian-Le Goff, Continuous administration of the glutamate uptake inhibitor L-trans-pyrrolidine-2,4dicarboxylate produces striatal lesion, Brain Research Molec Brain Res 50:181-189 (1997). 44. L. Massieu, and R. Tapia, Glutamate uptake impairment and neuronal damage in young and aged rats in vivo, J Neurochem 69:1151-1160 (1997). 45. J. Rothstein, M. Dykes-Hoberg, C. Pardo, et al., Knockout of glutamate transporters reveals a major role for astroglial transport in excitotoxicity and clearance of glutamate, Neuron 16:675-686 (1996). 46. L. Martin, A. Brambrink, C. Lehmann, et al., Hypoxia-ischemia causes abnormalities in glutamate transporters and death of astroglia and neurons in newborn striatum, Ann Neurol 42:335-348 (1997). 47. C. Pereira, and C. Oliveria, Oxidative glutamate toxicity involves mitochondrial dysfunction and perturbation of intracellular Ca2+ homeostasis, Neurosci Res 37:227-236 (2000). 48. R. Malenka, and R. Nicoll, Long-term potentiation - a decade of progress?, Science 285:1870-1874 (1999). 49. M. Beal, Does impairment of energy metabolism result in excitotoxic neuronal death in neurodegenerative illnesses?, Ann Neurol 31:119-130 (1992). 50. J. Schulz, R. Matthews, T. Klockgether, J. Dichgans, and M. Beal, The role of mitochondrial dysfunction and neuronal nitric oxide in animal models of neurodegenerative diseases, Molec Cell Biochem 174:193-197 (1997). 51. Z. Kovacevic, and J. McGivan, Mitochondrial metabolism of glutamine and glutamate and its physiological significance, Physiolog Rev 63:547-605 (1983). 52. M. McKenna, U. Sonnewald, X. Huang, J. Stevenson, Zielke, and HR, Exogenous glutamate concentration regulates the metabolic fate of glutamate in astrocytes, J Neurochem 66:386-393 (1996).

296

Chapter 14

53. M. McKenna, J. Stevenson, X. Huang, and I. Hopkins, Differential distribution of the enzymes glutamate dehydrogenase and aspartate aminotransferase in cortical synaptic mitochondria contributes to metabolic compartmentation in cortical synaptic terminals, Neurochem Intl 37:229-241 (2000). 54. D. Gincel, S. Silberberg, and V. Shoshan-Barmatz, Modulation of the voltagedependent anion channel (VDAC) by glutamate, J Bioenerg Biomembranes 32:571583 (2000). 55. Z. Pfund, D. Chugani, C. Juhász, et al., Evidence for coupling between glucose metabolism and glutamate cycling using FDG PET and 1H magnetic resonance spectroscopy in patients with epilepsy, J Cerebral Blood Flow Metab 20:871-878 (2000). 56. H. Qu, J. Konradsen, M. van Hengel, S. Wolt, and U. Sonnewald, Effect of glutamine and GABA on [U-(13)C]glutamate metabolism in cerebellar astrocytes and granule neurons, J Neurosci Res 66:885-890 (2001). 57. S. S. Korshunov, V. P. Skulachev, and A. A. Starkov, High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria, FEBS Lett 416:15-8 (1997). 58. S. S. Liu, Cooperation of a “reactive oxygen cycle” with the Q cycle and the proton cycle in the respiratory chain—superoxide generating and cycling mechanisms in mitochondria, J Bioenerg Biomembr 31:367-76 (1999). 59. I. Lee, E. Bender, and B. Kadenbach, Control of mitochondrial membrane potential and ROS formation by reversible phosphorylation of cytochrome c oxidase, Mol Cell Biochem 234-235:63-70 (2002). 60. D. Richard, S. Clavel, Q. Huang, D. Sanchis, and D. Ricquier, Uncoupling protein 2 in the brain: distribution and function, Biochem Soc Trans 29:812-817 (2001). 61. T. Horvath, C. Warden, M. Hajos, A. Lombardi, F. Goglia, and S. Diano, Brain uncoupling protein 2: uncoupled neuronal mitochondria predict thermal synapses in homeostatic centers, J Neurosci 19:10417-10427 (1999). 62. S. Diano, H. Urbanski, B. Horvath, et al., Mitochondrial uncoupling protein 2 (UCP2) in the nonhuman primate brain and pituitary, Endocrinol 141:4226-4238 (2000). 63. T. L. Horvath, S. Diano, and C. Barnstable, Mitochondrial uncoupling protein 2 in the central nervous system: neuromodulator and neuroprotector, Biochem Pharmacol 65:1917-21 (2003). 64. I. Reynolds, and T. Hastings, Glutamate induces the production of reactive oxygen species in cultured forebrain neurons following NMDA receptor activation, J Neurosci 15:3318-3327 (1995). 65. V. Skulachev, Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants, Quart Rev Biophys 29:169-202 (1996). 66. A. Nègre-Salvayre, C. Hirtz, G. Carrera, et al., A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation, FASEB J 11:809-815 (1997). 67. A. Stout, H. Raphael, B. Kanterewicz, E. Klann, and I. Reynolds, Glutamate-induced neuron death requires mitochondrial calcium uptake, Nature Neurosci 1:366-373 (1998). 68. S. Korshunov, O. Korkina, E. Ruuge, V. Skulachev, and A. Starkov, Fatty acids as natural uncouplers preventing generation of O2- and H2O2 by mitochondria in the resting state, FEBS Letts 435:215-218 (1998).


297

69. D. F. Rolfe, A. J. Hulbert, and M. D. Brand, Characteristics of mitochondrial proton leak and control of oxidative phosphorylation in the major oxygen-consuming tissues of the rat, Biochim Biophys Acta 1188:405-16 (1994). 70. Y. Ge, X. Wang, Z. Chen, N. Landman, E. H. Lo, and J. X. Kang, Gene transfer of the Caenorhabditis elegans n-3 fatty acid desaturase inhibits neuronal apoptosis, J Neurochem 82:1360-6 (2002). 71. D. S. Martin, P. E. Lonergan, B. Boland, et al., Apoptotic changes in the aged brain are triggered by interleukin-1beta-induced activation of p38 and reversed by treatment with eicosapentaenoic acid, J Biol Chem 277:34239-46 (2002). 72. P. E. Lonergan, D. S. Martin, D. F. Horrobin, and M. A. Lynch, Neuroprotective effect of eicosapentaenoic acid in hippocampus of rats exposed to gamma-irradiation, J Biol Chem 277:20804-11 (2002). 73. D. Yablonskiy, J. Ackerman, and M. Raichle, Coupling between changes in human brain temperature and oxidative metabolism during prolonged visual stimulation, Proc Natl Acad Sci USA 97:7603-7608 (2000). 74. D. Feldman, R. Nicoll, and R. Malenka, Synaptic plasticity at thalamocortical synapses in developing rat somatosensory cortex: LTP, LTD, and silent synapses, J Neurobiol 41:92-101 (1999). 75. J. Zhu, I. Esteban, Y. Hayashi, and R. Malinow, Postnatal synaptic potentiation: delivery of GluR4-containing AMPA receptors by spontaneous activity, Nature Neurosci 3:1098-1106 (2000). 76. E. J. Weeber, M. Levy, M. J. Sampson, et al., The role of mitochondrial porins and the permeability transition pore in learning and synaptic plasticity, J Biol Chem 277:18891-7 (2002). 77. J. Lisman, H. Schulman, and H. Cline, The molecular basis of CaMKII function in synaptic and behavioural memory, Nat Rev Neurosci 3:175-190 (2002). 78. M. Perkinton, J. Ip, G. Wood, A. Crossthwaite, and R. Williams, Phosphatidylinositol 3-kinase is a central mediator of NMDA receptor signalling to MAP kinase (Erk1/2), Akt/PKB and CREB in striatal neurones, J Neurochem 80:239254 (2002). 79. J. McCormack, and R. Denton, The role of intramitochondrial Ca2+ in the regulation of oxidative phosphorylation in mammalian tissues, Biochem Soc Trans 21:793-799 (1993). 80. O. Kann, S. Schuchmann, K. Buchheim, and U. Heinemann, Coupling of neuronal activity and mitochondrial metabolism as revealed by NAD(P)H fluorescence signals in organotypic hippocampal slice cultures of the rat, Neuroscience 119:87-100 (2003). 81. J. A. Esteban, S. H. Shi, C. Wilson, M. Nuriya, R. L. Huganir, and R. Malinow, PKA phosphorylation of AMPA receptor subunits controls synaptic trafficking underlying plasticity, Nat Neurosci 6:136-43 (2003). 82. Y. Hayashi, S. Shi, J. Esteban, A. Piccini, J. Poncer, and R. Malinow, Driving AMPA receptors into synapses by LTP and CaMKII: requirement for GluR1 and PDZ domain interaction, Science 287:2262-2267 (2000). 83. E. Beattie, D. Stellwagen, W. Morishita, et al., Control of synaptic strength by glial TNFalpha, Science 295:2282-2285 (2002). 84. O. Ozes, L. Mayo, J. Gustin, S. Pfeffer, L. Pfeffer, and D. Donner, NF-kappaB activation by tumour necrosis factor requires the Akt serine-threonine kinase, Nature 401:82-85 (1999). 85. Q. Wang, L. Liu, L. Pei, et al., Control of synaptic strength, a novel function of Akt, Neuron 38:915-28 (2003).

298

Chapter 14

86. P. Sanna, M. Cammalleri, F. Berton, et al., Phosphatidylinositol 3-kinase is required for the expression but not for the induction or the maintenance of long-term potentiation in the hippocampal CA1 region, J Neurosci 22:3359-3365 (2002). 87. P. Opazo, A. M. Watabe, S. G. Grant, and T. J. O’Dell, Phosphatidylinositol 3-kinase regulates the induction of long-term potentiation through extracellular signal-related kinase-independent mechanisms, J Neurosci 23:3679-88 (2003). 88. H. Y. Man, Q. Wang, W. Y. Lu, et al., Activation of PI3-kinase is required for AMPA receptor insertion during LTP of mEPSCs in cultured hippocampal neurons, Neuron 38:611-24 (2003). 89. J. Lin, W. Ju, K. Foster, et al., Distinct molecular mechanisms and divergent endocytotic pathways of AMPA receptor internalization, Nat Neurosci 3:1282-1290 (2000). 90. H. Y. Man, J. W. Lin, W. H. Ju, et al., Regulation of AMPA receptor-mediated synaptic transmission by clathrin-dependent receptor internalization, Neuron 25:649-62 (2000). 91. R. C. Carroll, E. C. Beattie, M. von Zastrow, and R. C. Malenka, Role of AMPA receptor endocytosis in synaptic plasticity, Nat Rev Neurosci 2:315-24 (2001). 92. I. Song, and R. L. Huganir, Regulation of AMPA receptors during synaptic plasticity, Trends Neurosci 25:578-88 (2002). 93. M. Delcommenne, C. Tan, V. Gray, L. Rue, J. Woodgett, and S. Dedhar, Phosphoinositide-3-OH kinase-dependent regulation of glycogen synthase kinase 3 and protein kinase B/AKT by the integrin-linked kinase, Proc Natl Acad Sci USA 95:11211-11216 (1998). 94. C. S. Chan, E. J. Weeber, S. Kurup, J. D. Sweatt, and R. L. Davis, Integrin requirement for hippocampal synaptic plasticity and spatial memory, J Neurosci 23:7107-16 (2003). 95. M. Ehlers, Reinsertion or degradation of AMPA receptors determined by activitydependent endocytic sorting, Neuron 28:511-525 (2000). 96. M. Sheng, and S. Hyoung Lee, AMPA receptor trafficking and synaptic plasticity: major unanswered questions, Neurosci Res 46:127-34 (2003). 97. T. Soderling, and V. Derkach, Postsynaptic protein phosphorylation and LTP, Trends Neurosci 23:75-80 (2000). 98. S. Hrabetova, P. Serrano, N. Blace, et al., Distinct NMDA receptor subpopulations contribute to long-term potentiation and long-term depression induction, J Neurosci 20:1-6 (2000). 99. C. Thornton, R. Yaka, S. Dinh, and D. Ron, H-Ras modulates N-methyl-D-aspartate receptor function via inhibition of Src tyrosine kinase activity, J Biol Chem 278:23823-9 (2003). 100. L. T. Knapp, and E. Klann, Role of reactive oxygen species in hippocampal longterm potentiation: contributory or inhibitory?, J Neurosci Res 70:1-7 (2002). 101. N. Z. Gerges, A. M. Aleisa, and K. A. Alkadhi, Impaired long-term potentiation in obese Zucker rats: possible involvement of presynaptic mechanism, Neuroscience 120:535-9 (2003). 102. M. Ankarcrona, J. Dypbukt, E. Bonfoco, et al., Glutamate-induced neuronal death: a succession of necrosis or apoptosis depending on mitochondrial function, Neuron 15:961-973 (1995). 103. M. Leist, and P. Nicotera, Apoptosis, excitotoxicity, and neuropathology, Exper Cell Res 239:183-201 (1998). 104. P. Nicotera, and S. Lipton, Excitotoxins in neuronal apoptosis and necrosis, J Cerebral Blood Flow Metab 19:583-591 (1999).


299

105. C. Wallin, S. Weber, and M. Sandberg, Glutathione efflux induced by NMDA and kainite: implications in neurotoxicity?, J Neurochem 73:1566-1572 (1999). 106. L. Dugan, S. Sensi, L. Canzoniero, et al., Mitochondrial production of reactive oxygen species in cortical neurons following exposure to N-methyl-D-aspartate, J Neurosci 15:6377-6388 (1995). 107. M. Schramm, S. Eimerl, and E. Costa, Serum and depolarizing agents cause acute neurotoxicity in cultured cerebellar granule cells: role of the glutamate receptor responsive to N-methyl-D-aspartate, Proc Natl Acad Sci USA 87:1193-1197 (1990). 108. M. Nishimura, K. Sato, T. Okada, P. Schloss, S. Shimada, and M. Tohyama, MK-801 blocks monoamine transporters expressed in HEK cells, FEBS Lett 423:376-380 (1998). 109. J. Lee, M. Grabb, G. Zipfel, and D. Choi, Brain tissue responses to ischemia, J Clin Invest 106:723-731 (2000). 110. C. Pereira, M. Santos, and C. Oliveira, Metabolic inhibition increases glutamate susceptibility on a PC12 cell line, J Neurosci Res 51:360-370 (1998). 111. M. Ward, A. Rego, B. Frenguelli, and D. Nicholls, Mitochondrial membrane potential and glutamate excitotoxicity in cultured cerebellar granule cells, J Neurosci 20:7208-7219 (2000). 112. A. Frandsen, and A. Schousboe, Dantrolene prevents glutamate cytotoxicity and Ca2+ release from intracellular stores in cultured cerebral cortical neurons, J Neurochem 56:1075-1078 (1991). 113. A. Frandsen, and A. Schousboe, Mobilization of dantrolene-sensitive intracellular calcium pools is involved in the cytotoxicity induced by quisqualate and N-methyl-Daspartate but not by 2-amino-3-(3-hydroxy-5-methylisoxazol-4-yl)propionate and kainate in cultured cerebral cortical neurons, Proc Natl Acad Sci USA 89:2590-2594 (1992). 114. S. Lei, D. Zhang, A. Abele, and S. Lipton, Blockade of NMDA receptor-mediated mobilization of intracellular Ca2+ prevents neurotoxicity, Brain Res 598:196-202 (1992). 115. S. Budd, and D. Nicholls, Mitochondria, calcium regulation, and acute glutamate excitotoxicity in cultured cerebellar granule cells, J Neurochem 67:2282-2291 (1996). 116. J. Dubinsky, and S. Rothman, Intracellular calcium concentrations during “chemical hypoxia” and excitotoxic neuronal injury, J Neurosci 11:2545-2551 (1991). 117. H. Wei, W. Wei, D. Bredesen, and D. Perry, Bcl-2 protects against apoptosis in neuronal cell line caused by thapsigargin-induced depletion of intracellular calcium stores, J Neurochem 70:2305-2314 (1998). 118. M. Villalba, A. Martínez-Serrano, P. Gómez-Puertas, et al., The role of pyruvate in neuronal calcium homeostasis: Effects on intracellular calcium pools, J Biol Chem 269:2468-2476 (1994). 119. F. Ruiz, G. Alvarez, R. Pereira, et al., Protection by pyruvate and malate against glutamate-mediated neurotoxicity, Neuroreport 9:1277-1282 (1998). 120. E. Bonfoco, D. Krainc, M. Ankarcrona, P. Nicotera, and S. Lipton, Apoptosis and necrosis: two distinct events induced respectively by mild and intense insults with Nmethyl-D-aspartate or nitric oxide/superoxide in cortical cell cultures, Proc Natl Acad Sci USA 92:7162-7166 (1995). 121. J. Carretero, E. Obrador, J. Pellicer, A. Pascual, and J. Estrela, Mitochondrial glutathione depletion by glutamine in growing tumor cells, Free Radical Biol Med 29:913-923 (2000).

300

Chapter 14

122. R. White, and I. Reynolds, Mitochondrial depolarization in glutamate-stimulated neurons: an early signal specific to excitotoxin exposure, J Neurosci 16:5688-5697 (1996). 123. D. G. Nicholls, Mitochondrial function and dysfunction in the cell: its relevance to aging and aging-related disease, Int J Biochem Cell Biol 34:1372-81 (2002). 124. A. Boldyrev, D. Carpenter, M. Huentelman, C. Peters, and P. Johnson, Sources of reactive oxygen species production in excitotoxin- stimulated cerebellar granule cells, Biochem Biophys Res Comm 256:320-324 (1999). 125. R. Castilho, M. Ward, and D. Nicholls, Oxidative stress, mitochondrial function, and acute glutamate excitotoxicity in cultured cerebellar granule cells, J Neurochem 72:1394-1401 (1999). 126. C. Sen, and L. Packer, Antioxidant and redox regulation of gene transcription, FASEB J 10:709-720 (1996). 127. V. Lakshminarayanan, E. Drab-Weiss, and K. Roebuck, H2O2 and tumor necrosis factor-alpha induce differential binding of the redox-responsive transcription factors AP-1 and NF-kappaB to the interleukin-8 promoter in endothelial and epithelial cells, J Biol Chem 273:32670-32678 (1998). 128. E. Shaulian, and M. Karin, AP-1 as a regulator of cell life and death, Nat Cell Biol 4:E131-6 (2002). 129. W. Kaufmann, P. Worley, J. Pegg, M. Bremer, and P. Isakson, COX-2, a synaptically induced enzyme, is expressed by excitatory neurons at postsynaptic sites in rat cerebral cortex, Proc Natl Acad Sci USA 93:2317-2321 (1996). 130. J. Marks, V. Bindokas, and X. Zhang, Maturation of vulnerability to excitotoxicity: intracellular mechanisms in cultured postnatal hippocampal neurons, Brain Res Devel Brain Res 124:101-116 (2000). 131. M. Grilli, M. Pizzi, M. Memo, Spano, and P, Neuroprotection by aspirin and sodium salicylate through blockade of NF-kappaB activation, Science 274:1383-1385 (1996). 132. H. Ko, K. Park, H. Kim, et al., Ca2+-mediated activation of c-Jun N-terminal kinase and nuclear factor kappa B by NMDA in cortical cell cultures, J Neurochem 71:13901395 (1998). 133. M. Grilli, and M. Memo, Possible role of NF-kappaB and p53 in the glutamateinduced pro-apoptotic nuonal pathway, Cell Death Differen 6:22-27 (1999). 134. G. Cambonie, L. Laplanche, J. Kamenka, and G. Barbanel, N-methyl-D-aspartate but not glutamate induces the release of hydroxyl radicals in the neonatal rat: modulation by group I metabotropic glutamate receptors, J Neurosci Res 62:84-90 (2000). 135. N. Perkins, The Rel/NF-kappa B family: friend and foe, Trends Biochem Sci 25:434440 (2000). 136. S. Budd, R. Castilho, and D. Nicholls, Mitochondrial membrane potential and hydroethidine-monitored superoxide generation in cultured cerebellar granule cells, FEBS Lett 415:21-4 (1997). 137. E. Chalecka-Franaszek, and D. Chuang, Lithium activates the serine/threonine kinase Akt-1 and suppresses glutamate-induced inhibition of Akt-1 activity in neurons, Proc Natl Acad Sci USA 96:8745-8750 (1999). 138. M. Hossain, J. Russell, R. Gomes, and J. Laterra, Neuroprotection by scatter factor/hepatocyte growth factor and FGF-1 in cerebellar granule neurons is phosphatidylinositol 3-kinase/Akt-dependent and MAPK/CREB-independent, J Neurochem 81:365-378 (2002). 139. S. Nonaka, C. Hough, and D. Chuang, Chronic lithium treatment robustly protects neurons in the central nervous system against excitotoxicity by inhibiting N-methyl-


301

D-aspartate receptor-mediated calcium influx, Proc Natl Acad Sci USA 95:2642-2647 (1998). 140. R. Hashimoto, C. Hough, T. Nakazawa, T. Yamamoto, and D. Chuang, Lithium protection against glutamate excitotoxicity in rat cerebral cortical neurons: involvement of NMDA receptor inhibition possibly by decreasing NR2B tyrosine phosphorylation, J Neurochem 80:589-597 (2002). 141. P. Rosenberg, S. Amin, and M. Leitner, Glutamate uptake disguises neurotoxic potency of glutamate agonists in cerebral cortex in dissociated cell culture, J Neurosci 12:56-61 (1992). 142. A. Volterra, D. Trotti, C. Tromba, S. Floridi, Racagni, and G, Glutamate uptake inhibition by oxygen free radicals in rat cortical astrocytes, J Neurosci 14:29242932 (1994). 143. D. Trotti, D. Rossi, O. Gjesdal, et al., Peroxynitrite inhibits glutamate transporter subtypes, J Biol Chem 271:5976-5979 (1996). 144. Y. Chen, W. Ying, V. Simma, et al., Overexpression of Cu,Zn superoxide dismutase attenuates oxidative inhibition of astrocyte glutamate uptake, J Neurochem 75:939945 (2000). 145. M. Harris, Y. Wang, N. Pedigo, Jr, K. Hensley, D. Butterfield, and J. Carney, Amyloid beta peptide (25-35) inhibits Na+-dependent glutamate uptake in rat hippocampal astrocyte cultures, J Neurochem 67:277-286 (1996). 146. S. Fine, R. Angel, S. Perry, et al., Tumor necrosis factor alpha inhibits glutamate uptake by primary human astrocytes: Implications for pathogenesis of HIV-1 dementia, J Biol Chem 271:15303-15306 (1996). 147. M. Beal, E. Brouillet, B. Jenkins, R. Henshaw, B. Rosen, and B. Hyman, Agedependent striatal excitotoxic lesions produced by the endogenous mitochondrial inhibitor malonate, J Neurochem 61:1147-1150 (1993). 148. J. Greene, R. Porter, R. Eller, and J. Greenamyre, Inhibition of succinate dehydrogenase by malonic acid produces an “excitotoxic” lesion in rat striatum, J Neurochem 61:1151-1154 (1993). 149. W. Maragos, and F. Silverstein, The mitochondrial inhibitor malonate enhances NMDA toxicity in the neonatal rat striatum, Dev Brain Res 88:117-121 (1995). 150. Z. Binienda, and C. Kim, Increase in levels of total free fatty acids in rat brain regions following 3-nitropropionic acid administration, Neurosci Lett 230:199-201 (1997). 151. M. Patel, R. Peoples, G. Yim, and G. Isom, Enhancement of NMDA-mediated responses by cyanide, Neurochem Res 19:1319-1323 (1994). 152. C. Pereira, and C. Oliveira, Glutamate toxicity on a PC12 cell line involves glutathione (GSH) depletion and oxidative stress, Free Radical Biol Med 23:637-647 (1997). 153. T. Murphy, R. Schnaar, J. Coyle, and A. Sastre, Glutamate cytotoxicity in a neuronal cell line is blocked by membrane depolarization, Brain Res 460:155-160 (1988). 154. G. Davey, S. Peuchen, and J. Clark, Energy thresholds in brain mitochondria: Potential involvement in neurodegeneration, J Biol Chem 273:12753-12757 (1998). 155. I. Inoue, S. Katayama, K. Takahashi, et al., Troglitazone has a scavenging effect on reactive oxygen species, Biochem Biophys Res Commun 235:113-116 (1997). 156. S. Uryu, J. Harada, M. Hisamoto, and T. Oda, Troglitazone inhibits both postglutamate neurotoxicity and low-potassium-induced apoptosis in cerebellar granule neurons, Brain Res 924:229-236 (2002).

302

Chapter 14

157. Y. Owada, T. Yoshimoto, and H. Kondo, Increased expression of the mRNA for brain- and skin-type but not heart-type fatty acid binding proteins following kainic acid systemic administration in the hippocampal glia of adult rats, Brain Res Molec Brain Res 42:156-160 (1996A). 158. D. Rhoads, R. Ockner, N. Peterson, and E. Raghupathy, Modulation of membrane transport by free fatty acids: inhibition of synaptosomal sodium-dependent amino acid uptake, Biochem 22:1965-1970 (1983). 159. A. Volterra, D. Trotti, P. Cassutti, et al., High sensitivity of glutamate uptake to extracellular free arachidonic acid levels in rat cortical synaptosomes and astrocytes, J Neurochem 59:600-606 (1992). 160. C. Blázquez, C. Sánchez, A. Daza, I. Galve-Roperh, and M. Guzmán, The stimulation of ketogenesis by cannabinoids in cultured astrocytes defines carnitine palmitoyltransferase I as a new ceramide-activated enzyme, J Neurochem 72:17591768 (1999A). 161. M. Guzmán, and C. Sánchez, Effects of cannabinoids on energy metabolism, Life Sci 65:657-664 (1999). 162. B. Costa, and M. Colleoni, Changes in rat brain energetic metabolism after exposure to anandamide or delta9-tetrahydrocannabinol, Eur J Pharmacol 395:1-7 (2000). 163. L. Walter, A. Franklin, A. Witting, T. Moller, and N. Stella, Astrocytes in culture produce anandamide and other acylethanolamides, J Biol Chem 277:20869-76 (2002). 164. H. H. Hansen, I. Azcoitia, S. Pons, et al., Blockade of cannabinoid CB(1) receptor function protects against in vivo disseminating brain damage following NMDAinduced excitotoxicity, J Neurochem 82:154-8 (2002). 165. T. Gomez Del Pulgar, M. L. De Ceballos, M. Guzman, and G. Velasco, Cannabinoids protect astrocytes from ceramide-induced apoptosis through the phosphatidylinositol 3-kinase/protein kinase B pathway, J Biol Chem 277:36527-33 (2002). 166. M. Egertova, B. F. Cravatt, and M. R. Elphick, Comparative analysis of fatty acid amide hydrolase and CB(1) cannabinoid receptor expression in the mouse brain: evidence of a widespread role for fatty acid amide hydrolase in regulation of endocannabinoid signaling, Neuroscience 119:481-96 (2003). 167. K. Kozak, and L. Marnett, Oxidative metabolism of endocannabinoids, Prostaglandins Leukotrienes and Essential Fatty Acids 66:211-220 (2002). 168. C. Fowler, Plant-derived, synthetic and endogenous cannabinoids as neuroprotective agents: non-psychoactive cannabinoids, “entourage” compounds and inhibitors of Nacyl ethanolamine breadkown as therapeutic strategies to avoid psychotropic effects, Brain Research Reviews 41:26-43 (2003).

Chapter 15 Pathogenesis of Alzheimer Disease: Metabolic Factors

15.1. INTRODUCTION Alzheimer disease is an increasingly common and incompletely understood cause of dementia, especially prevalent in the aging population. In an ongoing, widespread, intensive, and diverse research effort to better understand this disorder, the possible involvement of numerous genetic, metabolic, and environmental factors is being investigated, and impressive progress continues. Nevertheless, crucial questions remain unanswered, and conclusively established mechanisms of disease pathogenesis, prevention, and treatment are still lacking. In light of these unresolved issues, the present treatise, and its emphasis on interactions between intermediary metabolism and cell signaling, properly includes a consideration of Alzheimer disease. This conviction is justified by substantial evidence supportive of the hypothesis developed here: 1) that Alzheimer disease from the outset reflects a profound distortion of neuronal substrate utilization, metabolism, and mitochondrial function; 2) that abnormalities in fatty acid and ketone body metabolism are fundamental to the process; and 3) that these elements are critical determinants of the dementia. The hypothesis further proposes that this cascade of abnormal neuronal metabolism and energetics provides a milieu in which interaction between genetic and acquired factors intensifies the process, thereby augmenting the generation of ROS and oxidative stress, and driving the evolution of cognitive impairment and neuronal death. The essential elements of the hypothesis, and the evidence upon which they are based, are developed in the following pages. 303

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A detailed discussion of Alzheimer disease would exceed the scope and intent of the present analysis. It is clear, however, that despite substantial contributions provided by numerous investigators toward its characterization and elucidation1-4, confusion and uncertainty continue to cloud this important disorder. The picture that emerges is that of an apparently common form of dementia the incidence of which increases with age, yet which is not readily distinguished on clinical grounds from other causes of more or less similar changes in cognitive function. Even the histopathological and biochemical hallmarks that have been associated with dementia of the Alzheimer type, individually or collectively, are not necessary, sufficient, or specific for the diagnosis of Alzheimer disease. Thus, the characteristic extracellular amyloidcontaining neuritic plaques and intracellular neurofibrillary tangles may be found in non-demented individuals5 or in association with other conditions6. Prediction of Alzheimer disease among individuals who already exhibit mild cognitive impairment may be facilitated by demonstration of elevated cerebrospinal fluid levels of and decreased levels of amyloid7,8. However, these hallmarks do not necessarily correlate with severity or progression of the dementing process in those individuals with what appears clinically to be definite or probable Alzheimer disease. Finally, despite demonstrable early changes in mitochondria and mitochondrial DNA9, “typical” histopathological findings may be absent altogether in some individuals with Alzheimer type dementia, in whom findings instead may be consistent with normal, or may include features such as Lewy bodies that are more characteristic of other conditions. In the present analysis, focus will be directed at genetic and acquired elements that interact with, and potentially perturb, neuronal substrate utilization, mitochondrial energetics, oxidative stress, and antioxidant defense. Evidence will be presented that documents a strong yet unexpected linkage between neuronal regulation of these phenomena and similar signaling and regulatory mechanisms in other cell types, largely wg/Wnt-like PI3K/Akt/PKB-mediated. These processes in non-neuronal cells share with neuronal activation, and synaptic function and plasticity, a requirement for unpredictable yet immediate and rapid, largely carbohydrate-fueled increases in mitochondrial energetics and ATP generation, and suppression of potentially ROS-generating fatty acid oxidation. Disruption of this metabolic profile can be linked to Alzheimer disease through a “vicious cycle” of pathogenesis that encompasses and integrates genetic, metabolic, environmental, and nutritional determinants.

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15.2. GENETIC DETERMINANTS 15.2.1. Apolipoprotein E Isoforms As noted above in connection with most of the recognized accompaniments of Alzheimer disease, a lack of specificity also characterizes the genetic determinant apolipoprotein E (ApoE), the E4 isoform of which is associated with Alzheimer disease1-3,10,11. ApoE, in addition to its recognized relationship to plasma lipoprotein metabolism, exerts important effects on intracellular lipid metabolism12, is synthesized and secreted by astrocytes13, and is abundant in cytoplasm of normal and Alzheimer disease astrocytes and to a lesser extent neurons14. E4 differs from E3 and E2 isoforms in that an arginine is substituted for cysteine at position 112, resulting in its inability to form disulfide complexes and modifying its interaction with lipoprotein particles so as to favor association with triacylglycerol-rich VLDL and intermediate density lipoproteins. The E3 isoform enhances neurite outgrowth whereas E4 is inhibitory or neutral1,15, suggesting that E4 may participate in neuronal repair and remodeling after injury less effectively than E31,10. Absence of the cysteine sulfhydryl group may contribute to a relatively impaired antioxidant neuroprotective effect of E416, possibly reflecting in part a lipidation-dependent lower affinity of amyloid for E4 than for E317, an interaction which is also subject to modulation by allelic variation18. Recently an apoE-mediated provision of astrocyte-derived cholesterol for synaptogenesis has been identified19; this process is altered in Alzheimer brain by increased astrocyte expression of a cholesterol degrading enzyme20, and conceivably could be further distorted by E4. E4 also disrupts acetylcholine-mediated muscarinic signaling21. E4 induction of neuronal apoptosis involves its interaction with low-density lipoprotein receptor-related protein (LRP)22; LRP in turn appears to play a role in amyloid clearance23,24, which may be influenced by E4-LRP interaction25. E4 contributes to an earlier onset of Alzheimer disease2,26, and appears to accelerate its course in both symptomatic27 and cognitively normal subjects28,29. These effects are age-dependent30, and are greater in E4 homozygotes than in heterozygotes. In asymptomatic individuals the E4 isoform may impair aspects of visual attention31; it also is associated with relative impairment of 2-oxo-glutarate dehydrogenase, a key component of the citric acid cycle32 (see this Chapter 15.4.: Cerebral Metabolism and Insulin Signaling). E4 homozygotes exhibit increased formation of neuritic plaques, in which amyloid abundance is also increased, but this phenomenon appears to be independent of the course of the disease2,33. In Alzheimer disease, E4 is associated with, and

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may in part reflect, diminished expression and abundance of hippocampal insulin-degrading enzyme 34 . E4 predisposes to cognitive decline that occurs in subjects with HIV infection35 (also, see Chapter 13.4.2.: HIV Dementia), after cardiopulmonary bypass36, and among professional boxers37, and is associated with an unfavorable outcome after acute head injury 38 . These associations may reflect E4 isoform-specific increases in amyloid deposition and adverse intracellular effects39,40. Although most patients with confirmed Alzheimer disease carry at least one E4 allele, it is not sufficiently sensitive or specific to serve as the Together with PET-demonstrated cerebral sole diagnostic test41. metabolic impairment, however, it may permit preclinical detection42-44. Moreover, its prognostic significance is greatly influenced by gender, ethnicity, and apolipoprotein (a)45, and many or most individuals who carry it do not develop the disease46,47.

15.2.2. Presenilin and Amyloid Precursor Protein Mutations Germline mutations in presenilin-1 (PS1) and PS2 genes are present in most of the relatively small number of cases of early-onset familial Alzheimer disease4,11,48-51. However, the function of wild type presenilins is not limited to neuronal cells. Thus, while wild type PS1 modulates neurofilament-associated neurite outgrowth 52 , it is also regularly demonstrable in the inflammatory lesions of both genetically determined and sporadic inclusion-body myositis, along with ApoE and other Alzheimer-associated moieties including secretases, amyloid, and the paired helical filaments of phosphorylated that comprise neurofibrillary tangles53-55. Recent findings have linked PS1 to the formation of amyloid through cleavage of APP11, in an example of the more general process of regulated intramembrane proteolysis56. PS1 also participates in the regulation of catenin57,58(also, see this Chapter 15.4.2.1.: Wingless/Wnt-like Signaling), and thus the control of cell metabolism, cell adherens junctions, gene transcription, and cell proliferation58,59. Amyloid abundance has been linked to a locus on chromosome 10q60-62. Significantly, age-related amyloid deposition in a murine transgenic model overexpressing mutant PS1 and amyloid precursor protein (APP) Swedish mutant is associated with only limited neuronal loss63, suggesting that increased amyloid deposition may not be sufficient basis, per se, for the widespread and severe neuronal injury that characterizes Alzheimer disease. Thus, neither the recognized genetic markers nor the histopathological features associated with Alzheimer disease appear to be specific or sufficient causes of the condition by themselves, despite their


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prognostic significance and apparent relationship to pathogenesis, thereby suggesting a critical role for one or more other factors. These inconsistencies and lack of specificity futher suggest that the dementing process designated as Alzheimer disease may represent the end result of any of several underlying primary conditions and pathogenetic sequences, and/or that it is the consequence of a single, final common pathogenetic pathway that is activated by one or more primary causes. Consistent with this view and as will be developed, available evidence strongly suggests that altered neuronal energy metabolism and chronic oxidative stress play important roles in pathogenesis by contributing to the development of a “vicious cycle” of cell injury which influences, and is influenced by, effects of the recognized major genetic and non-genetic predisposing factors. Unfortunately, and despite the apparently critical pathogenetic role of oxidative stress in aging64, Alzheimer disease, and other chronic neurodegenerative disorders (e.g., Huntington and Parkinson diseases, and amyotrophic lateral sclerosis), the origins of this oxidative stress have not been identified65,66. In all of these conditions, moreover, clinical manifestations appear only after many years, implying that they represent the cumulative effects of underlying, sustained, and at least initially subtle abnormalities. In Alzheimer disease in particular, an understanding of its origins is impeded by its predominant occurrence in the aging population, in which cerebral metabolic rate for glucose decreases despite relatively stable cerebral oxygen consumption. Although differing in their regional distribution67, these characteristics of age-related change in cerebral metabolism are strikingly similar to those which may result from ketone body utilization, and which also occur in Alzheimer disease. Clues to their origins, and to those of oxidative stress and altered glucose utilization, may be provided by accumulating evidence that type II (noninsulin-dependent) diabetes mellitus contributes to dementia in the aging population.

15.3. ASSOCIATION OF ALZHEIMER DISEASE WITH TYPE II DIABETES MELLITUS Type II diabetes mellitus is associated with an increased incidence of impaired cognitive function68, disturbances in cerebral metabolism, and dementia69,70, but evidence concerning a possible relationship between diabetes mellitus and Alzheimer disease has been conflicting. Thus, several studies have suggested that there is no association between them, that there is no increase in Alzheimer-type pathology in diabetes

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mellitus71, or even that diabetes mellitus is protective. However, several methodological problems complicating the interpretation of studies in this area are recognized71,72. For example, some studies are largely clinicbased, in which the number of individuals enrolled may be small and subject to selection bias. Furthermore, there is potential uncertainty in the documentation of diabetes mellitus, or in the classification of dementia, especially in those individuals in whom the diagnosis of Alzheimer disease may be systematically excluded in the presence of diabetes mellitus because symptoms are attributed to vascular dementia73. However, similar processes related to diabetes mellitus and/or aging may occur elsewhere in the central nervous system. Thus, retinal neurodegeneration unrelated to vascular disease may occur as a relatively early consequence of human and experimental diabetes mellitus74, the evolution of age-related macular degeneration is associated with amyloid deposition75, and plasma FFA are associated with injury to microvascular endothelial cells and retinal pericytes76. Moreover, the recently demonstrated association of silent brain infarcts with cognitive decline77 could reflect a common underlying pathogenesis rather than, or in addition to, a causal connection. It is therefore highly significant that four large population-based or multicenter studies, conducted in the Netherlands, in Zutphen 78 and Rotterdam79, in Kuopio, Finland80, and at several centers in the United States81-83, demonstrate that type II diabetes mellitus represents a significant independent risk factor for accelerated cognitive decline and development of Alzheimer disease. In the Kuopio study80, the factors examined included elevated fasting insulin concentrations (indicating insulin resistance); in this group, an increased risk for Alzheimer disease was demonstrable only in those individuals who did not carry apolipoprotein E4. In the U.S. multicenter study82,83, women with type II diabetes mellitus had lower cognitive function scores on entry and accelerated cognitive decline over a several year follow up period. In the Zutphen study78, individuals with diabetes mellitus were significantly predisposed to dementia (not specifically designated as Alzheimer type), as were those non-diabetic subjects with high plasma insulin values and impaired glucose tolerance (i.e., insulin-resistant)78, and those in whom apolipoprotein E4 was expressed84. In addition to these population-based studies, three clinic-based studies also have demonstrated a significant association of Alzheimer disease with indicators of insulin resistance, i.e., increased fasting plasma insulin and glucose and/or elevated body mass index85, and with elevated Conflicting fasting plasma insulin86, and overt diabetes mellitus87. findings concerning cerebrospinal fluid insulin levels86,88 may in part


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reflect differences in experimental design. Of note, the Rotterdam population-based study also identified both high total and saturated fat intake89 and smoking90 as risk factors for Alzheimer disease, while lightto-moderate ethanol intake was protective against dementia91-93. Hypercholesterolemia has also been implicated as a risk factor on both epidemiological and experimental grounds94. What might account for these apparent linkages between Alzheimer disease and the systemic metabolic milieu?

15.4. CEREBRAL METABOLISM AND INSULIN SIGNALING To understand the possible basis for the association between Alzheimer disease and diabetes mellitus, it is necessary to re-examine the interactions among cell metabolism, cell signaling, and substrate-related differences in neuronal energy generation discussed in the earlier chapters of Part III. Important in this analysis is evidence that regulation of cerebral glucose metabolism by insulin is both direct and indirect95. Thus, increased cerebral utilization of ketone bodies has been demonstrated in untreated non-dementia subjects with insulin-dependent (type I) diabetes mellitus, associated with a relative shift of glucose metabolism to nonoxidative pathways as reflected in lactate release96. This effect of insulin lack implies that aerobic glycolysis in brain depends at least partially on insulin signaling. In companion studies overall glucose utilization was decreased or unchanged depending on the methodology employed96,97. Subsequent demonstration of unimpaired cerebral glucose utilization in type I diabetes mellitus during a hyperinsulinemic, slightly hypoglycemic clamp study98 would also be consistent with beneficial secondary effects on glucose utilization of insulin-induced suppression of plasma FFA and ketogenesis under experimental conditions. As noted above99, insulin does not appear to regulate brain glucose uptake (see Chapter12.2.: Glucose). However, widely distributed expression of the insulin receptor in brain100,101, and its regulation in response to training102, are indicative of an important insulin-signaling effect, presumably on other aspects of glucose metabolism96, as well as on carotid artery flow103. Moreover, streptozotocin-induced diabetes mellitus in rats is associated with oxidative stress and GLUT 3 adduct formation in hippocampal neurons104, with learning impairment70,105, and with diminished hippocampal blood flow, the latter ameliorated by the angiotensin converting enzyme inhibitor enalapril105 In addition, in view

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of the fact that glucose uptake is flow-dependent whereas that of FFA and ketone bodies is flow-independent (see Chapter 12: Utilization of Oxidizable Substrates in Brain), it is likely that conditions of reduced blood flow would result in a decrease in glucose uptake and utilization relative to, and more profound than, that of FFA and ketone bodies, thus further contributing to impaired learning. Conceivably, such diabetes mellitus-related changes might reflect direct cerebral effects of hyperglycemia106, or insulin lack. It seems more likely, however, that other consequences of insulin deficiency would contribute more importantly, especially the predictable increases in plasma concentrations of FFA and ketone bodies. As discussed, these latter factors are also subject to regulation by insulin, and may be important modulators of brain function, acutely and chronically. At least in part this modulation would reflect increased neuronal exposure to, and metabolism of, ketone bodies that are produced in excess in astrocyte and/or hepatocyte.

15.4.1. Glycolysis and the Citric Acid Cycle in Alzheimer Disease Important abnormalities in cerebral glucose metabolism that are similar to or consistent with diabetes mellitus-like changes have been demonstrated in Alzheimer patients. These include the decreased glucose utilization107 and oxidative phosphorylation in affected regions of brain that are recognized manifestations of early Alzheimer disease. Although neuroanatomical disconnection from other centers could contribute in part to these changes108, a basis for such disruption would be required in any case. Alzheimer disease-related metabolic changes precede the appearance of neuropsychological abnormalities and are potentially reversible109-113. Importantly, although atrophy of involved brain regions could contribute to metabolic changes114, assay of cerebral metabolic intermediates115 and correction for partial volume effect116 suggest that the observed abnormalities are not attributable to cerebral atrophy alone. Thus, while the functional changes are most pronounced in those areas in which pathological change is greatest71, they may involve relatively spared areas and neurons117. Moreover, activities of several enzymes involved in oxidative metabolism of glucose are decreased in Alzheimer brain tissue, including phosphofructokinase, pyruvate dehydrogenase (PDH), and neuronal 2-oxo-glutarate dehydrogenase (2–OGDH). The latter, a key citric acid cycle enzyme is thus critical in energy generation32,83,118-122 but is In addition, in Alzheimer susceptible to inhibition by ROS123. hippocampus and entorhinal cortex there is loss of mRNA for a subunit of cytochrome oxidase (complex IV of the electron transport chain); a


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similar change can occur reversibly in other settings in response to decreased neuronal activation, and thus it is consistent with the demonstrated decrease in neuronal activation in Alzheimer disease117. It has been suggested that increased insulin receptor abundance, reported in postmortem Alzheimer brain, may represent a compensatory response to impaired insulin signal transduction124. As discussed earlier (see Chapter 12: Utilization of Oxidizable Substrates in Brain), utilization of glucose via aerobic pathways, i.e., glycolysis and citric acid cycle-fueled oxidative phosphorylation, is maximally efficient for energy generation and minimizes generation of intramitochondrial ROS from the electron transport chain. In contrast, excessive oxidation of ketone bodies in neuronal mitochondria would contribute importantly to the pathogenesis of the major metabolic changes in Alzheimer disease, via several mechanisms. First, PDH activity would be suppressed (see Chapter 12.3.: Ketone Bodies, and Fig. 15.1 and Fig. 15.2), thereby inhibiting glycolysis and the tightly

Figure 15.1. Fatty acid and ketone body oxidation in mitochondria suppresses glycolysis. Mitochondrial of fatty acids (astrocytes) and ketone bodies (neurons) generates three moieties which activate kinase-mediated feedback inhibition of PDH, i.e., acetyl CoA, ATP, and NADH. This effect, together with an accompanying suppression of glycolysis, decreases mitochondrial oxidation of pyruvate, and overall utilization of glucose. In peripheral tissues, especially muscle, this effect of mitochondrial fatty acid oxidation contributes to the “Randle cycle” inhibitory effect on glucose utilization and insulin resistance. Dashed lines: inhibition of PDH by acetyl CoA/CoA, ATP/ADP, and NADH/NAD. Abbreviations — ATP: adenosine triphosphate; CPT-I: carnitine palmitoyltransferase-I; NADPH: nucleotide adenine triphosphate, reduced form; PDH: pyruvate dehydrogenase.

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controlled fueling of the citric acid cycle by pyruvate-derived acetyl CoA. Second, as demonstrated in studies of perfused heart, provision of ketone bodies as sole nutrient may inhibit the citric acid cycle at the 2OGDH reaction123,125,126; 2-OGDH is also inhibited by oxidative stress in nerve terminals127. Importantly, mitochondrial 2-oxo-acid dehydrogenase complexes may be inhibited by ROS generated in the redox reactions they themselves catalyze128. The somewhat inconsistent depression of cerebral 2-OGDH activity in Alzheimer brain as determined in vitro129, and its lack of correlation with enzyme protein130, would also be compatible with such a mechanism. Thus, ketone body inhibition of 2OGDH in the perfused heart (and presumably brain) is reversible125, as a result of which enzyme activity assayed in vitro would be influenced by experimental conditions. Similar considerations may explain the failure


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of in vitro assays of biopsied Alzheimer brain tissue to demonstrate consistent abnormalities in glucose metabolism and mitochondrial Finally, excessive ketone body-generated reducing function131. equivalents could increase electron flux and density in the electron transport chain, thus predisposing to generation of superoxide anion and derivative ROS, and oxidative stress. In addition to the more direct effects of ROS and oxidative stress, increased generation of superoxide would favor formation of peroxynitrite, creating a milieu in which NO has been shown to be injurious to the cell. Under other circumstances, in contrast, NO may be either non-injurious or even beneficial132-135, possibly reflecting in part a neuroprotective effect of peroxynitrite that is mediated by pentose phosphate pathway-generated NADPH136. The foregoing considerations suggest that the pathogenesis of depressed glucose utilization, impaired energy generation, and sustained oxidative stress that characterize Alzheimer disease can be understood, perhaps primarily, as the consequence of a shift in neuronal substrate utilization; this shift is characterized by an increase in mitochondrial oxidation of ketone bodies relative to glucose, oxidation of which is thereby suppressed. Furthermore, as shown in primary hepatocytes, transforming growth increases in mitochondrial fatty acid oxidation and ketogenesis137, would also contribute to inhibition of glycolysis138, and support the increase in gluconeogenesis implied by expression of phosphoenolpyruvate carboxykinase in these cells139. also induces 140-142 oxidative stress . Similar effects in the astrocyte could in part account for the Alzheimer-like neuronal pathology associated with chronic 143 overproduction of . A possible basis for the neuroprotective effect of under certain conditions144 is discussed below (see this Chapter 15.4.2.2.: Neuroprotection and Cell Cycle-Linked Neuronal Death). Recent evidence suggests that the energetically divergent effects of specific wg/Wnt-like signaling agonists and (see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable, and Chapter 6.3.: Transforming Growth may interact in regulating synaptogenesis145. By this line of reasoning, therefore, the decreased cerebral utilization of glucose that occurs in Alzheimer disease, as in diabetes mellitus, may represent the equivalent of a resistance to and attenuation of insulin-like metabolic effects, induced in this setting by the preferential neuronal utilization of alternative substrates, especially ketone bodies. The normal effects of insulin so compromised include promotion of oxidative metabolism of glucose, in which relative increases in the activity of both

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glycolysis and the citric acid cycle would otherwise augment oxidative phosphorylation and the generation of ATP. These considerations are consistent with abundant evidence that diminished glucose utilization in Alzheimer disease occurs in what at least initially are viable albeit functionally compromised areas of the brain109-113, implying that the surviving neurons are utilizing alternative fuel(s), among which locally or systemically generated ketone bodies would be especially important. This interpretation is consistent with evidence that in areas affected by Alzheimer disease, oxygen consumption is relatively stable despite decreased glucose utilization121, 146, 147. This relationship also characterizes the metabolic activity of normal brain in aging67 and during acute exogenous hyperketonemia148. Importantly, all three circumstances appear to represent the reverse of normal neuronal activation, in which oxygen consumption is relatively stable despite increased glucose utilization (see Chapter 14.2.: Activation-Related Glucose and Oxygen Consumption). In Alzheimer subjects, moreover, activation-induced increases in oxidative metabolism are associated with subsequent cognitive decline149. This relationship would also be consistent with, and at least in part explained by, neuronal utilization of alternative substrates that are energetically less efficient and potentially oxidative stressinducing; these would include ketone bodies, oxidation of which in generating a given amount of ATP obligates greater oxygen consumption than does glucose (see Chapter 12.3.: Ketone Bodies). These metabolic phenomena are further illuminated by their linkage to other aspects of insulin signal transduction and its relationship to intermediary metabolism, phosphorylation, and amyloid.

15.4.2. Wingless/Wnt-Like Signal Transduction and Effects of Lithium Recent findings have identified the perhaps surprising involvement in neuronal function and disease of the wingless/Wnt-like signal transduction pathway through phosphatidyl inositol-3-kinase (PI3K) and Akt/protein kinase B (Akt/PKB); in neural progenitor and non-neuronal cells this signaling cascade is linked to proliferation (see Chapter 3: Nutrient and Energy Metabolism in Cell Proliferation, and Chapter 13.4.3.: Cell Proliferation and Tumorigenesis). The key negative regulator of this pathway, glycogen synthase inhibits glycogen synthase, an important determinant of cellular storage and availability of glucose, and pyruvate dehydrogenase (PDH), the mitochondrial regulator of glucose (pyruvate) oxidation. Interaction of this pathway with neuronal substrate utilization and intermediary metabolism appears


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to exert a major influence on the generation of oxidative stress, amyloid, the phosphorylated form of the microtubule-associated protein and survival. 15.4.2.1. Wingless/Wnt-Like Signaling The demonstration that PS1 binds to both and 150-154 as well as 155,156 suggests that these interactions may regulate neuronal counterparts of the wingless (wg) cascade in Drosophila and the Wnt/APC (adenomatous polyposis coli)/E-cadherin system in mammals157-160(also, see this Chapter 15.2.2.: Presenilin and Amyloid Precursor Protein Mutations). This cascade is of critical importance in embryogenesis161 and in the pathogenesis of human familial adenomatous polyposis and, if untreated, the latter’s virtually invariable culmination in colon cancer (see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). In the absence of colonocyte wg/Wnt-like signaling, is sequestered in a complex that includes axin, the APC gene product, and This promotes phosphorylation of and its subsequent degradation, thereby suppressing transcriptional activation and cell proliferation while promoting the E-cadherin-mediated cell-cell adhesion that is associated with normal cell differentiation. Conversely, mutations in the APC or genes which prevent phosphorylation of increase the latter’s abundance in cytosol. In colonocytes and other non-neuronal cells, the unopposed wg/Wnt-like signaling that such changes represent enhances transcriptional regulation and promotion of cell proliferation and tumorigenesis. wg/Wnt-like signaling is activated directly or indirectly by several mitogenic factors. This concept is consistent with virtually ubiquitous expression of the mutually inhibitory insulin receptor substrate-1 (IRS-1) 162-164 and , and has been demonstrated for insulin165 and several other growth factors, e.g., hepatocyte growth factor166, and brain-derived neurotrophic factor167. Furthermore, survival signaling converges with that of neuronal depolarization at PI3K168, indicating even greater scope for this pathway in neuronal function and survival. Inhibition of by insulin during glycogen synthesis has long been recognized and is characteristic early in human and experimental hepatocarcinogenesis169,170. More broadly, however, these interactions appear to represent the pivotal convergence of four distinct and essential antecedents to mitosis, i.e., diminished cell-cell adhesion, enhanced storage and availability of energy-generating substrate, activation of transcriptional regulation, and

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promotion of cell cycle progression, as discussed above (see Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable). This unexpected neuronal expression of a pathway so intimately involved in cell proliferation may be understandable in terms of neuronal energy requirements. In neuronal activation as in mitosis, both rate and magnitude of ATP generation are critical, and are optimally supported by an insulin-like enhancement of glycolytic fueling of the citric acid cycle and oxidative phosphorylation. In cortical neurons, the role of PS1 superficially resembles the scaffolding function of APC or axin in the colonocyte171; that is, it participates in formation and regulation of an enzyme–substrate complex (i.e., In both mitosis and neuronal activation, a key element is that wg/Wnt-like (e.g., insulin) signaling is required to prevent formation of this complex. This critical action is accomplished through the initiation of PI3K/Akt/PKB(in neurons, mediated, S9 phosphorylation and inactivation of may also be designated protein kinase I). Conversely, withdrawal of the trophic factor (e.g., insulin) abrogates PI3K/Akt/PKB signaling, resulting in activation of through Y216 phosphorylation, 172,173 and in neuronal degeneration . In contrast to non-postmitotic epithelial cells, in which the transcriptional regulatory arm of wg/Wnt-like signaling through is also governed by the latter’s interaction with a scaffold-promoted complex (APC/axin), PS-1 does not regulate this mechanism in neuronal cells154. This observation suggests that in the neuron, wg/Wnt-like signaling may be related primarily to the regulation of metabolism, including support of synaptic plasticity and long term potentiation (see Chapter 14.5.2.1.: Long Term Potentiation), and the phosphorylation of as discussed below. This somewhat “truncated” neuronal wg/Wnt-like regulatory pathway may also serve to avoid an effect that is a normal process in cell proliferation but would be counterproductive in the synaptic neuron, i.e., wg/Wnt-like signaling-induced disruption of synapse-associated adherens junctions174-176. 15.4.2.2. Neuroprotection and Cell Cycle-Linked Neuronal Death A key element in neuronal wg/Wnt-like signaling involves its relationship to PS1 and phosphorylation. Thus, neuronal insulin signaling via IRS-1 and the PI3K/Akt/PKB cascade inhibits and therefore its phosphorylation of 177,178, and in addtion protects against apoptosis179. It also favorably influences APP processing and trafficking so as to minimize neuronal exposure to amyloid180-182, and mediates the


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183 neuroprotective effects of and of 184 metabotrophic glutamate receptors . As a corollary, compromise of the neuroprotective wg/Wnt-like signal cascade, e.g., resulting from impaired insulin signaling, could contribute significantly to the generation of important manifestations of Alzheimer disease. These include impaired synaptic AMPAR kinetics and LTP (see Chapter 14.5.2.1.: Long Term Potentiation), abnormal phosphorylation of and formation of neurofibrillary tangles177,178, amyloid exposure180-182, and programmed cell death179. It should be noted that phosphorylation status is not determined solely by PI3K/Akt/PKB. Thus, neuroprotective inactivating phosphorylation of may also be effected by protein kinase 185. In addition, phosphorylation may reflect activity of kinases 186,187 other than , and is also regulated by protein phosphatase 188 activity . Accordingly, phosphorylation status reflects both kinase activity (e.g., and phosphatase activity (e.g., protein phosphatase 2A — PP2A). For example, in the increased phosphorylation associated with fasting (a hypoinsulinemic state), suppressed PP2A rather than increased kinase activity appears to be the more important determinant189. Moreover, PI3K/Akt/PKB-mediated inactivation of (i.e., wg/Wnt-like signaling) would prevent phosphorylation not only of 177 but presumably also of PDH190, the pivotal determinant of glycolytic fueling of the citric acid cycle and oxidative phosphorylation. Similar mechanisms appear to account for the Akt/PKB induced protective activation of telomerase191,192(also, see Chapter 5.3.2.: wg/Wnt-Like Signaling: Convergence of Mitochondria, p53, and Telomerase), and the 194,195 potentially neuroprotective effects of nerve growth factor193, , 196 and insulin-like growth factor-1 , as well as certain other small molecules197. Clinical evidence pertaining to a possible neuroprotective effect of estrogens21,198-202 remains conflicting202,203. However, estrogens also activate neuronal protein kinase C204 and the mitogenic extracellularsignal regulated (ERK) pathway of the mitogen-activated protein (MAP) kinase cascade, thereby linking neurotrophin-initiated signaling205, and regulation of synaptic plasticity206, with the metabolic and energetic effects of wg/Wnt-like signaling, i.e., PI3K/Akt/PKB195(also, see Chapter 14.5.2.1.: Long-Term Potentiation). In the absence of wg/Wnt-like signaling, conversely, PS1 forms a complex with active and hyperphosphorylation of generates neurofibrillary tangles 177,178. binding, kinase activity152, and amyloid secretion49 are increased by those PS1 mutations that are associated with early onset familial Alzheimer disease48,50; a similar mechanism undoubtedly

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contributes to the activation and neuronal death associated with 172,173 trophic factor withdrawal . Although wild-type but not mutant PS1 was reported to facilitate constitutive turnover207, the relationship of this effect to wg/Wnt-like signaling was not defined. Together, these observations in neural and non-neural cells suggest that wg/Wnt-like signaling may be viewed as a general mechanism for the regulation of cellular energetics. When active, e.g., with insulin or hepatocyte growth factor signaling-induced inactivation of and increased abundance, it may be harnessed in support of maximal rates of ATP generation, e.g., in mitosis, neuronal activation, or other time-critical and energy-critical processes. As discussed above (Chapter 3.3.2.: Wingless/Wnt-Like Signaling: Convergence of Antecedents and the Unpredictable), operation of the pathway may be disturbed in either direction. In a non-neural cell such as the colonocyte, mutationally inactivated APC may permit inappropriately sustained activation of wg/Wnt-like signaling, leading to disruption of cell-cell adhesion, increased transactivational activity, mitosis, and tumorigenesis. In the neuron, conversely, inappropriate suppression of wg/Wnt-like signaling (e.g., insulin resistance and/or mutant PS1 expression) may permit inappropriately sustained activation of Decreased PI3K/Akt/PKB and increased activity would lead to inactivation of PDH and impaired intermediary metabolism, diminished post-synaptic neuronal AMPAR insertion/cycling and LTP, hyperphosphorylated and generation of neurofibrillary tangles. Unlike axin, which negatively regulates phosphorylation, wild type PS1 positively influences this reaction208, while PS1 mutations associated with familial Alzheimer disease accentuate the wild type effect152. These observations are consistent with evidence that, in addition to compromising wg/Wnt-like signaling, PS1 mutations associated with familial Alzheimer disease predispose to neuronal apoptosis. Thus, they: 1) augment phosphorylation152, accelerate 209,210 turnover and/or alter its intracellular trafficking211 ; 2) augment amyloid secretion48-50; 3) interfere with the antiapoptotic (pro-survival) 212,213 effect of ; and 4) predispose to oxidative stress, mitochondrial dysfunction, and excitotoxic death of hippocampal neurons214. These concepts identify a critical difference between wg/Wnt-like signaling in the neuron and the differentiating colonocyte. Thus, wg/Wnt-like (e.g., insulin) signaling normally is required during neuronal activation but is suppressed in the differentiating post-mitotic colonocyte. In the latter, E-cadherin-mediated cell-cell adhesion is stabilized, butyrate and other antiproliferative short chain fatty acids derived from colonic flora are dominant fuels, while glucose utilization and cell proliferation


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are suppressed (also, see Chapter 6.2.: Butyrate). Significantly, the antiproliferative short chain fatty acid butyrate induces phosphorylation in human neuroblastoma cells215, presumably reflecting suppressed wg/Wnt-like signaling and enhanced activity. Together, the findings imply a functional duality of wg/Wnt-like signaling and cell energetics, i.e., in their support of both cell cycle progression and neuronal activation. This concept is further suggested by recent evidence that, unlike glutamate, the inhibitory neurotransmitter GABA not only is poorly utilized in the citric acid cycle216 but also suppresses hepatocyte proliferation and liver regeneration after partial hepatectomy217’218. Moreover, it may be appropriate to regard wild-type PS1 as being a growth-promoting factor, superficially equivalent to a mediator of wg/Wnt-like signaling. Thus, suppression of PS1 in leukemia cell lines — by p53, or antisense PS1 cDNA — is associated with growth arrest and/or apoptosis219. Conversely, wild-type PS1 expression is increased during neuronal differentiation and in neuroblastoma cell lines; in astrocytes, its minimal basal expression is increased by proliferative stimuli, and it is actively expressed in glioma cell lines220. The perhaps unexpected induction of apoptosis by overexpression of the highly homologous PS2 under conditions of growth factor withdrawal221 may, perhaps analogous to apoptosis caused by amyloid, directly222,223 or indirectly224 reflect forced cell cycle progression in the absence of an appropriately supportive intracellular metabolic milieu225(also, see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer). The concept that neuronal cell cycle progression induced by amyloid is an important and closely associated accompaniment and precipitating factor in neuronal death in Alzheimer disease is increasingly well supported222,226-229. A direct linkage between the cell cycle and neuronal injury is suggested by cell division cycle (cdc) 2 kinasemediated phosphorylation of 230. Recent evidence also indicates a contributory role of ganglioside GD3 in this interaction226,227. While the mechanism underlying these associations awaits elucidation, they may reflect induction of cell cycle progression by moderate levels of ROS231(also, see Chapter 3.3.2.3.: Reactive Oxygen Species), under conditions in which this effect is potentially lethal because cellular energetic status is inadequate to support the demands of cell cycle progression (see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer). In this additional respect, i.e., an apparently critical relationship of the cell cycle to activation of programmed cell death, there exist important and somewhat unexpected parallels between neurons232-235 and non-

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neuronal cells225. In both settings, forced cell cycle progression in the absence of requisite cellular metabolic and energetic conditions may precipitate apoptosis; inhibition of cell cycle progression under these conditions may be protective225(also, see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer). It is plausible that such a mechanism accounts for the neuroprotection afforded by 144 , a cytokine that is antiproliferative in non-neural cells, that generates amyloid-like pathology when chronically overexpressed143, and that increases APP expression in astrocytes236. A similar effect 237 may contribute to complement-mediated neuroprotection . Moreover, an antiproliferative effect of certain nonsteroidal antiinflammatory drugs (NSAIDs), demonstrated in non-neural cells, may account for their neuroprotective effect, although evidence is conflicting238-243. NSAIDs may also directly suppress amyloid levels independently of their inhibition of COX2244-246, as well as through non-stereospecific inhibition 247,248 of . The possible efficacy of these agents in Alzheimer disease prevention is currently undergoing multicenter, placebocontrolled evaluation249. The complex interactions which link energy metabolism and cell proliferation to cell death are discussed in greater detail elsewhere225,250 (also, see Chapter 5.3.4.: Apoptosis and the Cell Cycle: Chronic Viral Hepatitis and Cancer). 15.4.2.3. Neurochemical Effects of Lithium The broader importance of wg/Wnt-like signaling is further suggested by the diverse effects of lithium ion, the smallest univalent cation except for the proton. Through both direct251,252 and indirect253 inhibition of and stimulation of PKB/Akt, lithium: 1) mimics aspects of insulin/PI3K-stimulated glucose metabolism254; 2) increases cytosolic 252 abundance of ; 3) induces Tcf transcriptional activity in 255 fibroblasts ; 4) augments cell proliferation256,257; 5) increases cerebral grey matter volume258; 6) protects against glutamate-induced excitotoxicity253,239; 7) mimics wg/Wnt-like signaling participation in axonal remodelling260; 8) inhibits phosphorylation of 178,252,261 ; 9) inhibits the isoform, thus suppressing amyloid 182 production in vitro and in vivo ; and 10) protects against the p53independent262, Bcl-2/Bax-modulated263, cell cycle-associated222 neuronal death induced by amyloid264-266. As a corollary, the therapeutic efficacy of lithium in bipolar (manic-depressive) disorder implies that the changes in energy metabolism documented in this and related conditions267-269 may be important in their pathogenesis. Thus, affective depression may


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represent the clinical manifestation of energetic depression in selected neuronal populations270. Recent studies suggest that abnormalities in substrate utilization and cell energetics similar to those considered above may also contribute to the evolution of other neurodegenerative processes the pathogenesis of which is incompletely understood, such as Parkinson disease66,271-273. For 274 example, abundance of the prion protein has been found to be increased in Alzheimer brain275, and has been linked to neuronal caveolin1 and the src family tyrosine kinase Fyn276. It has become clear that caveoli — in which are localized several tyrosine kinase receptors, src family tyrosine kinases, and related docking proteins — represent an important focus of receptor-initiated tyrosine kinase signaling activity in both neuronal and non-neuronal cells277,278. Fyn itself has been shown to bind to insulin receptor substrate-1 (IRS-1) during insulin stimulation in vitro and in CHO cells279, and to mediate diverse functions including mitogenic signaling280, long term potentiation281, and insulin-stimulated phosphorylation of caveolin-1 in 3T3-L1 cells282. Fyn also participates in the formation of a ternary complex, comprised of neural cell adhesion molecule (NCAM), Fyn, and focal adhesion kinase (FAK), that is A specific required for NCAM-mediated growth cone formation283. connection of these particular interactions to neural cell glucose utilization and cellular/mitochondrial energetics is strongly supported by recent evidence that the synthetic peptide corresponding to residues 106126 of human PrP induces apoptosis via a primary effect on mitochondria284; this action is associated with activation of GSK3 that is 285 suppressible by insulin and , i.e., by wg/Wnt-like-signaling. This relationship among PrP, mitochondria, and signaling by receptor and non-receptor tyrosine kinases, e.g., the insulin receptor, strongly suggests its pathogenetic importance, both for Alzheimer disease and for those conditions more explicitly linked to the prion protein family274.

15.5. METABOLIC DETERMINANTS IN ALZHEIMER DISEASE: SYNTHESIS 15.5.1. Oxidative Stress and Amyloid: A “Vicious Cycle” of Multiple Interacting Determinants As noted above, increasing evidence indicates that Alzheimer disease is associated with early and persisting oxidative stress286, altered neuronal energy metabolism, and the generation and deposition of amyloid.

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While these phenomena are not specific for Alzheimer disease, they are among its important characteristics. Of overriding importance, moreover, is evidence not only that each appears to contribute to the evolving structural and functional deterioration that characterizes Alzheimer disease progression, but also that each is able to amplify the deleterious effects of the others. Together, they comprise a “vicious cycle” of neurodegeneration. In so doing, they also provide a mechanistic framework into which interactions among genetic, metabolic, and environmental factors can be integrated in a unifying formulation of the dementing process. This formulation is described in the following paragaphs. 15.5.1.1.

Amyloid

Reflecting both direct and indirect effects, amyloid induces oxidative stress287. Thus, amyloid inhibits cellular uptake of glucose288-290, in addition to perturbing electron transport chain function and mitochondrial respiration291-296, and synaptic function 297 . Indeed, the electron transport chain has been shown to be required for manifestations of amyloid-induced cell death298. This important observation is consistent with evidence that amyloid (25-35)-induced cell injury requires its intracellular, not extracellular, location299,300, that amyloid oligomers may be the principal mediator of early injury 11 , that amyloid’s interaction with ApoE4 may promote lysosomal leakage40, and that amyloid oligomers may exist in a cytosolic location301,302 where they interact with heat shock (stress) proteins, and may lead to the latter’s Extramitochondrial origin of ROS, transcriptional upregulation303. especially early in the process, may also contribute to amyloid’s 304 injurious effects . Furthermore, intracellular accumulation of amyloid may be related to, and mediated by, expression of the nicotinic acetylcholine receptor305, and precedes development of both neurofibrillary tangles and extracellular amyloid plaques297,306. Failure of to prevent amyloid-induced oxidative stress266 is consistent with a direct interaction between amyloid and mitochondria, thereby circumventing the protective effects of wg/Wnt-like signaling as discussed above. In addition, however, amyloid peptide interaction with partially purified insulin receptor in vitro suggests that it may also interfere with insulin receptor (thus, wg/Wnt-like) signaling at a site external to the plasma membrane, by inhibiting insulin binding307. Amyloid (25-35) in a concentration-dependent manner may inhibit astrocyte removal of glutamate from the synaptic cleft308, but has also been shown to augment this process309. Deletion of the insulin degrading


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enzymes results in increased brain levels of amyloid310,311 and a type II diabetes mellitus-like systemic metabolic profile310. Given the association of type II diabetes mellitus with Alzheimer disease (see this Chapter 15.3.: Association of Alzheimer Disease with Type II Diabetes Mellitus), it is likely that diabetes mellitus also contributes to increased amyloid in brain. Increased E4 abundance in Alzheimer disease may also in part reflect diminished expression of hippocampal insulin-degrading enzyme34. Finally, as noted above, amyloid also activates in turn, active phosphorylates and thus inactivates PDH190, thereby inhibiting aerobic glycolysis and undoubtedly contributing to impaired neuronal uptake and utilization of glucose289. Conversely, activation of wg/Wnt-like PI3K/Akt/PKB signaling, and inhibit GSK3 and decrease amyloid exposure182. It is clear that wg/Wnt-like signaling, e.g., insulininduced, is integrally and inextricably linked to the phenomena that augment neuronal function and survival. 15.5.1.2. Oxidative Stress In addition to its direct adverse effects on the poorly antioxidantdefended neuron, oxidative stress alters post-translational processing of APP, thereby augmenting the production of amyloid312-317 and increases expression and activity318. Oxidative stress also impairs astrocyte uptake of glutamate319-321, potentially exposing the post-synaptic neuron to excitotoxic glutamate concentrations in the synaptic cleft. As discussed in the preceding paragraph, amyloid in turn induces electron transport chain disruption and additional oxidative stress. Thus, the interaction between amyloid and oxidative stress represents a mutually reinforcing destructive interplay, providing a foundation upon which genetic and environmental determinants of pathogenesis may interact. 15.5.1.3. Genetic Factors The Swedish APP 670/671 mutation increases cerebral amyloid-42 generation322; oxidative stress is especially prounced in this variant. The E4 isoform of apolipoprotein E predisposes to Alzheimer disease, apparently in large part through its effects on amyloid. Thus, E4 is also associated with diminished clearance of amyloid22-24, and E4’s lower binding affinity for amyloid relative to that of E317 could amplify the deleterious effects of amyloid, including their interaction in promoting lysosomal leakage, as noted40. In addition, as compared with E3, E4 Moreover, E4 exerts a weaker antioxidant neuroprotective effect16. fragments formed during intracellular processing may directly interact

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with cytoskeletal components, predisposing to formation of neurofibrillary tangle-like inclusions323. PS mutations associated with Alzheimer disease increase binding to PS and its kinase activity152. As a result, they augment hyperphosphorylation of generation of neurofibrillary tangles177,178, and amyloid secretion48-50. Mutant PS-associated augmentation of activity thus abrogates the protective effect of wg/Wnt-like signaling that reflects PI3K/Akt/PKB-mediated inactivation of (e.g., by insulin), that otherwise would prevent phosphorylation of l77 and PDH190, and promote cell survival. 15.5.1.4. Changes in Fatty Acid and Ketone Body Metabolism Sustained elevations of plasma [FFA], e.g., in diabetes mellitus, may further modulate the amyloid-oxidative stress cycle, overriding insulinactivated wg/Wnt-like signaling190,324-328 (also, see Chapter 7.2.: Adverse Effects of Fatty Acids on Mitochondrial Function), promoting astrocytic and thus neuronal oxidative stress, increasing amyloid generation, and increasing activity and phosphorylation. Adding to similar effects of amyloid and oxidative stress, FFA also inhibit astrocyte uptake of glutamate329, further contributing to the potential for excitotoxic-like neuronal injury. Increased neuronal utilization of ketone bodies (e.g., resulting from FFA-induced increases in astrocyte fatty acid oxidation-driven ketogenesis) would impair citric acid cycle activity126, as well as compromise energetics through their uncoupling-like effect and augmentation of mitochondrial matrix reducing potential and ROS generation (see Chapter 12.3.: Ketone Bodies, and Chapter 6.2.: Butyrate). Increased ketone body utilization also would suppress neuronal PDH activity and glycolysis, as demonstrated experimentally during increased cerebral utilization of ketone bodies in rats330 and human subjects148. Consistent with this general formulation, injurious products of fatty acid peroxidation indicative of oxidative stress, e.g., 4-hydroxynonenal (HNE), are increased in Alzheimer brain, ventricular fluid, and plasma331,332. Streptozotocin-induced diabetes mellitus in rats is associated with learning impairment70 and with formation of adducts between HNE and the neuronal glucose transporter GLUT 3104; HNE also mediates oxidative stress in vitro333. Finally, as discussed earlier (see this Chapter 15.4.1.: Glycolysis and the Citric Acid Cycle in Alzheimer Disease), increases hepatocyte mitochondrial fatty acid oxidation137 and oxidative stress140-142. Similar effects of signaling in astrocytes would serve to increase gluconeogenesis139, inhibit glycolysis138, and


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contribute to the generation of Alzheimer-like neuronal pathology143. This scenario is consistent with recent evidence that injection of into the cisterna magna induces EEC changes suggestive of altered cerebral energetics334, and increases systemic fatty acid oxidation335. 15.5.1.5. Neuroprotective Responses and the “Vicious Cycle” The importance of oxidative stress in Alzheimer disease is further suggessted by evidence that antioxidant neuroprotective responses are activated in this disorder. In neurons and especially in astrocytes, ROSinduced activation would help to protect against amyloidinduced oxidative stress-linked neuronal death336,337 through transcriptional regulation of redox-related protective mechanisms338-343. Other antioxidant defenses including superoxide dismutase and heat shock proteins are also activated. A similar response to oxidative stress may contribute to the reported lower incidence of dementia associated with limited ethanol ingestion91-93(also, see Chapter 7.4.2.2.: Paradoxical Benefits of Adversity). As noted earlier (see this Chapter15.4.2.2.: Neuroprotection and Cell Cycle-Linked Neuronal Death; and Chapter 14.5.2.1.: Long Term Potentiation), wg/Wnt-like signaling and PI3K/Akt/PKB-mediated inactivation of would prevent phosphorylation not only of 177; but presumably also of a amyloid-induced, inactivating phosphorylation of PDH190. In addition to participating in neuroprotective responses, wg/Wnt-like signaling plays a fundamental role in the regulated membranous expresssion of AMPAR in synaptic plasticity and long term potentiation344-347, implying that wg/Wnt-like signaling is directly involved in normal memory function (see Chapter 14.5.2.1.: Long Term Potentiation). Importantly, the suggested interactions among components of the oxidative amyloid cycle are also consistent with demonstrated or proposed cytoprotective actions of diverse agents such as in CHO cells182, and lipoic acid348, salicylate, and estradiol in astrocytes349(also, see Chapter 14.5.2.2.: Excitotoxicity — Origins of Oxidative Stress, and this Chapter 15.4.2.2.: Neuroprotection and Cell Cycle-Linked Neuronal Death). These agents may afford neuroprotection by decreasing intramitochondrial oxidative stress and/or augmenting wg/Wnt-like signaling; certain NSAIDs may also act through non-stereospecific 247 (implying COX2-independent) inhibition of . Moreover, if the demonstrated induction of ROS generation in astrocytes by amyloid349 is valid in vivo, it would constitute a mechanism, in addition to

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fatty acid oxidation and ketogenesis, for astrocytic production of diffusible oxidants as agents of neuronal injury. The oxidative amyloid cycle enunciated here, and other suggested formulations of the role of amyloid in Alzheimer disease350, are not mutually exclusive. Taken together, the above evidence suggests that although intrinsic neuronal antioxidant protective mechanisms are available they are inherently weak. As a result, neurons are largely dependent on the antioxidant defenses of the astrocyte. As a corollary, under those conditions in which potentially injurious oxidants emanate from the astrocyte itself, as the byproduct of its own oxidizing metabolic activity and oxidative stress (e.g., the hypothesized excessive plasma FFA-driven fatty acid oxidation and ketogenesis), the neuron faces double jeopardy.

15.5.2. Hypothesized Role of Fatty Acid Metabolism in Neurodegeneration Notwithstanding the abundance of evidence and the foregoing considerations, it remains to be determined whether neuronal utilization of ketone bodies in fact contributes to the oxidative stress, depressed utilization of glucose, and hypothesized “vicious cycle” that characterize Alzheimer disease pathogenesis. Plasma concentrations and brain utilization of FFA and ketone bodies have not been quantified in most reports. Thus, it is of considerable interest that in a study of global cerebral utilization of glucose, lactate, pyruvate, ketone bodies, FFA, and oxygen351, utilization of ketone bodies, fatty acids, and oxygen in Alzheimer patients did not differ significantly in absolute terms from controls, despite the expected and observed significantly lower rate of cerebral glucose metabolism in the Alzheimer group. However, it can be calculated from the data presented in this important experiment that in Alzheimer brain, utilization of ketone bodies and FFA were similar to controls but were increased relative to that of glucose, in terms of both substrate mass and maximal ATP-generating potential. Thus, assuming that 30 ATP molecules are generated for each molecule of glucose completely oxidized, and approximately 106 ATP molecules for each palmitic acid (whether direct or via oxidation of palmitate-derived ketone bodies), and correcting for measured pyruvate release, and lactate uptake (controls) or release (Alzheimers), the contribution of FFA and ketone bodies to ATP generation, as a percentage of that of glucose/lactate/pyruvate, could theoretically have increased from 49% in normals to 70% in the Alzheimer patients (both figures being somewhat high because of the relative inefficiency of fatty acid-supported ATP generation). Most importantly, as these observations necessarily


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described global cerebral metabolism, it seems virtually certain that changes in utilization of FFA and ketone bodies relative to glucose would have been significantly more pronounced in affected brain regions. The basis for such abnormalities in substrate utilization may reside within the neuron, or may be extraneuronal or extracranial. For example, local or systemic factors that augment fatty acid utilization and ketogenesis by astrocytes could increase the availability of ketone bodies in the interstitial space, shifting neuronal substrate utilization away from glucose (Fig. 15.2, and Chapter 12.3.: Ketone Bodies). Indeed, participation of systemic abnormalities in the pathogenesis of sporadic late onset Alzheimer disease, such as those affecting systemic fatty acid and FFA metabolism, would be consistent with, and possibly contribute to: 1) similar mitochondrial disturbances in blood platelets and brain in Alzheimer patients295; 2) evidence of increased susceptibility to apoptosis in periperal blood mononuclear cells from Alzheimer patients352; 3) an inverse relationship between cognitive decline and the ratio of fatty acids in erythrocyte membranes (see Chapter 4: Fatty Acids and Growth Regulation); and 4) other observations353. Similar findings in peripheral cells from patients with familial early-onset Alzheimer disease may reflect more widespread metabolic and energetic effects of mutant gene products or incidental systemic abnormalities354. Evidence that systemic processes may underlie sporadic late-onset Alzheimer disease is also provided by other observations. Thus, apoptotic death also occurs in glial cells in Alzheimer disease355, despite their stronger antioxidant defenses. This vulnerability may reflect the fact that 356 human brain astrocytes fail to express , and therefore may lack the protective adaptation to excess fatty acid exposure that mediates357. The potentially critical significance of astrocyte ketogenesis in neuronal metabolism and energetics is consistent with other evidence that supports a role for astrocytes in Alzheimer disease pathogenesis358. Over time, a neuronal substrate supply that is inappropriately enriched in astrocyte-generated ketone bodies and diffusible ROS would foster oxidative stress-induced injury to key components of neuronal energetics such as the electron transport chain and This metabolic milieu would account for decreases in glucose utilization and impaired cortical function that are characteristic in Alzheimer patients, and for the progressive and irreversible features of advancing disease. Systemic disorders that would foster such a sequence include obesity, insulin resistance, diabetes mellitus, and HIV infection. All of these may be associated with elevated plasma FFA and accelerated ketogenesis in liver (and astrocytes), and with sustained expression of proinflammatory cytokines which contribute to the inflammatory nature of the dementing

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process359, potentially compounded by fatty acid released through activation of brain phospholipase A2360. Of particular significance in this context is the fact that adipocyte expression is increased in obesity, may contribute to insulin resistance through its effects on insulin receptor signal transduction361, and interacts with both neuronal and glial cells in brain362-364. Consistent with these adverse consequences of abnormal lipid metabolism is evidence that dietary fat and glucose exert opposing effects on learning and memory in rats365, that ketogenic weight-reducing366 and seizure-suppressing367 diets have adverse cognitive effects, that long-term potentiation is impaired in genetic models of obesity368,369, and that brain dysfunction in aging rats and mice is associated with mitochondrial ROS generation64,370. Moreover, neuroprotection is afforded in rodents by measures which would attenuate this deleterious process, including dietary restriction371,372 and exercise373. In Alzheimer patients, protection may also be afforded by calorie restriction374, by dietary antioxidants375, by certain NSAIDs as discussed above, and by “statin” type hypocholesterolemic agents25,376,377. The latter agents modulate human cerebral cholesterol metabolism378, and diminish amyloid levels in vitro and in vivo379,380. Moreover, as discussed elsewhere in connection with the use of statins in atherosclerosis (see Chapter 7.4.1.: Atherosclerosis and Arterial Hypertension) the beneficial effects of these agents may transcend their modulation of serum lipid concentrations. Thus, at least in part, they may result from activation of SREBP-2 and the potentially antioxidant effects of a resulting acetyl CoA carboxylase/malonyl CoA-induced suppression of mitochondrial fatty acid oxidation. Statins also inhibit astrocyte secretion of apoE into the extracellular space, reflecting impaired prenylation of an as yet unidentified protein381. Aging rhesus monkeys are susceptible to amyloid-induced neurodegenerative changes, including phosphorylation, prior to plaque formation382. Finally, the prominent adrenergic effects of smoking-related nicotine administration increase adipocyte lipolysis and plasma [FFA]383-387, and by this mechanism could account at least in part for the suggested but epidemiologically still uncertain association between smoking and Alzheimer disease90,388-390. This issue is complicated by the fact that nicotinic agonists (e.g., nicotine) and muscarinic agonists are neuroprotective391-393, activate wg/Wnt-like PI3K/Akt/PKB signaling in respiratory epithelial cells394, enhance long term potentiation in vitro395, and acutely improve attention, information processing, and short term memory in human subjects396. Moreover, nicotine exerts anti-amyloid effects in vitro397. Thus, these relatively early, direct, and salutary neuronal effects of nicotine would appear to be inconsistent with Alzheimer-like sequelae and the mechanisms proposed here. However,


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recent studies indicate that amyloid-associated oxidative stress may induce loss of brain nicotinic receptors398, that accumulation of amyloid in Alzheimer disease is correlated with nicotinic acetylcholine receptor expression305, and that amyloid and nicotine may interact in suppressing long-term potentiation399. Further, the potential adverse effects of nicotine-stimulated adipocyte lipolysis and increases in plasma [FFA], sustained over many years, plausibly could override a more transient nicotinic receptor-mediated neuroprotection in some individuals. It will be important to resolve the relationship between these apparently conflicting consequences of chronic nicotine exposure and the pathogenesis of Alzheimer disease.

15.5.3. Integrated Model of Alzheimer Disease Pathogenesis The formulation developed in this chapter is based on the concept that changes in systemic fatty acid metabolism, and associated increases in hepatocellular and especially astrocytic fatty acid oxidation and ketogenesis, may contribute importantly to altered neuronal substrate utilization and oxidative stress. By themselves, these disturbances in Alzheimer disease may be sufficient: 1) to perturb neuronal intermediary metabolism and mitochondrial function; 2) to account for adverse changes in neuronal redox balance, glucose utilization, synaptic function and plasticity, and survival; and 3) to generate amyloid plaques and neurofibrillary tangles. In addition, they provide a mechanistic basis for the integration of numerous genetic and non-genetic determinants that are linked to this disorder. In this integrated model, a critical point of convergence may reside at the mutually amplifying interaction between oxidative stress and amyloid generation. Thus, available evidence suggests that in addition to its other adverse effects, oxidative stress-induced perturbation of APP processing is sufficient to promote amyloid generation, while amyloid’s effects on mitochondrial function are sufficient to induce oxidative stress. This “vicious cycle”, or feed-forward self-amplifying loop, may be augmented by genetic factors, such as mutated APP that increases amyloid generation, PS1 mutations that compromise neuroprotective wg/Wnt-like signaling and induce phosphorylation, and the E4 apolipoprotein isoform that, among its other potential effects, compromises redox balance, neuronal repair, and amyloid clearance. Importantly, the “vicious cycle” is subject to both activation and augmentation by non-genetic factors linked causally to altered fatty acid and ketone body metabolism, as may occur in obesity, insulin resistance, and type II diabetes mellitus; it is consistent with evidence for a role of

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insulin resistance in pathogenesis400, and may contribute importantly to the pathogenesis of sporadic, late-onset Alzheimer disease. Furthermore, variability among the components of this interplay could provide an explanation for inconsistent associations between various predisposing factors and clinically recognizeable disease. Similar mechanisms may also contribute to the pathogenesis of other neurodegenerative processes and HIV encephalopathy. As a corollary, the model may provide a basis for understanding the effects of certain agents potentially of preventive or therapeutic value401. These include dietary antioxidants, lithium, estrogens, NSAIDs, the “statin” and fibrate hypolipidemic drugs, ethanol, physical activity (see Chapter 7.4.: Consequences of Intramitochondrial Oxidative Stress: Atherosclerosis and Beyond), as well as cognitive activity itself402. Cognitive activity may be beneficial through its amelioration of oxidative stress as the result of increases in neuronal activation and cerebral blood flow, and therefore in glucose utilization. Verification of the hypothetical framework developed in this treatise will require further examination of the in vitro and in vivo interaction between astrocyte and neuron as a determinant of neuronal substrate utilization, redox balance, energetics, and synaptic function. If validated, it may open novel approaches to the prevention and treatment of Alzheimer disease and related disorders.

15.6.

REFERENCES

1. R. Mahley, B. Nathan, and R. Pitas, Apolipoprotein E: Structure, function, and possible roles in Alzheimer’s disease, Ann NY Acad Sci 777:139-145 (1996). 2. W. Strittmatter, and A. Roses, Apolipoprotein E and Alzheimer’s disease, Ann Rev Neurosci 19:53-77 (1996). 3. B. Yankner, Mechanisms of neuronal degeneration in Alzheimer’s disease, Neuron 16:921-932 (1996). 4. D. Selkoe, Alzheimer’s disease: genes, protein, and therapy, Physiol Rev 81:741-766 (2001). 5. J. Price, and J. Morris, Tangles and plaques in nondemented aging and “preclinical” Alzheimer’s disease, Ann Neural 45:358-368 (1999). 6. J. Hardy, and D. J. Selkoe, The amyloid hypothesis of Alzheimer’s disease: progress and problems on the road to therapeutics, Science 297:353-6 (2002). 7. N. Andreasen, E. Vanmechelen, H. Vanderstichele, P. Davidsson, and K. Blennow, Cerebrospinal fluid levels of total-tau, phospho-tau and A beta 42 predicts development of Alzheimer’s disease in patients with mild cognitive impairment, Acta Neurol Scand Suppl 170 (Suppl. 179):47-51 (2003). 8. T. Sunderland, G. Linker, N. Mirza, et al., Decreased beta-amyloid 1-42 and increased tau levels in cerebrospinal fluid of patients with Alzheimer disease, JAMA 289:2094103 (2003).


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9. K. Hirai, G. Aliev, A. Nunomura, et al., Mitochondrial abnormalities in Alzheimer’s disease, J Neurosci 21:3017-23 (2001). 10. R. Mahley, and S. Rall, Jr, Apolipoprotein E: Far more than a lipid transport protein, Annu Rev Genomics Hum Genet 1:507-37 (2000). 11. D. J. Selkoe, Deciphering the genesis and fate of amyloid beta-protein yields novel therapies for Alzheimer disease, J Clin Invest 110:1375-81 (2002). 12. B. Schwiegelshohn, J. Presley, M. Gorecki, et al., Effects of apoprotein E on intracellular metabolism of model triglyceride-rich particles are distinct from effects on cell particle uptake, J Biol Chem 270:1761-1769 (1995). 13. R. Pitas, J. Boyles, S. Lee, D. Foss, and R. Mahley, Astrocytes synthesize apolipoprotein E and metabolize apolipoprotein E-containing lipoproteins, Biochim Biophys Acta 917:148-161 (1987). 14. S. Han, G. Einstein, K. Weisgraber, et al., Apolipoprotein E is localized to the cytoplasm of human cortical neurons: a light and electron microscopic study, J Neuropathol Exp Neurol 53:535-544 (1994). 15. R. DeMattos, L. Curtiss, and D. Williams, A minimally lipidated form of cell-derived apolipoprotein E exhibits isoform-specific stimulation of neurite outgrowth in the absence of exogenous lipids or lipoproteins, J Biol Chem 273:4206-4212 (1998). 16. W. Pedersen, S. Chan, and M. Mattson, A mechanism for the neuroprotective effect of apolipoprotein E: isoform-specific modification by the lipid peroxidation product 4hydroxynonenal, J Neurochem 74:1426-1433 (2000). 17. T. Tokuda, M. Calero, E. Matsubara, et al., Lipidation of apolipoprotein E influences its isoform-specific interaction with Alzheimer’s amyloid beta peptides, Biochem J 348:358-365 (2000). 18. L. Myllykangas, T. Polvikoski, K. Reunanen, et al., ApoE epsilon3-haplotype modulates Alzheimer beta-amyloid deposition in the brain, Am J Med Genet 114:288291 (2002). 19. D. Mauch, K. Nagler, S. Schumacher, et al., CNS synaptogenesis promoted by gliaderived cholesterol, Science 294:1354-1357 (2001). 20. N. Bogdanovic, L. Bretillon, E. Lund, et al., On the turnover of brain cholesterol in patients with Alzheimer’s disease: Abnormal induction of the cholesterol-catabolic enzyme CYP46 in glial cells, Neurosci Lett 314:45-48 (2001B). 21. A. Cedazo-Minguez, and R. Cowburn, Apolipoprotein E isoform-specific disruption of phosphoinositide hydrolysis: protection by estrogen and glutathione, FEBS Lett 504:45-49 (2001). 22. Y. Hashimoto, H. Jiang, T. Niikura, et al., Neuronal apoptosis by apoplipoprotein E4 through low-density lipoprotein receptor-related protein and heterotrimeric GTPases, J Neurosci 20:8401-8409 (2000). 23. D. Kang, C. Pietrzik, L. Baum, et al., Modulation of amyloid beta-protein clearance and Alzheimer’s disease susceptibility by the LDL receptor-related protein pathway, J Clin Invest 106:1159-1166 (2000). 24. M. Shibata, S. Yamada, S. Kumar, et al., Clearance of Alzheimer’s amyloid-beta(1-40) peptide from brain by LDL receptor-related protein-1 at the blood-brain barrier, J Clin Invest 106:1489-1499 (2000). 25. L. Puglielli, R. E. Tanzi, and D. M. Kovacs, Alzheimer’s disease: the cholesterol connection, Nat Neurosci 6:345-51 (2003). 26. M. Meyer, J. Tschanz, M. Norton, et al., APOE genotype predicts when—not whether—one is predisposed to develop Alzheimer disease, Nature Genet 1199:321322 (1998).

332

Chapter 15

27. S. Craft, L. Teri, S. Edland, et al., Accelerated decline in apolipoprotein E-epsilon4 homozygotes with Alzheimer’s disease, Neurology 51:149-53 (1998B). 28. E. Reiman, R. Caselli, L. Yun, et al., Preclinical evidence of Alzheimer’s disease in persons homozygous for the epsilon 4 allele for apolipoprotein E, New Engl J Med 334:752-8 (1996). 29. E. Reiman, R. Caselli, K. Chen, G. Alexander, D. Bandy, and J. Frost, Declining brain activity in cognitively normal apolipoprotein E epsilon4 heterozygotes: A foundation for using positron emission tomography to efficiently test treatments to prevent Alzheimer’s disease, Proc Natl Acad Sci USA 98:3334-3339 (2001). 30. N. R. Graff-Radford, R. C. Green, R. C. Go, et al., Association between apolipoprotein E genotype and Alzheimer disease in African American subjects, Arch Neurol 59:594-600 (2002). 31. P. Greenwood, T. Sunderland, J. Friz, and R. Parasuraman, Genetics and visual attention: selective deficits in healthy adult carriers of the epsilon 4 allele of the apolipoprotein E gene, Proc Natl Acad Sci USA 97:11661-11666 (2000). 32. G. Gibson, V. Haroutunian, H. Zhang, et al., Mitochondrial damage in Alzheimer’s disease varies with apolipoprotein E genotype, Ann Neurol 48:297-303 (2000). 33. D. Schmechel, A. Saunders, W. Strittmatter, et al., Increased amyloid beta-peptide deposition in cerebral cortex as a consequence of apolipoprotein E genotype in lateonset Alzheimer disease, Proc Natl Acad Sci USA 90:9649-9653 (1993). 34. D. Cook, L. JB, P. McMillan, et al., Reduced hippocampal insulin-degrading enzyme in late-onset Alzheimer’s disease is associated with the apolipoprotein E-episilon4 allele, American Journal of Pathology 162:313-319 (2003). 35. E. Corder, K. Robertson, L. Lannfelt, et al., HIV-infected subjects with the E4 allele for APOE have excess dementia and peripheral neuropathy, Nature Med 4:1182-1184 (1998). 36. B. Tardiff, M. Newman, A. Saunders, et al., Preliminary report of a genetic basis for cognitive decline after cardiac operations: The Neurologic Outcome Research Group of the Duke Heart Center, Ann Thorac Surg 64:715-720 (1997). 37. B. Jordan, N. Relkin, L. Ravdin, A. Jacobs, A. Bennett, and S. Gandy, Apolipoprotein E epsilon4 associated with chronic traumatic brain injury in boxing, J Am Med Assoc 278:136-140 (1997). 38. G. Teasdale, J. Nicoll, G. Murray, and M. Fiddes, Association of apolipoprotein E polymorphism with outcome after head injury, Lancet 350:1069-1071 (1997). 39. D. M. Holtzman, K. R. Bales, T. Tenkova, et al., Apolipoprotein E isoformdependent amyloid deposition and neuritic degeneration in a mouse model of Alzheimer’s disease, Proc Natl Acad Sci U S A 97:2892-7 (2000). 40. Z. S. Ji, R. D. Miranda, Y. M. Newhouse, K. H. Weisgraber, Y. Huang, and R. W. Mahley, Apolipoprotein E4 potentiates amyloid beta peptide-induced lysosomal leakage and apoptosis in neuronal cells, J Biol Chem 277:21821-8 (2002). 41. R. Mayeux, A. Saunders, S. Shea, et al., Utility of the apolipoprotein E genotype in the diagnosis of Alzheimer’s disease. Alzheimer’s Disease Centers Consortium on Apolipoprotein E and Alzheimer’s Disease, New Engl J Med 338:506-511 (1998). 42. G. Small, L. Ercoli, D. Silverman, et al., Cerebral metabolic and cognitive decline in persons at genetic risk for Alzheimer’s disease, Proc Natl Acad Sci USA 97:60376042 (2000). 43. S. Rapoport, Functional brain imaging to identify affected subjects genetically at risk for Alzheimer’s disease, Proc Natl Acad Sci USA 97:5696-5698 (2000).


333

44. G. Alexander, K. Chen, P. Pietrini, S. Rapoport, and E. Reiman, Longitudinal PET Evaluation of Cerebral Metabolic Decline in Dementia: A Potential Outcome Measure in Alzheimer’s Disease Treatment Studies, Am J Psychiatry 159:738-745 (2002). 45. V. Mooser, N. Helbecque, J. Miklossy, S. Marcovina, P. Nicod, and P. Amouyel, Interactions between apolipoprotein E and apolipoprotein(a) in patients with lateonset Alzheimer disease, Annals Internal Med 132:533-537 (2000). 46. L. Fairer, L. Cupples, J. Haines, et al., Effects of age sex and ethnicity on the association between apolipoprotein E genotype and Alzheimer disease: A metaanalysis. APOE and Alzheimer Disease Meta Analysis Consortium, J Am Med Assoc 278:1349-1356 (1997). 47. M. Tang, Y. Stern, K. Marder, et al., The APOE-epsilon4 allele and the risk of Alzheimer disease among African Americans, Whites and Hispanics, J Am Med Assoc 279:751-755 (1998). 48. G. Thinakaran, The role of presenilins in Alzheimer’s disease, Journal of Clinical Investigation 104:1321-7 (1999). 49. O. Murayama, T. Tomita, N. Nihonmatsu, et al., Enhancement of amyloid beta 42 secretion by 28 different presenilin 1 mutations of familial Alzheimer’s disease, Neurosci Lett 265:61-63 (1999). 50. R. Kopan, and A. Goate, A common enzyme connects notch signaling and Alzheimer’s disease, Genes Devel 14:2799-2806 (2000). 51. S. Sisodia, and P. St. George-Hyslop, gamma-Secretase, Notch, Abeta and Alzheimer’s disease: where do the presenilins fit in?, Nat Rev Neurosci 3:281-290 (2002). 52. W. Dowjat, H. Wisniewski, and T. Wisniewski, Alzheimer’s disease presenilin-1 expression modulates the assembly of neurofilaments, Neurosci 103:1-8 (2001). 53. V. Askanas, W. Engel, C.-C. Yang, R. Alvarez, V.-Y. Lee, and T. Wisniewski, Light and electron microscopic immunolocalization of Presenilin 1 in abnormal muscle fibers of patients with sporadic inclusion-body myositis and autosomal-recessive inclusion-body myopathy, Am J Pathol 152:889-895 (1998A). 54. V. Askanas, and W. Engel, Does overexpression of betaAPP in aging muscle have a pathogenic role and a relevance to Alzheimer’s disease? Clues from inclusion body myositis cultured human muscle and transgenic mice, Am J Path 153:1673-1677 (1998B). 55. G. Vattemi, W. Engel, J. McFerrin, J. Buxbaum, L. Pastorino, and V. Askanas, Presence of BACE1 and BACE2 in muscle fibres of patients with sporadic inclusionbody myositis, Lancet 358:1962-1964 (2001). 56. M. S. Brown, J. Ye, R. B. Rawson, and J. L. Goldstein, Regulated intramembrane proteolysis: a control mechanism conserved from bacteria to humans, Cell 100:3918 (2000). 57. S. Soriano, D. E. Kang, M. Fu, et al., Presenilin 1 negatively regulates beta-catenin/T cell factor/lymphoid enhancer factor-1 signaling independently of beta-amyloid precursor protein and notch processing, J Cell Biol 152:785-94 (2001). 58. X. Xia, S. Qian, S. Soriano, et al., Loss of presenilin 1 is associated with enhanced beta-catenin signaling and skin tumorigenesis, Proc Natl Acad Sci U S A 98:10863-8 (2001). 59. D. Hartmann, J. Toumoy, P. Saftig, W. Annaert, and B. De Strooper, Implication of APP secretases in notch signaling, J Mol Neurosci 17:171-81 (2001). 60. L. Bertram, D. Blacker, K. Mullin, et al., Evidence for genetic linkage of Alzheimer’s disease to chromosome l0q, Science 290:2302-2303 (2000).

334

Chapter 15

61. N. Ertekin-Taner, N. Graff-Radford, L. Younkin, et al., Linkage of plasma Abeta42 to a quantitative locus on chromosome 10 in late-onset Alzheimer’s disease pedigrees, Science 290:2303-2304 (2000). 62. A. Myers, P. Holmans, H. Marshall, et al., Susceptibility locus for Alzheimer’s disease on chromosome 10, Science 290:2304-2305 (2000). 63. A. Takeuchi, M. Irizarry, K. Duff, et al., Age-related amyloid beta deposition in transgenic mice overexpressing both Alzheimer mutant presenilin 1 and amyloid beta precursor protein Swedish mutant is not associated with global neuronal loss, Am J Pathol 157:331-339 (2000). 64. J. Liu, D. W. Killilea, and B. N. Ames, Age-associated mitochondrial oxidative decay: improvement of carnitine acetyltransferase substrate-binding affinity and activity in brain by feeding old rats acetyl-L- carnitine and/or R-alpha -lipoic acid, Proc Natl Acad Sci U S A 99:1876-81 (2002). 65. G. Fiskum, A. Murphy, and M. Beal, Mitochondria in neurodegeneration: acute ischemia and chronic neurodegenerative diseases, J Cerebral Blood Flow Metab 19:351-369 (1999). 66. M. P. Mattson, S. L. Chan, and W. Duan, Modification of brain aging and neurodegenerative disorders by genes, diet, and behavior, Physiol Rev 82:637-72 (2002). 67. D. Kuhl, E. Metter, W. Riege, and R. Hawkins, The effect of normal aging on patterns of local cerebral glucose utilization, Ann Neurol 15:S133-S137 (1984). 68. G. Reaven, L. Thompson, D. Nahum, Haskins, and E, Relationship between hyperglycemia and cognitive function in older NIDDM patients, Diabetes Care 13:1621 (1990). 69. A. McCall, The impact of diabetes on the CNS, Diabetes 41:557-570 (1992). 70. G. J. Biessels, L. P. van der Heide, A. Kamal, R. L. Bleys, and W. H. Gispen, Ageing and diabetes: implications for brain function, Eur J Pharmacol 441:1-14 (2002). 71. J. Heitner, and D. Dickson, Diabetics do not have increased Alzheimer-type pathology compared with age-matched control subjects:A retrospective postmortem immunocytochemical and histofluorescent study, Neurol 49:1306-1311 (1997). 72. J. Halter, Alzheimer’s disease and non-insulin-dependent diabetes mellitus: common features do not make common bedfellows, J Am Geriatr Soc 44:992-993 (1996). 73. C. Brayne, F. Matthews, J. Xuereb, J. Broome, a. et, and m. o. t. N. G. o. t. M. C. F. a. A. Study), Pathological correlates of late-onset dementia in a multicentre, community-based population in England and Wales, Lancet 357:169-175 (2001). 74. A. Barber, E. Lieth, S. Khin, D. Antonetti, A. Buchanan, and T. Gardner, Neural apoptosis in the retina during experimental and human diabetes:Early onset and effect of insulin, J Clin Invest 102:783-791 (1998). 75. L. V. Johnson, W. P. Leitner, A. J. Rivest, M. K. Staples, M. J. Radeke, and D. H. Anderson, The Alzheimer’s A beta -peptide is deposited at sites of complement activation in pathologic deposits associated with aging and age-related macular degeneration, Proc Natl Acad Sci U S A 99:11830-5 (2002). 76. S. Yamagishi, T. Okamoto, S. Amano, et al., Palmitate-induced apoptosis of microvascular endothelial cells and pericytes, Mol Med 8:179-84 (2002). 77. S. E. Vermeer, N. D. Prins, T. den Heijer, A. Hofman, P. J. Koudstaal, and M. M. Breteler, Silent brain infarcts and the risk of dementia and cognitive decline, N Engl J Med 348:1215-22 (2003). 78. S. Kalmijn, E. Feskens, L. Launer, T. Stijnen, Kromhout, and D, Glucose intolerance, hyperinsulinaemia and cognitive function in a general population of elderly men, Diabetologia 38:1096-1102 (1995).


335

79. A. Ott, R. Stolk, A. Hofman, F. van Harskamp, D. Grobbee, and M. Breteler, Association of diabetes mellitus and dementia: the Rotterdam Study, Diabetologia 39:1392-1397 (1996). 80. J. Kuusisto, K. Koivisto, L. Mykkänen, et al., Association between features of the insulin resistance syndrome and Alzheimer’s disease independently of apolipoprotein E4 phenotype: cross sectional population based study, Brit Med J 315:1045-104 (1997). 81. C. Leibson, W. Rocca, V. Hanson, et al., Risk of dementia among persons with diabetes mellitus: a population-based cohort study, Am J Epidemiol 145:301-308 (1997). 82. E. Gregg, K. Yaffe, J. Cauley, et al., Is diabetes associated with cognitive impairment and cognitive decline among older women?, Arch Intern Med 160:174-180 (2000). 83. D. Bennett, Diabetes and change in cognitive function, Arch Intern Med 160:141143 (2000). 84. E. Feskens, L. Havekes, S. Kalmijn, P. de Knijff, L. Launer, and D. Kromhout, Apolipoprotein e4 allele and cognitive decline in elderly men, Brit Med J 309:12021206 (1994). 85. G. Razay, and G. Wilcock, Hyperinsulinaemia and Alzheimer’s disease, Age Ageing 23:396-399 (1994). 86. S. Craft, E. Peskind, M. Schwartz, G. Schellenberg, M. Raskind, and D. Porte, Jr, Cerebrospinal fluid and plasma insulin levels in Alzheimer’s disease: relationship to severity of dementia and apolipoprotein E genotype, Neurol 50:164-168 (1998A). 87. M. Haan, L. Shemanski, W. Jagust, T. Manolio, and L. Kuller, The role of APOE epsilon4 in modulating effects of other risk factors for cognitive decline in elderly persons, JAMA 282:40-46 (1999). 88. J. Molina, F. Jimenez-Jimenez, C. Vargas, et al., Cerebrospinal fluid levels of insulin in patients with Alzheimer’s disease, Acta Neurol Scand 106:347-350 (2002). 89. S. Kalmijn, L. Launer, A. Ott, J. Witteman, A. Hofman, and M. Breteler, Dietary fat intake and the risk of incident dementia in the Rotterdam Study, Ann Neurol 42:776782 (1997). 90. A. Ott, A. Slooter, A. Hofman, et al., Smoking and risk of dementia and Alzheimer’s disease in a population-based cohort study: the Rotterdam Study, Lancet 351:18401843 (1998). 91. J. Orgogozo, J. Dartigues, S. Lafont, et al., Wine consumption and dementia in the elderly: a prospective community study in the Bordeaux area., Rev Neurol 153:185192 (1997). 92. A. Ruitenberg, J. van Swieten, J. Witteman, et al., Alcohol consumption and risk of dementia: the Rotterdam Study, Lancet 359:281-286 (2002). 93. K. J. Mukamal, L. H. Kuller, A. L. Fitzpatrick, W. T. Longstreth, Jr., M. A. Mittleman, and D. S. Siscovick, Prospective study of alcohol consumption and risk of dementia in older adults, Jama 289:1405-13 (2003). 94. L. Refolo, M. Pappolla, B. Malester, et al., Hypercholesterolemia accelerates the Alzheimer’s amyloid pathology in a transgenic mouse model, Neurobiol Dis 7:321331 (2000). 95. S. Hoyer, The brain insulin signal transduction system and sporadic (type II) Alzheimer disease: an update, J Neural Transm 109:341-360 (2002). 96. V. Grill, M. Gutniak, O. Björkman, et al., Cerebral blood flow and substrate utilization in insulin-treated diabetic subjects, Am J Physiol 258:E813-E820 (1990).

336

Chapter 15

97. M. Gutniak, G. Blomqvist, L. Widén, S. Stone-Elander, B. Hamberger, and V. Grill, D-[U-11C]glucose uptake and metabolism in the brain of insulin-dependent diabetic subjects, Am J Physiol 258:E805-E812 (1990). 98. C. Fanelli, C. Dence, J. Markham, et al., Blood-to-brain glucose transport and cerebral glucose metabolism are not reduced in poorly controlled type 1 diabetes, Diabetes 47:1444-1450 (1998). 99. S. Hasselbalch, G. Knudsen, C. Videbaek, et al., No effect of insulin on glucose blood-brain barrier transport and cerebral metabolism in humans, Diabetes 48:19151921 (1999). 100. P. Freychet, Insulin receptors and insulin actions in the nervous system, Diabetes/Metab Res Rev 16:390-392 (2000). 101. C. Park, Cognitive effects of insulin in the central nervous system, Neurosci Biobehav Rev 25:311-323 (2001). 102. W. Zhao, H. Chen, H. Xu, et al., Brain insulin receptors and spatial memory: Correlated changes in gene expression, tyrosine phosphorylation, and signaling molecules in the hippocampus of water maze trained rats, J Biol Chem 274:3489334902 (1999). 103. A. Chaudhuri, Y. Kanjwal, P. Mohanty, et al., Insulin-induced vasodilatation of internal carotid artery, Metab Clin Exp 48:1470-1473 (1999). 104. L. Reagan, A. Magariños, D. Yee, et al., Oxidative stress and HNE conjugation of GLUT3 are increased in the hippocampus of diabetic rats subjected to stress, Brain Res 862:292-300 (2000). 105. S. M. Manschot, G. J. Biessels, N. E. Cameron, et al., Angiotensin converting enzyme inhibition partially prevents deficits in water maze performance, hippocampal synaptic plasticity and cerebral blood flow in streptozotocin-diabetic rats, Brain Res 966:274-82 (2003). 106. M. Aragno, S. Parola, E. Tamagno, et al., Oxidative derangement in rat synaptosomes induced by hyperglycaemia: restorative effect of dehydroepiandrosterone treatment, Biochem Pharmacol 60:389-395 (2000). 107. P. Pietrini, M. Furey, G. Alexander, et al., Association between brain functional failure and dementia severity in Alzheimer’s disease: Resting versus stimulation PET study, Am J Psych 156:470-473 (1999). 108. K. Meguro, X. Blaizot, Y. Kondoh, M. Le, C, J. Baron, and C. Chavoix, Neocortical and hippocampal glucose heypometabolism following neurotixic lesions of the entorhinal and perithinal cortices in the non-human primate as shown by PET: Implications for Alzheimer’s disease, Brain 122:1519-1531 (1999). 109. A. Kennedy, R. Frackowiak, S. Newman, et al., Deficits in cerebral glucose metabolism demonstrated by positron emission tomography in individuals at risk of familial Alzheimer’s disease, Neurosci Lett 186:17-20 (1995). 110. K. Chandrasekaran, K. Hatanpää, D. Brady, and S. Rapoport, Evidence for physiological down-regulation of brain oxidative phosphorylation in Alzheimer’s disease, Exp Neurol 142:80-88 (1996). 111. G. Small, S. Komo, A. La Rue, et al., Early detection of Alzheimer’s disease by combining apolipoprotein E and neuroimaging, Ann NY Acad Sci 802:70-78 (1996). 112. S. Rapoport, K. Hatanpää, D. Brady, Chandrasekaran, and K, Brain energy metabolism, cognitive function and down-regulated oxidative phosphorylation in Alzheimer disease, Neurodegeneration 5:473-476 (1996). 113. S. Rapoport, In vivo PET imaging and postmortem studies suggest potentially reversible and irreversible stages of brain metabolic failure in Alzheimer’s disease, Eur Arch Psych Clin Neurosci 249Suppl3:46-55 (1999B).


337

114. G. Chetelat, and J. C. Baron, Early diagnosis of Alzheimer’s disease: contribution of structural neuroimaging, Neuroimage 18:525-41 (2003). 115. G. Stoppe, H. Bruhn, P. Pouwels, W. Hänicke, and J. Frahm, Alzheimer disease: Absolute quantification of cerebral metabolites in vivo using localized proton magnetic resonance spectroscopy, Alzheimer Dis Assoc Disorders 14:112-119 (2000). 116. V. Ibáñez, P. Pietrini, G. Alexander, et al., Regional glucose metabolic abnormalities are not the result of atrophy in Alzheimer’s disease, Neurology 50:1585-1593 (1998). 117. K. Chandrasekaran, K. Hatanpää, D. Brady, J. Stoll, and S. Rapoport, Downregulation of oxidative phosphorylation in Alzheimer disease: loss of cytochrome oxidase subunit mRNA in the hippocampus and entorhinal cortex, Brain Res 796:13-19 (1998). 118. J. Blass, and G. Gibson, The role of oxidative abnormalities in the pathophysiology of Alzheimer’s disease, Rev Neurol 147:513-525 (1991). 119. F. Mastrogiacomo, C. Bergeron, and S. Kish, Brain alpha-ketoglutarate dehydrogenase complex activity in Alzheimer’s disease, J Neurochem 61:2007-2014 (1993). 120. K. Sheu, A. Cooper, K. Koike, M. Koike, J. Lindsay, and J. Blass, Abnormality of the alpha-ketoglutarate dehydrogenase complex in fibroblasts from familial Alzheimer’s disease, Ann Neurol 35:312-38 (1994). 121. S. Hoyer, Oxidative metabolism deficiencies in brains of patients with Alzheimer’s disease, Acta Neurol Scand Suppl 165:18-24 (1996). 122. G. Gibson, H. Zhang, K. Sheu, et al., Alpha-ketoglutarate dehydrogenase in Alzheimer brains bearing the APP670/671 mutation, Ann Neurol 44:676-681 (1998). 123. A. Nulton-Persson, and L. Szweda, Modulation of mitochondrial function by hydrogen peroxide, J Biol Chem 276:23357-23361 (2001). 124. L. Frölich, D. Blum-Degen, H. Bernstein, et al., Brain insulin and insulin receptors in aging and sporadic Alzheimer’s disease, J Neural Transmission 105:423-438 (1998). 125. R. Russell, 3d, and H. Taegtmeyer, Changes in citric acid cycle flux and anaplerosis antedate the functional decline in isolated rat hearts utilizing acetoacetate, J Clin Invest 87:384-390 (1991). 126. R. Russell, 3d, and H. Taegtmeyer, Coenzyme A sequestration in rat hearts oxidizing ketone bodies, J Clin Invest 89:968-973 (1992). 127. L. Tretter, and V. Adam-Vizi, Inhibition of Krebs cycle enzymes by hydrogen peroxide: A key role of [alpha]-ketoglutarate dehydrogenase in limiting NADH production under oxidative stress, J Neurosci 20:8972-8979 (2000). 128. V. I. Bunik, and C. Sievers, Inactivation of the 2-oxo acid dehydrogenase complexes upon generation of intrinsic radical species, Eur J Biochem 269:5004-15 (2002). 129. S. Kish, Brain energy metabolizing enzymes in Alzheimer’s disease: alphaketoglutarate dehydrogenase complex and cytochrome oxidase, Ann NY Acad Sci 826:218-228 (1997). 130. F. Mastrogiacomo, J. Lindsay, L. Bettendorff, J. Rice, and S. Kish, Brain protein and alpha-ketoglutarate dehydrogenase complex activity in Alzheimer’s disease, Ann Neurol 39:592-598 (1996). 131. N. Sims, Energy metabolism, oxidative stress and neuronal degeneration in Alzheimer’s disease, Neurodegeneration 5:435-440 (1996).

338

Chapter 15

132. J. Bolaños, A. Almeida, E. Fernández, et al., Potential mechanisms for nitric oxidemediated impairment of brain mitochondrial energy metabolism, Biochem Soc Transact 25:944-949 (1997A). 133. J. Bolaños, A. Almeida, V. Stewart, et al., Nitric oxide-mediated mitochondrial damage in the brain: mechanisms and implications for neurodegenerative diseases, J Neurochem 68:2227-2240 (1997B). 134. J. Wilson, Antioxidant defense of the brain: a role for astrocytes, Can J Physiol Pharmacol 75:1149-1163 (1997). 135. B. Beltrán, A. Mathur, M. Duchen, J. Erusalimsky, and S. Moncada, The effect of nitric oxide on cell respiration: A key to understanding its role in cell survival or death, Proc Natl Acad Sci USA 97:14602-14607 (2000). 136. P. García-Nogales, A. Almeida, and J. Bolaños, Peroxynitrite protects neurons against nitric oxide-mediated apoptosis. A key role for glucose-6-phosphate dehydrogenase in neuroprotection., Journal of Biological Chemistry 278:864-874 (2003). 137. R. Ockner, N. Lysenko, N. Wu, and N. Bass, Hepatocyte growth inhibitors modulate mitochondrial and extramitochondrial fatty acid oxidation [Abstract], Hepatology 24:253A (1996). 138. M. Joaquin, J. Rosa, C. Salvado, et al., Hepatocyte growth factor and transforming growth factor beta regulate 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase gene expression in rat hepatocyte primary cultures, Biochem J 314:235-240 (1996). 139. G. Thoresen, and T. Christoffersen, Transforming growth factor beta 1 increases the phosphoenolpyruvate carboxykinase mRNA level in cultured rat hepatocytes, Cell Biol Intl 18:171-175 (1994). 140. Y. H. Hong, H. B. Peng, V. La Fata, and J. K. Liao, Hydrogen peroxide-mediated transcriptional induction of macrophage colony-stimulating factor by TGF-beta1, J Immunol 159:2418-23 (1997). 141. B. Herrera, A. M. Alvarez, A. Sanchez, et al., Reactive oxygen species (ROS) mediates the mitochondrial-dependent apoptosis induced by transforming growth factor (beta) in fetal hepatocytes, Faseb J 15:741-751 (2001B). 142. B. Herrera, M. Fernandez, A. M. Alvarez, et al., Activation of caspases occurs downstream from radical oxygen species production, Bcl-xL down-regulation, and early cytochrome C release in apoptosis induced by transforming growth factor beta in rat fetal hepatocytes, Hepatology 34:548-56 (2001 A). 143. E. Masliah, G. Ho, and T. Wyss-Coray, Functional role of TGF beta in Alzheimer’s disease microvascular injury: lessons from transgenic mice, Neurochem Int 39:393400 (2001). 144. A. Ruocco, O. Nicole, F. Docagne, et al., A transforming growth factor-beta antagonist unmasks the neuroprotective role of this endogenous cytokine in excitotoxic and ischemic brain injury, J Cereb Blood Flow Metab 19:1345-1353 (1999). 145. M. Packard, D. Mathew, and V. Budnik, Wnts and TGF beta in synaptogenesis: old friends signalling at new places, Nat Rev Neurosci 4:113-20 (2003). 146. S. Hoyer, Oxidative energy metabolism in Alzheimer brain: Studies in early-onset and late-onset cases, Molec Chem Neuropath 16:207-224 (1992). 147. H. Fukuyama, M. Ogawa, H. Yamauchi, et al., Altered cerebral energy metabolism in Alzheimer’s disease: a PET study, J Nucl Med 35:1-6 (1994). 148. S. Hasselbalch, P. Madsen, L. Hageman, et al., Changes in cerebral blood flow and carbohydrate metabolism during acute hyperketonemia, Am J Physiol 270:E746-51 (1996).


339

149. S. Bookheimer, M. Strojwas, M. Cohen, et al., Patterns of brain activation in people at risk for Alzheimer’s disease, N Engl J Med 343:450-456 (2000). 150. J. Zhou, U. Liyanage, M. Medina, et al., Presenilin 1 interaction in the brain with a novel member of the Armadillo family, Neuroreport 8:2085-2090 (1997). 151. G. Yu, F. Chen, G. Levesque, et al., The presenilin 1 protein is a component of a high molecular weight intracellular complex that contains beta-catenin, J Biol Chem 273:16470-16475 (1998). 152. A. Takashima, M. Murayama, O. Murayama, et al., Presenilin 1 associates with glycogen synthase kinase-3beta and its substrate tau, Proc Natl Acad Sci USA 95:9637-9641 (1998). 153. G. Levesque, G. Yu, M. Nishimura, et al., Presenilins interact with armadillo proteins including neural-specific plakophilin-related protein and beta-catenin, J Neurochem 72:999-1008 (1999). 154. F. Kirschenbaum, S. Hsu, B. Cordell, and J. McCarthy, Substitution of a glycogen synthase kinase-3beta phosphorylation site in presenilin 1 separates presenilin function from beta-catenin signaling, J Biol Chem 276:7366-7375 (2001). 155. M. Murayama, S. Tanaka, J. Palacino, et al., Direct association of presenilin-1 with beta-catenin, FEBS Lett 433:73-77 (1998). 156. E. Planel, X. Sun, and A. Takeshima, Role of GSK-3beta in Alzheimer’s disease pathology, Drug Dev Res 56:491-510 (2002). 157. M. Peifer, Beta-catenin as oncogene: the smoking gun, Science 275:1752-1753 (1997). 158. M. Hart, R. de los Santos, I. Albert, B. Rubinfeld, and P. Polakis, Downregulation of beta-catenin by human Axin and its association with the APC tumor suppressor, betacatenin and GSK3 beta, Current Biol 8:573-581 (1998). 159. J. Brakeman, S. Gu, X. Wang, G. Dolin, and J. Baraban, Neuronal localization of the Adenomatous polyposis coli tumor suppressor protein, Neurosci 91:661-572 (1999). 160. G. De Ferrari, and N. Inestrosa, Wnt signaling function in Alzheimer’s disease, Brain Res Rev 33:1-12 (2000). 161. A. Patapoutian, and L. Reichardt, Roles of Wnt proteins in neural development and maintenance, Curr Opin Neurobiol 10:392-399 (2000). 162. C. Sutherland, I. Leighton, and P. Cohen, Inactivation of glycogen synthase kinase3 beta by phosphorylation: new kinase connections in insulin and growth-factor signalling, Biochem J 296:15-19 (1993). 163. H. Eldar-Finkelman, and E. Krebs, Phosphorylation of insulin receptor substrate 1 by glycogen synthase kinase 3 impairs insulin action, Proc Natl Acad Sci USA 94:9660-9664 (1997). 164. P. Shepherd, D. Withers, and K. Siddle, Phosphoinositide 3-kinase: the key switch mechanism in insulin signalling, Biochem J 333:471-490 (1998). 165. M. Delcommenne, C. Tan, V. Gray, L. Rue, J. Woodgett, and S. Dedhar, Phosphoinositide-3-OH kinase-dependent regulation of glycogen synthase kinase 3 and protein kinase B/AKT by the integrin-linked kinase, Proc Natl Acad Sci USA 95:11211-11216 (1998). 166. J. Papkoff, and M. Aikawa, WNT-1 and HGF regulate GSK3- beta activity and betacatenin signaling in mammary epithelial cells, Biochem Biophys Res Comm 247:851-858 (1998). 167. N. Klöcker, P. Kermer, J. Weishaupt, M. Labes, R. Ankerhold, and M. Bähr, Brainderived neurotrophic factor-mediated neuroprotection of adult rat retinal ganglion cells in vivo does not exclusively depend on phosphatidyl-inositol-3’-kinase/protein kinase B signaling, J Neurosci 20:6962-6967 (2000).

340

Chapter 15

168. A. Vaillant, I. Mazzoni, C. Tudan, M. Boudreau, D. Kaplan, and F. Miller, Depolarization and neurotrophins converge on the phosphatidylinositol 3-kinase-Akt pathway to synergistically regulate neuronal survival, J Cell Biol 146:955-966 (1999). 169. P. Bannasch, F. Klimek, and D. Mayer, Early bioenergetic changes in hepatocarcinogenesis: preneoplastic phenotypes mimic responses to insulin and thyroid hormone, J Bioenergetics Biomembranes 29:303-313 (1997). 170. D. Nehrbass, F. Klimek, and P. Bannasch, Overexpression of insulin receptor substrate-1 emerges early in hepatocarcinogenesis and elicits preneoplastic hepatic glycogenosis, Am J Pathol 152:341-345 (1998). 171. B. Anderton, Alzheimer’s disease: clues from flies and worms, Curr Biol 9:R106R109 (1999). 172. R. Bhat, J. Shanley, M. Correll, et al., Regulation and localization of tyrosine216 phosphorylation of glycogen synthase kinase-3beta in cellular and animal models of neuronal degeneration, Proc Natl Acad Sci USA 97:11074-11079 (2000). 173. M. Hetman, J. Cavanaugh, D. Kimelman, and Z. Xia, Role of glycogen synthase kinase-3beta in neuronal apoptosis induced by trophic withdrawal, J Neurosci 20:2567-2574 (2000). 174. A. Georgakopoulos, P. Marambaud, S. Efthimiopoulos, et al., Presenilin-1 forms complexes with the cadherin/catenin cell-cell adhesion system and is recruited to intercellular and synaptic contacts, Mol Cell 4:893-902 (1999). 175. P. Marambaud, J. Shioi, G. Serban, et al., A presenilin-1/gamma-secretase cleavage releases the E-cadherin intracellular domain and regulates disassembly of adherens junctions, EMBO J 21:1948-1956 (2002). 176. W. Nishimura, I. Yao, J. Iida, N. Tanaka, and Y. Hata, Interaction of synaptic scaffolding molecule and Beta -catenin, J Neurosci 22:757-765 (2002). 177. M. Hong, and V. Lee, Insulin and insulin-like growth factor-1 regulate tau phosphorylation in cultured human neurons, J Biol Chem 272:19547-19553 (1997A). 178. J. Muñoz-Montaño, F. Moreno, J. Avila, and J. Diaz-Nido, Lithium inhibits Alzheimer’s disease-like tau protein phosphorylation in neurons, FEBS Lett 411:183-188 (1997). 179. B. Ryu, H. Ko, I. Jou, J. Noh, and B. Gwag, Phosphatidylinositol 3-kinase-mediated regulation of neuronal apoptosis and necrosis by insulin and IGF-I, J Neurobiol 39:536-546 (1999). 180. L. Gasparini, W. J. Netzer, P. Greengard, and H. Xu, Does insulin dysfunction play a role in Alzheimer’s disease?, Trends Pharmacol Sci 23:288-93 (2002). 181. D. C. Solano, M. Sironi, C. Bonfini, S. B. Solerte, S. Govoni, and M. Racchi, Insulin regulates soluble amyloid precursor protein release via phosphatidyl inositol 3 kinase-dependent pathway, Faseb J 14:1015-22 (2000). 182. C. J. Phiel, C. A. Wilson, V. M. Lee, and P. S. Klein, GSK-3alpha regulates production of Alzheimer’s disease amyloid-beta peptides, Nature 423:435-9 (2003). 183. G. Cheng, Z. Yu, D. Zhou, and M. P. Mattson, Phosphatidylinositol-3-kinase-Akt kinase and p42/p44 mitogen-activated protein kinases mediate neurotrophic and excitoprotective actions of a secreted form of amyloid precursor protein, Exp Neural 175:407-14 (2002). 184. L. Iacovelli, V. Bruno, L. Salvatore, et al., Native group-III metabotropic glutamate receptors are coupled to the mitogen-activated protein kinase/phosphatidylinositol-3kinase pathways, J Neurochem 82:216-23 (2002).


341

185. I. Tsujio, T. Tanaka, T. Kudo, et al., Inactivation of glycogen synthase kinase-3 by protein kinase C delta: implications for regulation of tau phosphorylation, FEBS Letters 469:111-117 (2000). 186. A. Shanavas, and S. Papasozomenos, tau kinases in the rat heat shock model: possible implications for Alzheimer disease, Proc Natl Acad Sci USA 97:1413914144 (2000). 187. X. Zhu, C. Rottkamp, H. Boux, A. Takeda, G. Perry, and M. Smith, Activation of p38 kinase links tau phosphorylation, oxidative stress, and cell cycle-related events in Alzheimer disease, J Neuropathol Exper Neurol 59:880-888 (2000). 188. M. Bennecib, C. Gong, I. Grundke-Iqbal, and K. Iqbal, Role of protein phosphatase2A and -1 in the regulation of GSK-3, cdk5 and cdc2 and the phosphorylation of tau in rat forebrain, FEBS Lett 485:87-93 (2000). 189. E. Planel, K. Yasutake, S. Fujita, and K. Ishiguro, Inhibition of protein phosphatase 2A overrides tau protein kinase I/glycogen synthase kinase 3 beta and cyclindependent kinase 5 inhibition and results in tau hyperphosphorylation in the hippocampus of starved mouse, J Biol Chem 276:34298-34306 (2001). 190. M. Hoshi, A. Takashima, K. Noguchi, et al., Regulation of mitochondrial pyruvate dehydrogenase activity by tau protein kinase I/glycogen synthase kinase 3beta in brain, Proc Natl Acad Sci USA 93:2719-2723 (1996). 191. S. Kang, T. Kwon, D. Kwon, and S. Do, Akt protein kinase enhances human telomerase activity through phosphorylation of telomerase reverse transcriptase subunit, J Biol Chem 274:13085-13090 (1999). 192. M. Mattson, and W. Klapper, Emerging roles for telomerase in neuronal development and apoptosis, J Neurosci Res 63:1-9 (2001). 193. R. Takano, S. Hisahara, K. Namikawa, H. Kiyama, H. Okano, and M. Miura, Nerve growth factor protects oligodendrocytes from tumor necrosis factor-alpha-induced injury through Akt-mediated signaling mechanisms, J Biol Chem 275:16360-16365 (2000). 194. O. Ozes, L. Mayo, J. Gustin, S. Pfeffer, L. Pfeffer, and D. Donner, NF-kappaB activation by tumour necrosis factor requires the Akt serine-threonine kinase, Nature 401:82-85 (1999). 195. E. Beattie, D. Stellwagen, W. Morishita, et al., Control of synaptic strength by glial TNFalpha, Science 295:2282-2285 (2002). 196. A. Yamaguchi, M. Tamatani, H. Matsuzaki, et al., Akt activation protects hippocampal neurons from apoptosis by inhibiting transcriptional activity of p53, J Biol Chem 276:5256-5264 (2001). 197. D. Cross, A. Culbert, K. Chalmers, L. Facci, S. Skaper, and A. Reith, Selective small-molecule inhibitors of glycogen synthase kinase-3 activity protect primary neurones from death, J Neurochem 77:94-102 (2001). 198. I. Skoog, and D. Gustafson, HRT and dementia, J Epidemiol Biostat 4:227-251 (1999). 199. K. Honda, H. Sawada, T. Kihara, et al., Phosphatidylinositol 3-kinase mediates neuroprotection by estrogen in cultured cortical neurons, J Neurosci Res 60:321-327 (2000). 200. K. Yaffe, L. Lui, D. Grady, J. Cauley, J. Kramer, and S. Cummings, Cognitive decline in women in relation to non-protein-bound oestradiol concentrations, Lancet 356:708-712 (2000). 201. E. LeBlanc, J. Janowsky, B. Chan, and H. Nelson, Hormone replacement therapy and cognition: systematic review and meta-analysis, J Amer Med Assoc 285:14891499 (2001).

342

Chapter 15

202. B. Cholerton, C. E. Gleason, L. D. Baker, and S. Asthana, Estrogen and Alzheimer’s disease: the story so far, Drugs Aging 19:405-27 (2002). 203. C. M. Clark, and J. H. Karlawish, Alzheimer disease: current concepts and emerging diagnostic and therapeutic strategies, Ann Intern Med 138:400-10 (2003). 204. M. Cordey, U. Gundimeda, R. Gopalakrishna, and C. J. Pike, Estrogen activates protein kinase C in neurons: role in neuroprotection, J Neurochem 84:1340-8 (2003). 205. C. Toran-Allerand, M. Singh, and G. Sétáló, Jr, Novel mechanisms of estrogen action in the brain: new players in an old story, Frontiers Neuroendocrinol 20:97-121 (1999). 206. R. Bi, M. Foy, R. Vouimba, R. Thompson, and M. Baudry, Cyclic changes in estradiol regulate synaptic plasticity through the MAP kinase pathway, Proc Natl Acad Sci U S A 98:13391-13395 (2001). 207. D. Kang, S. Soriano, M. Frosch, et al., Presenilin 1 facilitates the constitutive turnover of beta-catenin: differential activity of Alzheimer’s disease-linked PS1 mutants in the beta-catenin-signaling pathway, J Neurosci 19:4229-4237 (1999). 208. W. Stoothoff, C. Bailey, K. Mi, S. Lin, and G. Johnson, Axin negatively affects tau phosphorylation by glycogen synthase kinase 3-beta, Journal of Neurochemistry 83:904-913 (2002). 209. Z. Zhang, H. Hartmann, V. Do, et al., Destabilization of beta-catenin by mutations in presenilin-1 potentiates neuronal apoptosis, Nature 395:698-702 (1998). 210. C. Weihl, G. Ghadge, S. Kennedy, N. Hay, R. Miller, and R. Roos, Mutant presenilin-1 induces apoptosis and downregulates Akt/PKB, J Neurosci 19:53605369 (1999). 211. M. Nishimura, G. Yu, G. Levesque, et al., Presenilin mutations associated with Alzheimer disease cause defective intracellular trafficking of beta-catenin a component of the presenilin protein complex, Nature Med 5:164-169 (1999). 212. B. Passer, L. Pellegrini, P. Vito, J. Ganjei, and L. D’Adamio, Interaction of Alzheimer’s presenilin-1 and presenilin-2 with Bcl-X(L): A potential role in modulating the threshold of cell death, J Biol Chem 274:24007-24013 (1999). 213. J. Tan, T. Town, A. Placzek, A. Kundtz, H. Yu, and M. Mullan, Bcl-X(L) inhibits apoptosis and necrosis produced by Alzheimer’s beta-amyloid 1-40 peptide in PC 12 cells, Neuroscience Letters 272:5-8 (1999). 214. Q. Guo, W. Fu, B. Sopher, et al., Increased vulnerability of hippocampal neurons to excitotoxic necrosis in presenilin-1 mutant knock-in mice, Nature Med 5:101-106 (1999). 215. R. Nuydens, C. Heers, A. Chadarevian, et al., Sodium butyrate induces aberrant tau phosphorylation and programmed cell death in human neuroblastoma cells, Brain Res 688:86-94 (1995). 216. M. Erecinska, D. Nelson, Y. Daikhin, and M. Yudkoff, Regulation of GAB A level in rat brain synaptosomes: fluxes through enzymes of the GABA shunt and effects of glutamate, calcium, and ketone bodies, J Neurochem 67:2325-2334 (1996C). 217. M. Zhang, Y. Gong, and G. Minuk, The effects of ethanol and gamma aminobutyric acid alone and in combination on hepatic regenerative activity in the rat, Journal of Hepatology 29:638-41 (1998). 218. G. Minuk, R. Erlitzki, Y. Gong, and M. Zhang, Identification and characterization of GAB A A receptors and GABA transport proteins in the liver, Hepatology 30:308 A (1999). 219. J.-P. Roperch, V. Alvaro, S. Prieur, et al., Inhibition of presenilin 1 expression is promoted by p53 and p21WAF-1 and results in apoptosis and tumor suppression, Nature Medicine 4:835-838 (1998).


343

220. R. Ren, J. Lah, A. Diehlmann, et al., Differential effects of transforming growth factor-betaS and glial cell line-derived neurotrophic factor on gene expression of presenilin-1 in human post-mitotic neurons and astrocytes, Neurosci 93:1041-1049 (1999). 221. B. Wolozin, K. Iwasaki, P. Vito, et al., Participation of presenilin 2 in apoptosis: enhanced basal activity conferred by an Alzheimer mutation, Science 274:1710-3 (1996). 222. A. Copani, F. Condorelli, A. Caruso, et al., Mitotic signaling by beta-amyloid causes neuronal death, FASEB J 13:2225-2234 (1999). 223. A. Giovanni, E. Keramaris, E. Morris, et al., E2F1 mediates death of B-amyloidtreated cortical neurons in a manner independent of p53 and dependent on Bax and caspase 3, J Biol Chem 275:11553-11560 (2000). 224. Q. Wu, C. Combs, S. Cannady, D. Geldmacher, and K. Herrup, Beta-amyloid activated microglia induce cell cycling and cell death in cultured cortical neurons, Neurobiol Aging 21:797-806 (2000). 225. R. Ockner, Apoptosis and liver diseases: Recent concepts of mechanism and significance, J Gastroenterol Hepatol 16:248-260 (2001). 226. S. Fukumoto, T. Mutoh, T. Hasegawa, et al., GD3 synthase gene expression in PC12 cells results in the continuous activation of TrkA and ERK1/2 and enhanced proliferation, J Biol Chem 275:5832-8 (2000). 227. A. Copani, D. Melchiorri, A. Caricasole, et al., Beta-amyloid-induced synthesis of the ganglioside GD3 is a requisite for cell cycle reactivation and apoptosis in neurons, J Neurosci 22:3963-8 (2002). 228. K. L. Jordan-Sciutto, L. M. Malaiyandi, and R. Bowser, Altered distribution of cell cycle transcriptional regulators during Alzheimer disease, J Neuropathol Exp Neurol 61:358-67 (2002). 229. Y. Yang, E. J. Mufson, and K. Herrup, Neuronal cell death is preceded by cell cycle events at all stages of Alzheimer’s disease, J Neurosci 23:2557-63 (2003). 230. J. J. Pei, H. Braak, C. X. Gong, et al., Up-regulation of cell division cycle (cdc) 2 kinase in neurons with early stage Alzheimer’s disease neurofibrillary degeneration, Acta Neuropathol (Berl) 104:369-76 (2002). 231. J. Dypbukt, M. Ankarcrona, M. Burkitt, et al., Different prooxidant levels stimulate growth, trigger apoptosis, or produce necrosis of insulin-secreting RINm5F cells: The role of intracellular polyamines., J Biol Chem 269:30553-30560 (1994). 232. A. Raina, M. Monteiro, A. McShea, and M. Smith, The role of cell cycle -mediated events in Alzheimer’s disease, Int J Exp Path 80:71-76 (1999). 233. J. Busser, D. Geldmacher, and K. Herrup, Ectopic cell cycle proteins predict the sites of neuronal cell death in Alzheimer’s disease brain, J Neurosci 18:2801-2807 (1998). 234. D. Liu, and L. Greene, Neuronal apoptosis at the G1/S cell cycle checkpoint, Cell Tissue Res 305:217-228 (2001). 235. E. Morris, E. Keramaris, H. Rideout, et al., Cyclin-dependent kinases and P53 pathways are activated independently and mediate Bax activation in neurons after DNA damage, J Neurosci 21:5017-5026 (2001). 236. T. Burton, B. Liang, A. Dibrov, and F. Amara, Transcriptional activation and increase in expression of Alzheimer’s beta-amyloid precursor protein gene is mediated by TGF-beta in normal human astrocytes, Biochem Biophys Res Commun 295:70212 (2002). 237. T. Wyss-Coray, F. Yan, A. H. Lin, et al., Prominent neurodegeneration and increased plaque formation in complement-inhibited Alzheimer’s mice, Proc Natl Acad Sci U S A 99:10837-42 (2002).

344

Chapter 15

238. G. Halliday, S. Robinson, C. Shepherd, and J. Kril, Alzheimer’s disease and inflammation: a review of cellular and therapeutic mechanisms, Clin Exper Pharmacol Physiol 27:1-8 (2000). 239. P. McGeer, Cyclo-oxygenase-2 inhibitors: rationale and therapeutic potential for Alzheimer’s disease, Drugs Aging 17:1-11 (2000). 240. T. Wyss-Coray, and L. Mucke, Ibuprofen, inflammation and Alzheimer disease, Nature Med 6:973-974 (2000). 241. B. in t’ Veld, A. Ruitenberg, A. Hofman, et al., Nonsteroidal antiinflammatory drugs and the risk of Alzheimer’s disease, N Engl J Med 345:1515-1521 (2001). 242. P. Zandi, and J. Breitner, Do NSAIDs prevent Alzheimer’s disease? And, if so, why? The epidemiological evidence, Neurobiol Aging 22:811-817 (2001). 243. S. E. Nilsson, B. Johansson, S. Takkinen, et al., Does aspirin protect against Alzheimer’s dementia? A study in a Swedish population-based sample years, Eur J Clin Pharmacol 59:313-9 (2003). 244. T. Thomas, T. Nadackal, and K. Thomas, Aspirin and non-steroidal antiinflammatory drugs inhibit amyloid-beta aggregation, Neuroreport 12:3263-3267 (2001). 245. S. Weggen, J. Eriksen, P. Das, et al., A subset of NSAIDs lower amyloidogenic Abeta42 independently of cyclooxygenase activity, Nature 414:212-216 (2001). 246. T. Morihara, T. Chu, O. Ubeda, W. Beech, and G. Cole, Selective inhibition of Abeta42 production, Journal of Neurochemistry 83:1009-1012 (2002). 247. J. L. Eriksen, S. A. Sagi, T. E. Smith, et al., NSAIDs and enantiomers of flurbiprofen target gamma-secretase and lower Abeta 42 in vivo, J Clin Invest 112:440-9 (2003). 248. S. A. Sagi, S. Weggen, J. Eriksen, T. E. Golde, and E. H. Koo, The noncyclooxygenase targets of non-steroidal anti-inflammatory drugs, lipoxygenases, peroxisome proliferator-activated receptor, inhibitor of kappa B kinase, and NF kappa B, do not reduce amyloid beta 42 production, J Biol Chem 278:31825-30 (2003). 249. P. S. Aisen, K. A. Schafer, M. Grundman, et al., Effects of rofecoxib or naproxen vs placebo on Alzheimer disease progression: a randomized controlled trial, JAMA 289:2819-26 (2003). 250. J. Yuan, and B. Yankner, Apoptosis in the nervous system, Nature 407:802-809 (2000). 251. P. Klein, and D. Melton, A molecular mechanism for the effect of lithium on development, Proc Natl Acad Sci USA 93:8455-8459 (1996). 252. V. Stambolic, L. Ruel, and J. Woodgett, Lithium inhibits glycogen synthase kinase3 activity and mimics wingless signalling in intact cells, Curr Biol 6:1664-1668 (1996). 253. E. Chalecka-Franaszek, and D. Chuang, Lithium activates the serine/threonine kinase Akt-1 and suppresses glutamate-induced inhibition of Akt-1 activity in neurons, Proc Natl Acad Sci USA 96:8745-8750 (1999). 254. S. Summers, A. Kao, A. Kohn, et al., The role of glycogen synthase kinase 3beta in insulin-stimulated glucose metabolism, J Biol Chem 274:17934-17940 (1999). 255. F. Staal, B. Burgering, M. van de Wetering, and H. Clevers, Tcf-1-mediated transcription in T lymphocytes: differential role for glycogen synthase kinase-3 in fibroblasts and T cells, Intl Immunol 11:317-323 (1999). 256. K. Ptashne, F. Stockdale, and S. Conlon, Initiation of DNA synthesis in mammary epithelium and mammary tumors by lithium ions, J Cellular Physiol 103:41-6 (1980).


345

257. H. Cui, Y. Meng, and R. Bulleit, Inhibition of glycogen synthase kinase 3beta activity regulates proliferation of cultured cerebellar granule cells, Brain Research Devel Brain Res 111:177-188 (1998). 258. G. Moore, J. Bebchuk, I. Wilds, G. Chen, and H. Menji, Lithium-induced increase in human brain grey matter, Lancet 356:1241-1242 (2000). 259. S. Nonaka, C. Hough, and D. Chuang, Chronic lithium treatment robustly protects neurons in the central nervous system against excitotoxicity by inhibiting N-methylD-aspartate receptor-mediated calcium influx, Proc Natl Acad Sci USA 95:2642-2647 (1998). 260. F. Lucas, R. Goold, P. Gordon-Weeks, and P. Salinas, Inhibition of GSK-3beta leading to the loss of phosphorylated MAP-1B is an early event in axonal remodelling induced by WNT-7a or lithium, J Cell Sce 111:1351-1361 (1998). 261. M. Hong, D. Chen, P. Klein, and V. Lee, Lithium reduces tau phosphorylation by inhibition of glycogen synthase kinase-3, J Biol Chem 272:25326-25332 (1997B). 262. I. Blasko, M. Wagner, N. Whitaker, B. Grubeck-Loebenstein, and P. Jansen-Dürr, The amyloid beta peptide abeta (25-35) induces apoptosis independent of p53, FEBS Letters 470:221-225 (2000). 263. E. Paradis, H. Douillard, M. Koutroumanis, C. Goodyer, and A. LeBlanc, Amyloid beta peptide of Alzheimer’s disease downregulates Bcl-2 and upregulates bax expression in human neurons, J Neurosci 16:7533-7539 (1996). 264. G. Alvarez, J. Muñoz-Montaño, J. Satrústegui, J. Avila, E. Bogónez, and J. DíazNido, Lithium protects cultured neurons against beta-amyloid-induced neurodegeneration, FEBS Letters 453:260-264 (1999). 265. H. Wei, P. Leeds, Y. Qian, W. Wei, R.-W. Chen, and D.-M. Chuang, Beta-amyloid peptide-induced death of PC 12 cells and cerebellar granule cell neurons is inhibited by long-term lithium treatment, Eur J Pharmacol 392:117-123 (2000). 266. O. Ghribi, M. M. Herman, and J. Savory, Lithium inhibits Abeta-induced stress in endoplasmic reticulum of rabbit hippocampus but does not prevent oxidative damage and tau phosphorylation, J Neurosci Res 71:853-62 (2003). 267. R. S. Jope, and G. N. Bijur, Mood stabilizers, glycogen synthase kinase-3beta and cell survival, Mol Psychiatry 7 Suppl 1:S35-45 (2002). 268. J. C. Soares, Can brain-imaging studies provide a ‘mood stabilizer signature?’ Mol Psychiatry 7 Suppl 1:S64-70 (2002). 269. S. Washizuka, C. Kakiuchi, K. Mori, et al., Association of mitochondrial complex I subunit gene NDUFV2 at 18p11 with bipolar disorder, Am J Med Genet Part B 120B:72-8 (2003). 270. P. Videbech, B. Ravnkilde, T. H. Pedersen, et al., The Danish PET/depression project: clinical symptoms and cerebral blood flow. A regions-of-interest analysis, Acta Psychiatr Scand 106:35-44 (2002). 271. N. Jha, O. Jurma, G. Lalli, et al., Glutathione depletion in PC12 results in selective inhibition of mitochondrial complex I activity: Implications for Parkinson’s disease, J Biol Chem 275:26096-26101 (2000). 272. K. Conway, J. Rochet, R. Bieganski, and P. Lansbury, Jr, Kinetic stabilization of the alpha-synuclein protofibril by a dopamine-alpha-synuclein adduct, Science 294:1346-1349 (2001). 273. R. Sharon, M. Goldberg, I. Bar-Josef, R. Betensky, J. Shen, and D. Selkoe, alphaSynuclein occurs in lipid-rich high molecular weight complexes, binds fatty acids, and shows homology to the fatty acid-binding proteins, Proc Natl Acad Sci U S A 98:9110-9115 (2001). 274. S. Prusiner, Prions, Proc Natl Acad Sci USA 95:13363-13383 (1998).

346

Chapter 15

275. T. Voigtlander, S. Kloppel, P. Birner, et al., Marked increase of neuronal prion protein immunoreactivity in Alzheimer’s disease and human prion diseases, Acta Neuropathol (Berl) 101:417-423 (2001). 276. S. Mouillet-Richard, M. Ermonval, C. Chebassier, et al., Signal transduction through prion protein, Science 289:1925-1928 (2000). 277. C. Wu, S. Butz, Y. Ying, and R. Anderson, Tyrosine kinase receptors concentrated in caveolae-like domains from neuronal plasma membrane, J Biol Chem 272:35543559 (1997). 278. H. Chapman, Y. Wei, D. Simon, and D. Waltz, Role of urokinase receptor and caveolin in regulation of integrin signaling, Thromb Haemostasis 82:291-297 (1999). 279. X. Sun, S. Pons, T. Asano, M. Myers, Jr, E. Glasheen, and M. White, The Fyn tyrosine kinase binds Irs-1 and forms a distinct signaling complex during insulin stimulation, J Biol Chem 271:10583-10587 (1996). 280. M. Resh, Fyn a Src family tyrosine kinase, Intl J Biochem Cell Biol 30:1159-1162 (1998). 281. S. Grant, T. O’Dell, K. Karl, P. Stein, P. Soriano, and E. Kandel, Impaired long-term potentiation spatial learning and hippocampal development in fyn mutant mice, Science 258:1903-1910 (1992). 282. C. Mastick, and A. Saltiel, Insulin-stimulated tyrosine phosphorylation of caveolin is specific for the differentiated adipocyte phenotype in 3T3-L1 cells, J Biol Chem 272:20706-20714 (1997). 283. H. Beggs, S. Baragona, J. Hemperly, and P. Maness, NCAM140 interacts with the focal adhesion kinase p125(fak) and the SRC-related tyrosine kinase p59(fyn), J Biol Chem 272:8310-8319 (1997). 284. C. O’Donovan, D. Tobin, and T. Cotter, Prion protein fragment PrP-(106-126) induces apoptosis via mitochondrial disruption in human neuronal SH-SY5Y cells, J Biol Chem 276:43516-43523 (2001). 285. M. Perez, A. I. Rojo, F. Wandosell, J. Diaz-Nido, and J. Avila, Prion peptide induces neuronal cell death through a pathway involving glycogen synthase kinase 3, Biochem J 372:129-36 (2003). 286. D. Pratico, Alzheimer’s disease and oxygen radicals: new insights, Biochem Pharmacol 63:563-567 (2002). 287. D. A. Butterfield, and C. B. Pocernich, The glutamatergic system and Alzheimer’s disease: therapeutic implications, CNS Drugs 17:641-52 (2003). 288. R. J. Mark, Z. Pang, J. W. Geddes, K. Uchida, and M. P. Mattson, Amyloid betapeptide impairs glucose transport in hippocampal and cortical neurons: involvement of membrane lipid peroxidation, J Neurosci 17:1046-54 (1997). 289. T. Prapong, J. Buss, W. Hsu, P. Heine, H. West Greenlee, and E. Uemura, Amyloid beta-peptide decreases neuronal glucose uptake despite causing increase in GLUT3 mRNA transcription and GLUT3 translocation to the plasma membrane, Exp Neurol 174:253-258 (2002). 290. E. Uemura, and H. Greenlee, Amyloid beta-peptide inhibits neuronal glucose uptake by preventing exocytosis, Exp Neurol 170:270-276 (2001). 291. I. Kaneko, N. Yamada, Y. Sakuraba, M. Kamenosono, and S. Tutumi, Suppression of mitochondrial succinate dehydrogenase, a primary target of beta-amyloid, and its derivative racemized at Ser residue, J Neurochem 65:2585-2593 (1995). 292. J. N. Keller, Z. Pang, J. W. Geddes, et al., Impairment of glucose and glutamate transport and induction of mitochondrial oxidative stress and dysfunction in


347

synaptosomes by amyloid beta-peptide: role of the lipid peroxidation product 4hydroxynonenal, J Neurochem 69:273-84 (1997). 293. L. Canevari, J. B. Clark, and T. E. Bates, beta-Amyloid fragment 25-35 selectively decreases complex IV activity in isolated mitochondria, FEBS Lett 457:131-4 (1999). 294. C. Rodrigues, S. Sola, M. Brito, C. Brondino, D. Brites, and J. Moura, Amyloid beta-peptide disrupts mitochondrial membrane lipid and protein structure: protective role of tauroursodeoxycholate, Biochem Biophys Res Commun 281:468-474 (2001). 295. F. Bosetti, F. Brizzi, S. Barogi, et al., Cytochrome c oxidase and mitochondrial F(1)F(0)-ATPase (ATP synthase) activities in platelets and brain from patients with Alzheimer’s disease, Neurobiol Aging 23:371-376 (2002). 296. C. Casley, L. Canevari, J. Land, J. Clark, and M. Sharpe, Beta-amyloid inhibits integrated mitochondrial respiration and key enzyme activities, J Neurochem 80:91100 (2002). 297. L. Mucke, E. Masliah, G. Q. Yu, et al., High-level neuronal expression of abeta 1-42 in wild-type human amyloid protein precursor transgenic mice: synaptotoxicity without plaque formation, J Neurosci 20:4050-8 (2000). 298. S. Cardoso, R. Swerdlow, and C. Oliveira, Induction of cytochrome c-mediated apoptosis by amyloid beta 25-35 requires functional mitochondria, Brain Res 931:117-125 (2002). 299. P. Kienlen-Campard, S. Miolet, B. Tasiaux, and J. N. Octave, Intracellular amyloidbeta 1-42, but not extracellular soluble amyloid-beta peptides, induces neuronal apoptosis, J Biol Chem 277:15666-70 (2002). 300. Y. Zhang, R. McLaughlin, C. Goodyer, and A. LeBlanc, Selective cytotoxicity of intracellular amyloid beta peptide1-42 through p53 and Bax in cultured primary human neurons, J Cell Biol 156:519-29 (2002). 301. L. E. Goldstein, J. A. Muffat, R. A. Cherny, et al., Cytosolic beta-amyloid deposition and supranuclear cataracts in lenses from people with Alzheimer’s disease, Lancet 361:1258-65 (2003). 302. R. Kayed, E. Head, J. L. Thompson, et al., Common structure of soluble amyloid oligomers implies common mechanism of pathogenesis, Science 300:486-9 (2003). 303. V. Fonte, V. Kapulkin, A. Taft, A. Fluet, D. Friedman, and C. D. Link, Interaction of intracellular beta amyloid peptide with chaperone proteins, Proc Natl Acad Sci U S A 99:9439-44 (2002). 304. G. Alvarez, M. Ramos, F. Ruiz, J. Satrustegui, and E. Bogonez, Pyruvate protection against beta-amyloid-induced neuronal death: role of mitochondrial redox state, J Neurosci Res 73:260-9 (2003). 305. H. Y. Wang, M. R. D’Andrea, and R. G. Nagele, Cerebellar diffuse amyloid plaques are derived from dendritic Abeta42 accumulations in Purkinje cells, Neurobiol Aging 23:213-23 (2002). 306. G. K. Gouras, J. Tsai, J. Naslund, et al., Intraneuronal Abeta42 accumulation in human brain, Am J Pathol 156:15-20 (2000). 307. L. Xie, E. Helmerhorst, K. Taddei, B. Plewright, W. Van Bronswijk, and R. Martins, Alzheimer’s beta-amyloid peptides compete for insulin binding to the insulin receptor, J Neurosci 22:RC221-5 (2002). 308. M. Harris, Y. Wang, N. Pedigo, Jr, K. Hensley, D. Butterfield, and J. Carney, Amyloid beta peptide (25-35) inhibits Na+-dependent glutamate uptake in rat hippocampal astrocyte cultures, J Neurochem 67:277-286 (1996). 309. Y. Ikegaya, S. Matsuura, S. Ueno, et al., Beta-amyloid enhances glial glutamate uptake activity and attenuates synaptic efficacy, J Biol Chem 277:32180-6 (2002).

348

Chapter 15

310. W. Farris, S. Mansourian, Y. Chang, et al., Insulin-degrading enzyme regulates the levels of insulin, amyloid beta-protein, and the beta-amyloid precursor protein intracellular domain in vivo, Proc Natl Acad Sci U S A 100:4162-7 (2003). 311. B. C. Miller, E. A. Eckman, K. Sambamurti, et al., Amyloid-beta peptide levels in brain are inversely correlated with insulysin activity levels in vivo, Proc Natl Acad Sci U S A 100:6221-6 (2003). 312. D. Gabuzda, J. Busciglio, L. Chen, P. Matsudaira, and B. Yankner, Inhibition of energy metabolism alters the processing of amyloid precursor protein and induces a potentially amyloidogenic derivative, J Biol Chem 269:13623-13628 (1994). 313. W. Meier-Ruge, and C. Bertoni-Freddari, The significance of glucose turnover in the brain in the pathogenetic mechanisms of Alzheimer’s disease, Reviews in the Neurosciences 7:1-19 (1996). 314. L. Zhang, B. Zhao, D. T. Yew, J. W. Kusiak, and G. S. Roth, Processing of Alzheimer’s amyloid precursor protein during H2O2-induced apoptosis in human neuronal cells, Biochem Biophys Res Commun 235:845-8 (1997). 315. F. Ekinci, M. Linsley, and T. Shea, Beta-amyloid-induced calcium influx induces apoptosis in culture by oxidative stress rather than tau phosphorylation, Molec Brain Res 76:389-395 (2000). 316. H. Misonou, M. Morishima-Kawashima, and Y. Ihara, Oxidative stress induces intracellular accumulation of amyloid beta-protein (Abeta) in human neuroblastoma cells, Biochemistry 39:6951-9 (2000). 317. D. Paola, C. Domenicotti, M. Nitti, et al., Oxidative stress induces increase in intracellular amyloid beta-protein production and selective activation of betaI and betaII PKCs in NT2 cells, Biochem Biophys Research Commun 268:642-646 (2000). 318. E. Tamagno, P. Bardini, A. Obbili, et al., Oxidative stress increases expression and activity of BACE in NT2 neurons, Neurobiol Dis 10:279-88 (2002). 319. A. Volterra, D. Trotti, C. Tromba, S. Floridi, Racagni, and G, Glutamate uptake inhibition by oxygen free radicals in rat cortical astrocytes, J Neurosci 14:29242932 (1994). 320. D. Trotti, D. Rossi, O. Gjesdal, et al., Peroxynitrite inhibits glutamate transporter subtypes, J Biol Chem 271:5976-5979 (1996). 321. Y. Chen, W. Ying, V. Simma, et al., Overexpression of Cu,Zn superoxide dismutase attenuates oxidative inhibition of astrocyte glutamate uptake, J Neurochem 75:939945 (2000). 322. N. Bogdanovic, M. Zilmer, K. Zilmer, A. Rehema, and E. Karelson, The Swedish APP670/671 Alzheimer’s disease mutation: the first evidence for strikingly increased oxidative injury in the temporal inferior cortex, Dement Geriatr Cogn Disord 12:364370 (2001 A). 323. Y. Huang, X. Q. Liu, T. Wyss-Coray, W. J. Brecht, D. A. Sanan, and R. W. Mahley, Apolipoprotein E fragments present in Alzheimer’s disease brains induce neurofibrillary tangle-like intracellular inclusions in neurons, Proc Natl Acad Sci U S A 98:8838-43 (2001). 324. A. Dresner, D. Laurent, M. Marcucci, et al., Effects of free fatty acids on glucose transport and IRS-1-associated phosphatidylinositol 3-kinase activity, J Clin Invest 103:253-259 (1999). 325. Z. Jiang, Y. Lin, A. Clemont, et al., Characterization of selective resistance to insulin signaling in the vasculature of obese Zucker (fa/fa) rats, J Clin Invest 104:447-457 (1999).


349

326. K. Cusi, K. Maezono, A. Osman, et al., Insulin resistance differentially affects the PI 3-kinase- and MAP kinase-mediated signaling in human muscle, J Clin Invest 105:311-320 (2000). 327. H. Steinberg, G. Paradisi, G. Hook, K. Crowder, J. Cronin, and A. Baron, Free fatty acid elevation impairs insulin-mediated vasodilation and nitric oxide production, Diabetes 49:1231-1238 (2000). 328. Y. Kruszynska, D. Worrall, J. Ofrecio, J. Frias, G. Macaraeg, and J. Olefsky, Fatty acid-induced insulin resistance: decreased muscle PI3K activation but unchanged Akt phosphorylation, J Clin Endocrinol Metab 87:226-234 (2002). 329. A. Volterra, D. Trotti, P. Cassutti, et al., High sensitivity of glutamate uptake to extracellular free arachidonic acid levels in rat cortical synaptosomes and astrocytes, J Neurochem 59:600-606 (1992). 330. N. Ruderman, P. Ross, M. Berger, Goodman, and MN, Regulation of glucose and ketone-body metabolism in brain of anaesthetized rats, Biochem J 138:1-10 (1974). 331. M. Lovell, W. Ehmann, M. Mattson, Markesbery, and WR, Elevated 4hydroxynonenal in ventricular fluid in Alzheimer’s disease, Neurobiol Aging 18:457461 (1997). 332. L. McGrath, B. McGleenon, S. Brennan, D. McColl, S. McILroy, and A. Passmore, Increased oxidative stress in Alzheimer’s disease as assessed with 4-hydroxynonenal but not malondialdehyde, QJM 94:485-490 (2001). 333. I. Kruman, A. Bruce-Keller, D. Bredesen, G. Waeg, and M. Mattson, Evidence that 4hydroxynonenal mediates oxidative stress-induced neuronal apoptosis, J Neurosci 17:5089-5100 (1997). 334. M. Arai, H. Yamazaki, K. Inoue, and T. Fushiki, Effects of intracranial injection of transforming growth factor-beta relevant to central fatigue on the waking electroencephalogram of rats: Comparison with effects of exercise, Prog NeuroPsychopharmacol Biol Psych 26:307-312 (2002). 335. H. Yamazaki, M. Arai, S. Matsumura, K. Inoue, and T. Fushiki, Intracranial administration of transforming growth factor-beta3 increases fat oxidation in rats, Am J Physiol Endocrinol Metab 283:E536-44 (2002). 336. F. Lezoualc’h, Y. Sagara, F. Holsboer, and C. Behl, High constitutive NF-kappa B activity mediates resistance to oxidative stress in neuronal cells, J Neurosci 18:32243232 (1998). 337. B. Kaltschmidt, M. Uherek, H. Wellmann, B. Volk, and C. Kaltschmidt, Inhibition of NF-kappaB potentiates amyloid beta-mediated neuronal apoptosis, Proc Natl Acad Sci USA 96:9409-14 (1999). 338. A. Beg, and D. Baltimore, An essential role for NK-kappaB in preventing TNFalpha-induced cell death, Science 274:782-784 (1996). 339. D. Van Antwerp, S. Martin, T. Kafri, D. Green, and I. Verma, Suppression of TNFalpha-induced apoptosis by NF-kappaB, Science 274:787-789 (1996). 340. C.-Y. Wang, M. Mayo, and A. Baldwin, Jr, TNF- and cancer therapy-induced apoptosis: potentiation by inhibition of NF-kappaB, Science 274:784-787 (1996). 341. C. Sen, and L. Packer, Antioxidant and redox regulation of gene transcription, FASEB J 10:709-720 (1996). 342. V. Lakshminarayanan, E. Drab-Weiss, and K. Roebuck, H2O2 and tumor necrosis factor-alpha induce differential binding of the redox-responsive transcription factors AP-1 and NF-kappaB to the interleukin-8 promoter in endothelial and epithelial cells, J Biol Chem 273:32670-32678 (1998). 343. E. Shaulian, and M. Karin, AP-1 as a regulator of cell life and death, Nat Cell Biol 4:E131-6 (2002).

350

Chapter 15

344. P. Sanna, M. Cammalleri, F. Berton, et al., Phosphatidylinositol 3-kinase is required for the expression but not for the induction or the maintenance of long-term potentiation in the hippocampal CA1 region, J Neurosci 22:3359-3365 (2002). 345. H. Y. Man, Q. Wang, W. Y. Lu, et al., Activation of PI3-kinase is required for AMPA receptor insertion during LTP of mEPSCs in cultured hippocampal neurons, Neuron 38:611-24 (2003). 346. P. Opazo, A. M. Watabe, S. G. Grant, and T. J. O’Dell, Phosphatidylinositol 3kinase regulates the induction of long-term potentiation through extracellular signalrelated kinase-independent mechanisms, J Neurosci 23:3679-88 (2003). 347. Q. Wang, L. Liu, L. Pei, et al., Control of synaptic strength, a novel function of Akt, Neuron 38:915-28 (2003). 348. L. Zhang, G. Xing, J. Barker, et al., Alpha-lipoic acid protects rat cortical neurons against cell death induced by amyloid and hydrogen peroxide through the Akt signalling pathway, Neurosci Lett 312:125-128 (2001). 349. R. Dodel, Y. Du, K. Bales, F. Gao, and S. Paul, Sodium salicylate and 17betaestradiol attenuate nuclear transcription factor NF-kappaB translocation in cultured rat astroglial cultures following exposure to amyloid A beta(1-40) and lipopolysaccharides, J Neurochem 73:1453-1460 (1999). 350. C. Glabe, Intracellular mechanisms of amyloid accumulation and pathogenesis in Alzheimer’s disease, J Mol Neurosci 17:137-145 (2001). 351. M. Ogawa, H. Fukuyama, Y. Ouchi, H. Yamauchi, and J. Kimura, Altered energy metabolism in Alzheimer’s disease, J Neurol Sci 139:78-82 (1996). 352. M. Bergman, H. Salman, Y. Beloosesky, M. Djaldetti, and H. Bessler, Are peripheral blood cells from patients with Alzheimer disease more sensitive to apoptotic stimuli?, Alzheimer Dis Assoc Disord 16:156-60 (2002). 353. D. Uberti, T. Carsana, E. Bernardi, et al., Selective impairment of p53-mediated cell death in fibroblasts from sporadic Alzheimer’s disease patients, J Cell Sci 115:31318 (2002). 354. C. Cecchi, C. Fiorillo, S. Sorbi, et al., Oxidative stress and reduced antioxidant defenses in peripheral cells from familial Alzheimer’s patients, Free Radical Biol Med 33:1372-1379 (2002). 355. Y. Kitamura, T. Taniguchi, and S. Shimohama, Apoptotic cell death in neurons and glial cells: implications for Alzheimer’s disease, Jpn J Pharmacol 79:1-5 (1999). 356. N. Chattopadhyay, D. Singh, O. Heese, et al., Expression of peroxisome proliferator-activated receptors (PPARs) in human astrocytic cells: PPARgamma agonists as inducers of apoptosis, J Neurosci Res 61:67-74 (2000). 357. M. C. Sugden, K. Bulmer, G. F. Gibbons, B. L. Knight, and M. J. Holness, Peroxisome-proliferator-activated receptor-alpha (PPARalpha) deficiency leads to dysregulation of hepatic lipid and carbohydrate metabolism by fatty acids and insulin, Biochem J 364:361-8 (2002). 358. D. Slosman, C, Ludwig, S. Zerarka, et al., Brain energy metabolism in Alzheimer’s disease: 99mTc-HMPAO SPECT imaging during verbal fluency and role of astrocytes in the cellular mechanism of 99mTc-HMPAO retention, Brain Res Rev 36:230-240 (2001). 359. Neuroinflammation Working Group, Inflammation and Alzheimer’s disease, Neurobiol Aging 21:383-421 (2000). 360. A. A. Farooqui, H. C. Yang, T. A. Rosenberger, and L. A. Horrocks, Phospholipase A2 and its role in brain tissue, J Neurochem 69:889-901 (1997).


351

361. G. Hotamisligil, P. Peraldi, A. Budavari, R. Ellis, M. White, and B. Spiegelman, IRS-1-mediated inhibition of insulin receptor tyrosine kinase activity in TNF-alphaand obesity-induced insulin resistance, Science 271:665-8 (1996A). 362. E. Sternberg, Neural-immune interactions in health and disease, J Clin Invest 100:2641-2647 (1997). 363. N. Rothwell, S. Allan, and S. Toulmond, The role of interleukin 1 in acute neurodegeneration and stroke: pathophysiological and therapeutic implications, J Clin Invest 100:2648-2652 (1997). 364. J. Licinio, and M.-L. Wong, Pathways and mechanisms for cytokine signaling of the central nervous system, J Clin Invest 100:2941-2947 (1997). 365. C. Greenwood, and G. Winocur, Glucose treatment reduces memory deficits in young adult rats fed high-fat diets, Neurobiol Learning Memory 75:179-189 (2001). 366. R. Wing, J. Vazquez, and C. Ryan, Cognitive effects of ketogenic weight-reducing diets, Intl J Obesity Rel Metab Disord 19:811-816 (1995). 367. S. Su, M. Cilio, Y. Sogawa, D. Silveira, G. Holmes, and C. Stafstrom, Timing of ketogenic diet initiation in an experimental epilepsy model, Devel Brain Res 29125:131-138 (2000). 368. X. L. Li, S. Aou, Y. Oomura, N. Hori, K. Fukunaga, and T. Hori, Impairment of longterm potentiation and spatial memory in leptin receptor-deficient rodents, Neuroscience 113:607-15 (2002). 369. N. Z. Gerges, A. M. Aleisa, and K. A. Alkadhi, Impaired long-term potentiation in obese Zucker rats: possible involvement of presynaptic mechanism, Neuroscience 120:535-9 (2003). 370. A. Navarro, M. Sanchez Del Pino, C. Gomez, J. Peralta, and A. Boveris, Behavioral dysfunction, brain oxidative stress, and impaired mitochondrial electron transfer in aging mice, Am J Physiol Regul Integr Comp Physiol 282:R985-R992 (2002). 371. Z. Guo, A. Ersoz, D. Butterfield, and M. Mattson, Beneficial effects of dietary restriction on cerebral cortical synaptic terminals: Preservation of glucose and glutamate transport and mitochondrial function after exposure to amyloid betapeptide, iron, and 3-nitropropionic acid, J Neurochem 75:314-320 (2000). 372. M. Mattson, Emerging neuroprotective strategies for Alzheimer’s disease: dietary restriction telomerase activation and stem cell therapy, Exper Gerontol 35:489-502 (2000). 373. Z. Radák, T. Kaneko, S. Tahara, et al., Regular exercise improves cognitive function and decreases oxidative damage in rat brain, Neurochem Intl 38:17-23 (2001). 374. J. A. Luchsinger, M. X. Tang, S. Shea, and R. Mayeux, Caloric intake and the risk of Alzheimer disease, Arch Neurol 59:1258-63 (2002). 375. D. J. Foley, and L. R. White, Dietary intake of antioxidants and risk of Alzheimer disease: food for thought, Jama 287:3261-3 (2002). 376. H. Jick, G. Zornberg, S. Jick, S. Seshadri, and D. Drachman, Statins and the risk of dementia, Lancet 356:1627-1631 (2000). 377. B. Wolozin, W. Kellman, P. Ruosseau, G. Celesia, and G. Siegel, Decreased prevalence of Alzheimer disease associated with 3-hydroxy-3-methyglutaryl coenzyme A reductase inhibitors, Arch Neurol 57:1439-1443 (2000). 378. K. Fassbender, M. Stroick, T. Bertsch, et al., Effects of statins on human cerebral cholesterol metabolism and secretion of Alzheimer amyloid peptide, Neurology 59:1257-8 (2002). 379. K. Fassbender, M. Simons, C. Bergmann, et al., Simvastatin strongly reduces levels of Alzheimer’s disease beta -amyloid peptides Abeta 42 and Abeta 40 in vitro and in vivo, Proc Natl Acad Sci USA 98:5856-5861 (2001).

352

Chapter 15

380. R. L. Raffai, and K. H. Weisgraber, Cholesterol: from heart attacks to Alzheimer’s disease, J Lipid Res 44:1423-30 (2003). 381. A. Naidu, Q. Xu, R. Catalano, and B. Cordell, Secretion of apolipoprotein E by brain glia requires protein prenylation and is suppressed by statins, Brain Research 958:100-111 (2002). 382. C. Geula, C.-K. Wu, D. Saroff, A. Lorenzo, M. Yuan, and B. Yankner, Aging renders the brain vulnerable to amyloid beta-protein neurotoxicity, Nature Med 4:827-831 (1998). 383. P. Cryer, M. Haymond, J. Santiago, and S. Shah, Norepinephrine and epinephrine release and adrenergic mediation of smoking-associated hemodynamic and metabolic events, New England Journal of Medicine 295:573-7 (1976). 384. M. Hellerstein, N. Benowitz, R. Neese, et al., Effects of cigarette smoking and its cessation on lipid metabolism and energy expenditure in heavy smokers, Journal of Clinical Investigation 93:265-72 (1994). 385. K. Fattinger, D. Verotta, and N. Benowitz, Pharmacodynamics of acute tolerance to multiple nicotinic effects in humans, J Pharmacol Exp Ther 281:1238-46 (1997). 386. J. Rincón, A. Krook, D. Galuska, H. Wallberg-Henriksson, and J. Zierath, Altered skeletal muscle glucose transport and blood lipid levels in habitual cigarette smokers, Clin Physiol 19:135-142 (1999). 387. J. Manson, U. Ajani, S. Liu, D. Nathan, and C. Hennekens, A prospective study of cigarette smoking and the incidence of diabetes mellitus among US male physicians, Am J Med 109:538-542 (2000). 388. L. Fratiglioni, and H. Wang, Smoking and Parkinson’s and Alzheimer’s disease: review of the epidemiological studies, Behaviour Brain Res 113:117-120 (2000). 389. W. Kukull, The association between smoking and Alzheimer’s disease: effects of study design and bias, Biol Psych 49:194-199 (2001). 390. S. L. Tyas, L. R. White, H. Petrovitch, et al., Mid-life smoking and late-life dementia: the Honolulu-Asia Aging Study, Neurobiol Aging 24:589-96 (2003). 391. O. Forlenza, J. Spink, R. Dayanandan, B. Anderton, O. Olesen, and S. Lovestone, Muscarinic agonists reduce tau phosphorylation in non-neuronal cells via GSK-3beta inhibition and in neurons, J Neural Transmission 107:1201-1212 (2000). 392. R. Garrido, A. Malecki, B. Hennig, and M. Toborek, Nicotine attenuates arachidonic acid-induced neurotoxicity in cultured spinal cord neurons, Brain Res 861:59-68 (2000). 393. T. Kihara, S. Shimohama, H. Sawada, et al., alpha 7 nicotinic receptor transduces signals to phosphatidylinositol 3-kinase to block A beta-amyloid-induced neurotoxicity, J Biol Chem 276:13541-13546 (2001). 394. K. West, J. Brognard, A. Clark, et al., Rapid Akt activation by nicotine and a tobacco carcinogen modulates the phenotype of normal human airway epithelial cells, Journal of Clinical Investigation 111:81-90 (2003). 395. S. Fujii, and K. Sumikawa, Acute and chronic nicotine exposure reverse age-related declines in the induction of long-term potentiation in the rat hippocampus, Brain Res 894:347-353 (2001). 396. G. Jones, B. Sahakian, R. Levy, D. Warburton, and J. Gray, Effects of acute subcutaneous nicotine on attention, information processing and short-term memory in Alzheimer’s disease, Psychopharmacol 108:485-494 (1992). 397. K. Ono, K. Hasegawa, M. Yamada, and H. Naiki, Nicotine breaks down preformed Alzheimer’s beta-amyloid fibrils in vitro, Biol Psychiatry 52:880-6 (2002).


353

398. Z. Z. Guan, W. F. Yu, K. R. Shan, T. Nordman, J. Olsson, and A. Nordberg, Loss of nicotinic receptors induced by beta-amyloid peptides in PC 12 cells: possible mechanism involving lipid peroxidation, J Neurosci Res 71:397-406 (2003). 399. D. B. Freir, and C. E. Herron, Nicotine enhances the depressive actions of A beta 140 on long-term potentiation in the rat hippocampal CA1 region in vivo, J Neurophysiol 89:2917-22 (2003). 400. G. Watson, and S. Craft, The role of insulin resistance in the pathogenesis of Alzheimer’s disease. Implications for treatment, CNS Drugs 17:27-45 (2003). 401. A. M. Palmer, Pharmacotherapy for Alzheimer’s disease: progress and prospects, Trends Pharmacol Sci 23:426-33 (2002). 402. R. Wilson, C. Mendes De Leon, L. Barnes, et al., Participation in cognitively stimulating activities and risk of incident Alzheimer disease, JAMA 287:742-748 (2002).

Chapter 16 Part III Conclusions

In this analysis, aspects of normal and abnormal brain function have been examined in the context of neuronal substrate utilization and energy metabolism, emphasizing: a) availability of alternative and competing substrates and their differing effects on mitochondrial function, ATP production, and generation of oxidative stress; b) interaction between neurons and astrocytes as a major determinant of neuronal substrate utilization and redox balance; and c) systemic factors that influence neuronal substrate utilization in health and disease. Based on the available evidence, the following concepts emerge: 1. Complete oxidation of glucose through the glycolytic and citric acid cycles is optimal for energizing neuronal activation, reflecting several important factors: a) compared with other substrates glucose provides the highest yield of ATP per carbon atom and (except for glutamine/glutamate-derived 2-oxo-glutarate) per oxygen molecule consumed; b) entry of glucose-derived carbon (as acetyl CoA) into the citric acid cycle is tightly regulated by pyruvate dehydrogenase, thus minimizing electron transport chain overload and generation of oxidative stress; c) glucose and its metabolic derivatives at physiological concentrations do not uncouple or otherwise compromise the efficiency of oxidative phosphorylation; d) wg/Wnt-like signaling activated by insulin and other growth factors supports optimal glucose utilization, and stimulation of wg/Wnt-like signaling by lithium ion appears to account for this small univalent cation’s surprisingly diverse effects. In contrast to glucose, fatty acids are not utilized by most neurons as energy substrate. However, hypothalamic neurons involved in neuroendocrine regulation, metabolism, and feeding behavior appear to represent an important exception; in this vitally important neuronal subpopulation, 355

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leptin-induced mitochondrial fatty acid oxidation may be a key regulatory determinant. 2. Astrocytes are highly versatile metabolically, and critically important to neuronal substrate supply. Similar to hepatocytes and unlike most neurons, their of fatty acids may generate ROS while supporting ketogenesis and/or gluconeogenesis. Astrocytes do not export glucose; although they export lactate, neuronal utilization of this substrate alone may not be adequate to support normal neuronal activation and synaptic function. Ketone bodies, which are generated via mitochondrial fatty acid oxidation in hepatocytes or locally in astrocytes, are utilized by neurons in preference to glucose under all conditions. However, ketone bodies compromise efficiency of neuronal energy generation by suppressing glucose utilization, by their largely unregulated mitochondrial oxidation with augmentation of matrix reducing potential and ROS generation, and by a non-antioxidant uncoupling-like effect on mitochondrial energetics. Acting together with diffusible oxidants originating in the astrocyte, these phenomena would not only subject the neuron to oxidative stress but in addition compromise normal neuroprotective astrocytic antioxidant defense mechanisms. Glutamine exported by astrocytes is converted in neurons to glutamate, for utilization in neurotransmission and/or energy metabolism. 3. Neuronal activation requires a rapid and substantial increase in the rate of ATP generation. While it is associated with increased regional blood flow and glucose utilization, oxygen consumption remains relatively constant. This apparent paradox may reflect an activationinduced shift to more efficient utilization of oxygen, implying increased use of energetically more efficient substrate, e.g., glucose or glutaminederived 2-oxo-glutarate. Conversely, it is postulated that the resting, nonactivated, neuron is fueled to a relatively greater extent by the energetically less efficient ketone bodies. Inefficiency of ketone bodies as fuel, and their suppression of neuronal glucose utilization and glucosefueled citric acid cycle activity, may contribute to control of seizure activity by the ketogenic diet. 4. Increased neuronal utilization of glutamate as an alternative energy substrate may contribute to long-term potentiation (LTP) and to the pathogenesis of excitotoxicity. In both instances, evidence is consistent with fueling of the postsynaptic neuronal citric acid cycle and electron transport chain by 2-oxo-glutarate, derived from synaptic cleft glutamate. It is hypothesized that in LTP, heightened glutamate exposure and uptake is physiological and repetitive; it thus elicits an adaptive response, i.e., postsynaptic neuronal expression of the AMPA-activated glutamate

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receptor. Excitotoxicity, in contrast, reflects acute exposure of the postsynaptic neuron to excessive concentrations of glutamate; under these conditions, adaptive responses are precluded and neuronal oxidative stress and death ensue. 5. A hypothetical model for the pathogenesis of late-onset Alzheimer disease is presented. The model holds that the functional and structural changes associated with this disorder reflect relatively subtle but sustained generation of neuronal intramitochondrial oxidative stress, which interacts with amyloid in a mutually reinforcing “vicious cycle”. Augmented oxidative stress is hypothesized to reflect neuronal exposure to ketone bodies and ROS, excessive astrocyte generation of which is driven by chronic increases in plasma [FFA] as may occur in association with obesity, insulin resistance, and type II diabetes mellitus. Sustained preferential neuronal ketone body utilization further affects neuronal energetics and survival by suppressing glucose utilization and attenuating wg/Wnt-like signaling. The proposed model integrates these pathogenetic factors with genetic determinants such as apolipoprotein E4 and mutations in presenilin and APP genes; it also provides a mechanistic basis for understanding the apparent neuroprotective effects of diverse pharmacological agents. Similar mechanisms may contribute to the pathogenesis of HIV-associated dementia, in which significant alterations in systemic fatty acid metabolism are well-documented effects of proinflammatory cytokines. The foregoing concepts are plausible and consistent with currently available evidence in this complex and rapidly evolving field; their possible validation will require experimental testing. Particular attention is warranted to: 1) regional quantification and characterization of plasma FFA utilization and ketogenesis in brain; 2) characterization of the potential role of glutamate-induced increases in electron transport chain flux in the generation of long-term potentiation and excitotoxicity; 3) elucidation of the relationship of normal and disease-related neuronal function to astrocyte fatty acid metabolism and ketogenesis, and to substrate-induced intramitochondrial oxidative stress in both cell types; and 4) identification of the interactions among mitochondrial fatty acid oxidation, leptin signaling, and feeding behavior, in hypothalamic neurons involved in metabolic regulation. Aspects of this review that address pathogenesis have focussed largely on Alzheimer disease. However, evidence suggests that the mechanisms considered may be relevant to other chronic neurodegenerative processes as well as to diverse conditions that may also be linked to changes in brain metabolism, including depressive disorders, use and effects of addictive drugs, and the pathogenesis of neoplastic brain disease.

PART IV: EPILOGUE

Chapter 17 Looking Back‚ Looking Ahead

These final paragraphs are intended to provide additional perspective on the concepts developed in the preceding chapters‚ the objectives of which have been: 1) to identify and characterize the integration of cell metabolism and energetics with signal transduction as a central determinant of cell growth‚ injury‚ and death; and 2) to critically analyze the impact of this interaction in the broader context of human health and disease. Do the conclusions and hypotheses generated by this endeavor have merit? Examination of the available evidence suggests for many of them a tentative “yes”. Although their rigorous assessment will require considerable effort‚ the analysis has identified certain basic themes‚ repeated emergence of which in a variety of settings suggests that they may not only prove to be valid‚ but that they may be major determinants in diverse patterns of normal and pathological response. Thus‚ certain sudden‚ unpredictable‚ and survival-critical events are identified in which large increases in chemical energy are required with maximal speed and efficiency. Among these are cell cycle progression‚ neuronal activation‚ epithelial restitution‚ and hypoxia-induced angiogenesis. These events almost always depend upon glycolysis and wg/Wnt-like signaling‚ and require that mitochondrial fatty acid oxidation be suppressed. In contrast‚ more predictable and sustained processes such as maintenance of posture‚ long-distance migratory flight‚ or myocardial contraction‚ are readily fueled by mitochondrial fatty acid oxidation‚ reflecting for these processes the apparent selection of efficiency in energy storage in preference to efficiency in energy generation. Furthermore‚ significant differences in metabolism and mitochondrial effects among classes of fatty acids — e.g.‚ short chain vs. long chain‚ or vs. — are closely linked to their similarly contrasting effects on cell proliferation. 361

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Despite the importance of fatty acids in energizing certain vital processes‚ however‚ their utilization clearly may represent a “double-edged sword”. This dichotomy reflects the ability of cellular fatty acids in excess abundance‚ e.g.‚ as a result of elevated plasma [FFA]‚ to override the controls that normally govern their mitochondrial oxidation. Under such conditions‚ corresponding increases in electron transport chain flux may promote electron “leakage” with one-electron reduction of molecular oxygen‚ forming superoxide anion and derivative reactive oxygen species (ROS). ROS may also be produced by peroxisomal especially in the case of “trans” fatty acids. If antioxidant defense capabilities are exceeded‚ genomic instability and other forms of cell injury may ensue. These adverse consequences of excessive fatty acid oxidation and ROS generation are potentially ubiquitous within the organism. Accordingly‚ they may be implicated and perhaps dominant in the pathogenesis of diverse and widely distributed injurious processes‚ among them aging‚ insulin resistance and type II diabetes mellitus‚ atherosclerosis‚ neurodegeneration‚ and carcinogenesis. Moreover‚ they may facilitate and/or augment adverse effects of exogenous toxic or infectious agents. On this basis‚ identification of the opposing forces of endogenous “good” (antioxidant defenses) and “evil” (ROS) as determinants of the organism’s normal and abnormal function would appear to be relatively straightforward. Unfortunately‚ so simplistic a view is clearly untenable. Thus‚ although ROS are potentially injurious‚ they may in certain settings initiate survivalsupporting responses‚ e.g.‚ activation of transcription factors and AP1. Moreover‚ hypoxia intensifies electron transport chain generation of ROS; ROS‚ in turn‚ inhibit certain protein phosphatases‚ thereby augmenting PI3K and wg/Wnt-like signaling. This sequence contributes to activation and its transcriptional regulation of genes that encode mediators of protective responses‚ including increased expression of VEGF‚ erythropoietin‚ and glycolytic enzymes. Finally‚ ROS generation during a brief‚ “pre-conditioning” period of ischemia activates other protective responses‚ e.g.‚ expression of heat shock and uncoupling proteins‚ that may contribute to survival during subsequent exposure to otherwise lethal ischemia-reperfusion. These multifaceted‚ contrasting‚ concentrationdependent and time-dependent impacts of ROS on cell function and survival are demonstrated perhaps most simply and dramatically by the response of certain cell types to at low concentrations‚ cell proliferation; at moderate concentrations‚ programmed death; and at high concentrations‚ necrosis. Should these seemingly contradictory effects of ROS be viewed as representing the irrational manifestations of a biochemical “Jekyll and

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Hyde”‚ a surviving maladjustment to atmospheric oxygenation a billion-anda-half years ago that somehow escaped deletion at the hands of natural selection? To the contrary‚ available evidence can be interpreted as suggesting that‚ within limits‚ generation of ROS by the mitochondrial electron transport chain may serve as an essential and critical indicator of cellular energetics‚ and perhaps what originally may have been a primitive metabolic proliferation “signal”. Such a signal would have indicated that cell cycle progression is both temporally feasible and energetically optimal. While admittedly speculative‚ this suggestion is supported by the circumstances that pertain at peak mitochondrial energization. Under these conditions‚ and [ATP] are maximal; little ADP is available to drive the ATP synthase reaction (oxidative phsophorylation)‚ the velocity of which determines the rate at which electrons are removed from the electron transport chain and oxygen is consumed. Thus‚ limited ADP supply constrains the rate at which electrons are utilized in the biosynthesis of ATP; as a result‚ they accumulate in the electron transport chain‚ increasing their propensity to “leak” and thereby to generate ROS. Similar to the sequence that has been described in hypoxia‚ ROS activation of would lead to transcriptional regulation of genes important in antioxidant defense‚ while ROS inhibition of certain protein phosphatases would augment PI3K activity and wg/Wnt-like signaling‚ leading to cell cycle progression. Moreover‚ in addition to generating low levels of ROS‚ maximal [ATP] and minimal [ADP] would assure that [AMP] is also minimal and therefore that AMPK is inactive. As a result of AMPK silencing‚ mitochondrial fatty acid oxidation would be suppressed by both malonyl CoA-dependent and malonyl CoAindependent mechanisms‚ thereby promoting biosynthesis of fatty acids and cholesterol as required for membrane biogenesis in mitosis. Thus‚ an energetic/metabolic “code” that drives (or is permissive for) cell cycle progression might comprise a combination of “signal settings”‚ among them: 1) high; 2) [ATP]: high; 3) [AMP]: very low; 4) [ATP]/[AMP]: very high; 5) AMPK: inactive; 6) ROS: moderately increased; 7) PI3K‚ and wg/Wnt-like signaling: active; and 8) lipogenesis: active. Their net effect would be subject to variable modulation by (and to some extent reflect) DNA integrity‚ growth factor signaling‚ extracellular matrix interactions‚ and other potential determinants. Accordingly‚ “good” ROS might be those that are generated through a normal ongoing process (e.g.‚ as the low-level by-product of normal electron transport chain flux)‚ at a rate that can be contained by cellular antioxidant defenses. If such ROS generation were to become slightly and/or transiently increased‚ it would likely initiate survival-promoting cellular signaling. “Evil” ROS‚ in contrast‚ would be those that are generated in greater abundance over longer periods of time (perhaps even years or decades)‚ e.g.‚

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as the direct or indirect of xenobiotics‚ and/or in locations in which antioxidant defenses are weak. Such criteria would satisfy the requirements of diverse circumstances in which ROS are strongly implicated in disease pathogenesis and aging. One example of this would appear to be the consequences of exposure of vascular endothelium to large quantities of lipoprotein lipid and FFA. Another‚ excessive astrocyte fatty acid oxidation driven by sustained high plasma [FFA]‚ resulting in export of ketone bodies and diffusible ROS. Uptake and utilization of these astrocyte-generated derivatives of fatty acid oxidation by the weakly antioxidant-defended brain neuron would suppress neuronal glucose utilization and predispose to oxidative stress-induced neuronal injury. Although the obviously important role of humoral and paracrine mediators themselves has figured less prominently in the foregoing paragraphs‚ their very entry into the extracellular milieu‚ through which they access cognate receptors on cell surfaces‚ is also linked to such considerations. For example‚ insulin secretion is activated by conditions within the pancreatic beta cell that closely resemble those that would support cell cycle progression. Thus‚ the prime determinant is beta cell glycolysis activated by an unpredictable rise in blood glucose concentration: ATP generation is high‚ wg/Wnt-like signaling is permissive or active‚ and fatty acid oxidation is suppressed. Conversely‚ excessive beta cell mitochondrial fatty acid oxidation and ROS generation‚ sustained over years‚ may eventually impair the augmented insulin response that is required to promote glucose utilization in fatty acid-associated insulin resistant extrapancreatic tissues‚ eventually leading to beta cell death and type II diabetes mellitus. Consistent with this concept‚ stimulation of insulin secretion by some oral hypoglycemic agents may reflect their inhibition of beta cell CPT-I. In a quite different and far more transient setting‚ macrophage (Kupffer cell) secretion of early after partial hepatectomy reflects transcriptional regulation of expression by the latter activated at least in part by mitochondrial fatty acid oxidation-induced ROS generation. In hepatocyte as well as Kupffer cell‚ predominant fatty acid oxidation precludes cell proliferation during this several hour long initial phase. Through its hepatocyte receptor‚ however‚ activates the PI3K- and responses that are essential to the normal ensuing hepatocyte proliferation and liver regeneration. Thus‚ metabolic determinants appear not only to energize cell cycle progression and other intracellular processes‚ but in addition contribute importantly to the regulated secretion of intercellular mediators of signals that both communicate and promote circumstances conducive to the activation of such processes.


365

The concepts advanced in this volume also may provide a basis for newer understandings of human disease‚ and its prevention and treatment. Among the questions and hypotheses so generated are the following: Do sustained increases in plasma [FFA] associated with obesity‚ insulin resistance‚ and type II diabetes mellitus‚ or HIV infection — the latter reflecting proinflammatory cytokine effects on adipocyte lipolysis — contribute to neurodegeneration (Alzheimer disease or HIV encephalopathy‚ respectively) by driving increased astrocyte mitochondrial fatty acid oxidation and ketogenesis? Might similar mechanisms account for diabetes mellitusrelated effects on the kidney‚ or on fetal development‚ incidence of preeclampsia‚ and placental insufficiency? Does heightened ROS generation in diverse organs and tissues caused by sustained‚ nicotine-induced increases in plasma [FFA] contribute to the pathogenesis of various smoking-related disorders‚ neoplastic and otherwise? Quite unexpectedly‚ this endeavor has also brought to light hints at what might represent a basis for understanding aspects of an interaction between conscious thought and cellular function. For example‚ — smallest of all charged atomic particles except for the proton — augments wg/Wnt-like signaling and‚ in certain circumstances‚ subjective mood. These parallel effects suggest the possibility that affect may be related to metabolism and energetics in specific brain neurons. Also‚ regulation of the sensation of hunger and its associated behavior patterns may be linked to the combined effects of insulin and leptin on intermediary metabolism in specific hypothalamic neuronal populations‚ which also appears to influence the activity of systemic adrenergic neurotransmission. Possibly related in this context are the differing metabolic and behavioral reponses to activation of cannabinoid receptors by endocannabinoids‚ as compared with tetrahydrocannabinol. Whether such implied and hypothetical “mind-body” linkages are valid remains to be determined‚ as does their possibly broader significance for human thought and feeling. What about cancer? Did the mechanisms that enabled the survival and evolution of our primordial‚ “sea”-dwelling ancestral eukaryotes (Chapter 1) also endow their distant progeny with the genomic plasticity that underlies development of many or most human cancers‚ especially those that do not result from germline mutations? In a setting imposed by human disease‚ cells struggle for survival in a battle zone‚ subjected to potentially lethal injury by immunocytes‚ cytokines‚ toxic xenobiotics or biotransformation products‚ and reactive oxygen species‚ or through deprivation of nutrient‚ oxygen‚ or growth factors. In this environment‚ cell death is expected‚ programmed in part by an exogenous toxin or invading organism‚ or by the host in its defensive response. Under such conditions‚ perhaps the only hope

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for survival may lie in a progressively radical metamorphosis that offers the possibility of reversion to that earlier‚ free-living‚ primordial existence. In this metamorphosis‚ involving incremental modifications in gene expression‚ the cell would discard the trappings of life in its multicellular‚ interdependent‚ and regulated environment. Such a scenario might be exemplified by the following positive or negative changes: Cell-cell (adherens junctions) and cell-matrix associations are severed (wg/Wnt-like signaling and matrilysin expression‚ respectively). Antiproliferative and pro-apoptotic influences are suppressed‚ whether internal (PTEN‚ p53) or external signaling). Energizing nutient is acquired‚ hoarded‚ and utilized aggressively (GLUT 1‚ glucose-6-phosphatase‚ hexokinase II‚ pyruvate kinase). Signaling pathways such as wg/Wnt-like (PI3K/Akt/PKB) integration of fueling‚ energetics‚ and mitosis may be constitutively activated (APC and PTEN mutations)‚ and these in turn may effect coordinate upregulation of biosynthetic pathways for fatty acids‚ isoprenoids‚ and sterols (SREBPs). Oxygen availability is maximized (VEGF and other angiogenic factors); and competition from neighboring cells is counteracted secretion). Among the few antiproliferative mechanisms retained by the transformed malignant cell are those required for survival‚ such as the Cip/Kip protein family. Uniquely among inhibitors of the early cell cycle‚ the Cip/Kip proteins escape inactivating mutation in virtually all human cancers. The malignant cell thus retains access to the vital services of Cip/Kip: 1) activation of D cyclin/cdk4‚6 complexes and of G1 cell cycle progression; and 2) prevention of premature and potentially fatal entry into S phase‚ with its sustained and intensive commitment of energy (i.e.‚ consumption of ATP) to the demands of genome replication. Fully developed‚ the metamorphosis launches the cell upon a solo journey. Recapitulating its ancestral origins‚ the cell now enjoys the possibility of unfettered proliferation‚ limited only by those constraints that are imposed by nutrient and oxygen availability or adverse external factors. Clonal expansion can be accomplished far more rapidly than would have been imaginable under the cumbersome “old” rules that governed regulated growth in a multicellular organism. In fact‚ as clonal expansion progresses‚ the host may be subject to type II diabetes mellitus-like changes in metabolism and a wasting process‚ both of which effectively and preferentially but inefficiently divert nutrient to the growing tumor. In an anthropomorphic extension of its metamorphosis‚ the cell that might be viewed by its progeny as an heroic founding ancestor is seen by us as the arch-enemy. Importantly‚ the transitions that make this new existence possible do not appear to have been dictated by any specific pre-ordained strategy. Rather‚ as was true of its ancestors in the primordial “sea”‚ the transitions that endure are those‚ encoded by more or less random gene


367

mutations (or epigenetic regulation of wild-type genes) that enable the cell to succeed‚ i.e.‚ to survive‚ proliferate‚ and “write the history” of its period. In this setting‚ unfortunately‚ the history is that of an often intractable and devastating illness. But perhaps the growth advantage acquired by the cancer cell in its Darwinian struggle for survival also provides a point of vulnerability (an “Achilles heel”) for those who would seek to defeat it. Although fatty acids‚ salicylate‚ and statins‚ all acting through different mechanisms‚ oppose tumor development and growth by suppressing cell proliferation‚ a more decisive blow is required to deal with the established malignancy. One potential target at which such a blow might be directed is the prolonged and energy-intensive S (DNA synthesis) phase of the cell cycle. During this critical period of genome duplication‚ cellular demand for energy (ATP) is high and sustained‚ and the respite that might otherwise be afforded by cell cycle checkpoint arrest is not available as an option until S phase is completed. Might the implicit unacceptability of compromised energy generation during this period of genome replication represent the basis for a novel therapeutic approach to cancer? In such an approach‚ might it be possible‚ in vivo‚ to achieve near synchronous‚ pharmacologically-induced tumor cell G1 arrest‚ then to activate near synchronous cell cycle progression? If so‚ perhaps a subsequent pharmacological disabling of mitochondrial energetics (e.g.‚ through inhibition of a key metabolic pathway‚ uncoupling of oxidative phosphorylation‚ and/or inhibition of the electron transport chain)‚ and timed for delivery during S phase‚ would trigger a consumption-driven‚ inexorable and precipitous fall in [ATP] and its sequelae: adenylate kinase-mediated rise in [AMP]‚ activation of AMPK‚ and programmed cell death. Because some cancer cells may transiently fuel proliferation anaerobically‚ interference with glycolysis may also be important to augment cell killing. Unavoidably‚ normal‚ non-transformed cells might also be exposed to the reagents employed in such a program. However‚ because these cells remain subject to physiological growth controls‚ they would be far less susceptible to pharmacologically induced cell cycle progression‚ and therefore less vulnerable to the subsequent disruptive manipulation of mitochondrial energetics. While frankly hypothetical — and certainly challenging in regard to its potential for clinical application — the theoretical basis for such an approach appears to be sound. The present treatise has generated many concepts that address important and unresolved scientific and clinical questions. Some of these concepts appear to be strongly supported by the published findings of many scientists‚ variously reinterpreted in light of the hypotheses here advanced. Other concepts are consistent with published reports‚ but need to be specifically

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addressed. Still others are hypothetical or even speculative. For few of the concepts and hypotheses if any‚ however‚ would currently available information justify outright dismissal. Rather‚ their validity or lack thereof will be determined only through direct experimentation. In that sense‚ perhaps the volume itself can be viewed as an experiment‚ hopefully one that will have generated not only questions‚ but also answers that are “fit” enough to survive.

INDEX

Acetoacetate: see Ketone bodies Acetyl CoA carboxylase; see also AMPK; and Fatty acid biosynthesis AMPK‚ 4‚ 5‚ 55‚ 57‚ 79‚ 96 BRCA1,55 fatty acid oxidation‚ 232‚ 237; see also Fatty acid oxidation “statin” agents‚ SREBP‚ 153‚ 154‚ 328 Activator protein-1 (AP-1): see Nuclear factor Adenine nucleotide translocase (ANT): see Mitochondrial energetics‚ VDAC Adenomatous polyposis coli (APC)‚ 27‚ 30‚ 31‚ 33‚ 84-85‚ 97‚ 127‚ 315-316‚ 318‚ 366; see also Cancer colon cancer‚ 27‚ 30‚ 33 glycogen synthase see PI3K/Akt/PKB murine model‚ ‚ 84‚ 127 85 wg/Wnt-like signaling: see PI3K/Akt/PKB signaling Adenosine diphosphate (ADP)‚ ROS generation and‚ 147‚ 158‚ 285‚ 290-291‚ 363 Adenosine monophosphate (AMP): see AMP-activated protein kinase Adenosine triphosphate (ATP) generation‚ stoichiometry‚ 58-60; see also Cell cycle Adenylate kinase and AMPK‚ 94‚ 96‚ 367 Adherens junctions‚ 92‚ 306‚ 316‚ 366 Adipocyte fatty acid binding protein (aP2): see Fatty acid binding proteins Adipose tissue; see also Diabetes mellitus; and Insulin resistance; and Obesity visceral‚ 148‚ 179 Advanced glycation endproducts (AGE)‚ 147-148‚ 151; see also Hyperglycemia Affect: see Depression Aging‚ 8‚ 15‚ 82‚ 149‚ 158‚ 303‚ 307‚ 308‚ 314‚ 328‚ 362‚ 364 Agouti-related peptide‚ 236 Akt-protein kinase B (Akt/PKB): see PI3K/Akt/PKB signaling Alternative substrates‚ 124‚ 217‚ 260‚ 313-314 Alzheimer disease‚ 6‚ 8‚ 9‚ 32-33‚ 155‚ 208‚ 225‚ 267‚277‚280‚ 303-330‚ 365; see also Astrocyte; and Nicotine; and Synaptic function amyloid‚ 33‚ 34‚ 152‚ 208‚ 223‚ 225‚ 240‚ 291‚ 304-306‚ 314-323‚ 328-329 cell cycle induction‚ 319

369

370

Index

Alzheimer disease (cont.) amyloid precursor protein (APP)‚ 306‚ 316‚ 317‚ 320‚ 323‚ 329 Swedish mutant APP‚ 306‚ 323 apolipoprotein (a)‚ 306 apolipoprotein E (apoE)‚ 305-306‚ 328 amyloid and‚ 306‚ 320‚ 323 E2‚ E3 isoforms‚ 305 E4 isoform‚ 305-306‚ 308‚ 322-324 effects on neuronal repair‚ synaptic function‚ 305 cerebral metabolism‚ 309-310 citric acid cycle‚ 305‚ 310‚ 311‚ 314‚ 316‚ 317‚ 319‚ 324 glucose-oxygen dissociation‚ 314; see also Neuronal activation GLUT 3 adducts‚ 309‚ 324 insulin effects‚ 309-310 ketone body utilization and effects‚ 303‚ 309-314‚ 324‚ 326‚ 327‚ 329‚ 364 2-oxo-glutarate dehydrogenase‚ 305‚ 310 regional differences‚ 310 cerebrospinal fluid in diagnosis‚ 304‚ 308 cholesterol cholesterol-degrading enzyme‚ 305 risk factor‚ 309 “statins”‚ 15‚ 55‚ 147‚ 152-154‚ 328‚ 330‚ 367 synaptic function‚ 305 diabetes mellitus‚ insulin resistance‚ 307-309 analogous retinal pathology‚ 308 fatty acid metabolism‚ systemic manifestations‚ potential modulators‚ 327-330 nicotine‚ 322‚ 328-329 insulin-degrading enzyme‚ 306‚ 322‚ 323 neurofibrillary tangles‚ 32-33‚ 304‚ 306‚ 317‚ 318‚ 322‚ 324‚ 329 PI3K/Akt/PKB‚ 304‚ 314-325‚ 328; see also PI3K/Akt/PKB signaling and‚ 314-324 effects and depression‚ 320-321 neurofibrillary tangles‚ 32-33‚ 304‚ 306‚ 317‚ 318‚ 322‚ 324‚ 329 neuronal death and the cell cycle‚ 319-320 presenilins (PS)‚ 31-32‚ 306‚ 315-319‚ 324‚ 329 prion protein interactions‚ 321 phosphorylation‚ 317 “truncated” signaling‚ 316 ROS‚ oxidative stress: see Reactive oxygen species “vicious cycle” in pathogenesis‚ 321-326 5-Aminoimidazole-4-carboxamide ribonucleoside (AICAR): see AMP-activated protein kinase receptor (AMPAR): see Synaptic transmission AMP-activated protein kinase (AMPK)‚ 4-5‚ 54‚ 55‚ 57-59‚ 79‚ 94‚ 96‚ 97‚151‚ 179‚ 189‚ 264‚ 363‚ 367 adenylate kinase‚ 94‚ 96‚ 367 AICAR‚ 96-97 apoptosis and‚ 96-97

Index AMP-activated protein kinase (AMPK)‚ (cont.) fatty acid oxidation‚ 96-97‚ 151‚ 264 regulation of growth‚ lipogenesis‚ 4-5 Anandamide: see Cannabinoids Angiogenesis: see Hypoxia Apolipoprotein (a)‚ 306; see also Alzheimer disease Apolipoprotein B‚ 150 Apoptosis‚ 85-98‚ facing p. 85 Bcl-2 protein family‚ 88-90 caspase(s)‚ 28‚ 34‚ 77‚ facing p. 85‚ 86‚ 87‚ 90‚ 93-96‚ 98‚ 125‚ 126‚ 144‚ 188‚ 239 caspase 3 targets Akt/PKB‚ Bcl-2‚ cytokeratins 8/18‚ 95 cell cycle and‚ 95-98 AICAR‚ 96-98 AMPK‚ 96-98 carcinogenesis‚ 97 hepatitis B and C‚ 97 insulin‚ leptin‚ 97 ceramide‚ 92-94‚ 144‚ 188‚ 239‚ 292 ganglioside GD3‚ 92 mitochondrial function‚ 77-98‚ facing p. 85 apoptosome‚ 87 apoptotic protease activating factor-1 (Apaf-1)‚ 87 inhibition by Hsp27‚9 0 procaspase 9 activation‚ 87 intermembrane protein release‚ 86‚ 89-90 cardiolipin‚ 89 cytochrome c‚ 89-90 PI3K/Akt/PKB‚ 88-93 ursodeoxycholic acid‚ 94‚ 126 VDAC/permeability transition pore‚ 86‚ 91-92; see also Mitochondrial energetics‚ VDAC tumor necrosis factor receptor (TNFR) family‚ 85‚ facing p. 85‚ 86‚ 93-95; see also Proinflammatory cytokines Fas‚ Fas ligand (FasL)‚ 85‚ 86‚ 92-95 Fas mediation of butyrate-induced apoptosis‚ 123 FasL-independent Fas signaling‚ 93-95 AMPK‚ 94-95 ATP‚ ROS‚ 94 bile acids‚ menadione‚ UVB light‚ 93-94 TNFR1.85‚ 86 PI3K/Akt/PKB-mediated signaling‚ 86‚ 91‚ 129‚ 185‚ 187‚ 188‚ 240‚ 287 Apoptotic protease activating factor-1 (APAF-1)‚ 87; see also Apoptosis‚ apoptosome Appetite: see Feeding behavior Arterial hypertension‚ 154 Aspirin: see NSAIDs‚ salicylate Astrocyte; see also Alzheimer disease antioxidant defenses‚ 229‚ 231‚ 232‚ 239‚ 262‚ 266‚ 268‚ 326 fatty acid binding protein‚ expression‚ 262 fatty acid oxidation‚ 207-208‚ 210‚ 211‚ 262-264 gluconeogenesis‚ 7‚ 211-212‚ 255-258‚ 324

371

372

Index

Astrocyte (cont.) glutamate‚ 255‚ 257‚ 259-262‚ 265‚ 268 glutamine export‚ glutamine shuttle‚ 259-262 ketogenesis‚ 6‚ 155‚ 208‚ 211‚ 212‚ 263-268‚ 325‚ 327‚ 364 cannabinoid effects‚ 264 lactate export‚ 212‚ 255-258 Atherosclerosis ABCA1‚ 151 C-reactive protein‚ 151-153 CD36 scavenger receptor‚ 150-151 cholesterol‚ 150-153 endothelial lesion‚ 150-154 fatty acid binding protein (aP2)‚ 151 fatty acid oxidation‚ 147-154 leptin‚ 151 trans fatty acids‚ 8‚ 144‚ 150‚ 151‚ 157‚ 362 peroxisome proliferator-activated receptors‚ 82‚ 84‚ 151-153; see also hypolipidemic drugs fibrate peroxisome proliferators‚ 154 “statin” agents‚ 152-154 antiinflammatory‚ antiproliferative effects‚ 15‚ 152-154 fatty acid oxidation‚ 153 HMG CoA reductase‚ 152-153 LXR‚ 55‚ 92‚ 152‚ 154 SREBP‚ 152-154 oxidized LDL‚ 149-150 Bipolar disorder: see Depression Brain substrate utilization fatty acids: see also Feeding behavior adverse effects on astrocyte‚ neuron‚ 263-268 astrocyte utilization and oxidation‚ 226-232; see also Astrocyte ketogenesis‚ 6‚ 208‚ 211‚ 230‚ 263 MCFA‚ SCFA oxidation‚ 230 ROS generation‚ antioxidant defense‚ 231‚ 264-266 flow-independent uptake‚ 213‚ 226 neuronal SREBP‚ 232 PPAR expression and activity‚ 231 glucose flow-dependent uptake‚ 213 glucose transporters‚ GLUT 1‚ GLUT 3‚ 218 GLUT 3 adducts‚ 309‚ 324; see also Alzheimer disease‚ cerebral metabolism glycolysis‚ 218 insulin‚ insulin receptor‚ 218‚ 235‚ 238‚ 309‚ 311‚ 315‚ 321‚ 322‚ 328 suppression by ketone bodies‚ 221‚ 222 ketone bodies flow-independent uptake‚ 213‚ 221‚ 310 in brain‚ 207‚ 208‚ 212‚ 220-225‚ 232‚ 261‚ 263‚ 264‚ 267‚ 268‚ 277-279‚ 282‚ 292‚ 309-314‚ 324‚ 326‚ 327

Index

373

Brain substrate utilization (cont.) fasting-independent‚ 220-221 inhibition of PDH‚ glycolysis‚ 222-223 neuronal energetics‚ 6‚ 207‚ 208‚ 223-225 neuronal utilization in preference to glucose‚ 207‚ 208‚ 221-225 short term neuroprotection‚ 223-225 uncoupling‚ uncoupling-like effects‚ 122‚ 123‚ 129‚ 208‚ 211‚ 222-224‚ 231‚263‚ 282‚ 324 ROS generation‚ 208‚ 222-224‚ 264-268‚ 282‚ 292‚ 324 synaptic function‚ 224-225 neuroprotection vs. acute insults‚ 223-225 BRCA1 and acetyl CoA carboxylase‚ 55 Butyrate antiproliferative effects‚ 121-124‚ facing p. 123 apoptosis‚ 123-124 121 colonic epithelium‚ 124 fatty acid oxidation and ketogenesis‚ 122-124 uncoupling‚ uncoupling-like effects‚ 122-123‚ 129‚ 222-223 C75‚ 237; see also Feeding behavior E-Cadherin‚ 29‚ 32‚ 121‚ 125‚ 315‚ 318 P-Cadherin‚ 125 protein kinase II (CaMKII)‚ 287 Calcium stimulation of mitochondrial dehydrogenases‚ 61‚ 287‚ 288; see also Mitochondrial energetics Caloric restriction‚ 14‚ 146‚ 149‚ 328 Cancer‚ cancer cells‚ 15‚ 23-27‚ 29-31‚ 45‚ 47‚ 84‚ 365-367 cancer cachexia‚ 15‚ 47‚ 50‚ 52‚ 61-64 colon cancer butyrate‚ 124 cell lines‚ 51‚ 84‚ 122‚ 127 growth inhibition by MCT‚ ketone bodies‚ 49-50 salicylate‚ NSAID effects‚ 127 transplantable MAC-13/16‚ host and tumor metabolism‚ 49‚ 52‚ 61‚ 62 fatty acid biosynthesis‚ 53-56 Cannabinoids‚ 59‚ 237‚ 292‚ 293‚ 365; see also Feeding behavior cannabinoid receptor CB1‚ 59‚ 292 endocannabinoids‚ 292‚ 293‚ 365 anandamide‚ 292 fatty acid amide hydrolase‚ 292 tetrahydrocannabinol‚ 264‚ 292‚ 293‚ 365 Cardiac myocyte adverse effects of ketone bodies‚ 223 fatty acid oxidation‚ 7‚ 58‚ 96‚ 97‚ 125‚ 151‚ 154‚ 232‚ 234 and mitochondria‚ 125 oxidative stress‚ 154-155 Carnitine palmitoyltransferase-I (CPT-I): see Fatty acid oxidation see also PI3K/Akt/PKB signaling butyrate‚ 121

374

Index

(cont.) caspase 3‚ 95 colonocyte‚ 30‚ 33‚ 84‚ 315 liver regeneration‚ 180 neuron‚ 32‚ 33‚ 306‚ 315-318‚ 320 phosphorylation by 28‚ 29‚ 31‚ 33‚ 315-320 transcriptional regulation‚ 27-35‚ 92‚ 315-320 84-85 125 C. elegans‚ 29 Cell cycle apoptosis and‚ 95-98‚ 319-320 energy (ATP) demands‚ 5‚ 6‚ 8‚ 13-15‚ 22-25‚ 30-32‚ 59‚ 361‚ 363‚ 364‚ 366-367 inhibition butyrate‚ 121-122 cyclin-dependent kinase inhibitors Cip/Kip family 29-30‚ 95-98‚ 366 29 121 Forkhead transcription factors‚ 29 salicylate‚ NS AIDs‚ 127 125 PI3K/Akt/PKB signaling‚ 27-31; see also PI3K/Akt/PKB signaling inhibition by fatty acids‚ 31‚ 83‚ 86‚ 144‚ 145 ROS‚ 33-35 Cell proliferation; see also Cell cycle; and Hypoxia; and Mitochondrial energetics hypoxic‚ 14‚ 19‚ 26‚ 27‚ 30‚ 34-35‚ 50‚ 77‚ 81‚ 91‚ 93‚ 128‚ 158‚ 264‚ 361-363 requirements for model‚ 14-16 substrates aerobic glycolysis and citric acid cycle‚ 14‚ 16‚ 20-23‚ facing p. 21‚ 25‚ 26‚ 32‚ 35‚ 5558 fatty acids‚ 53-57 exclusion of fatty acid oxidation‚ 19‚ 59‚ 81-83‚ 144-145‚ 160‚ 181-184 fatty acid biosynthesis‚ 53-55 liver fatty acid binding protein‚ 49 81-83 glutamine‚ glutamate‚ 58‚ 124‚ 148 sterols‚ isoprenoids‚ 55-56‚ 363 Ceramide‚ 92-94‚ 144‚ 188‚ 239‚ 292 Cholesterol: see Alzheimer disease; and Atherosclerosis Citric acid cycle (tricarboxylic acid cycle): see Cell proliferation‚ substrates; and Mitochondrial energetics effect of fatty acids‚ 53 NSAIDs‚ 127‚ 129-131 and 84-85 COX: see Cyclo-oxygenase CPT-I (Carnitine palmitoyltransferase-I): see Fatty acid oxidation Crabtree effect‚ 27‚ 148; see also Hyperglycemia C-reactive protein (CRP)‚ 151-153; see also Atherosclerosis

Index

375

Cyclin D and PI3K/Akt/PKB‚ 28-30‚ 156 Cyclo-oxygenase (COX‚ prostaglandin G/H synthase)‚ 28‚ 53‚ 84‚ 85‚ 187‚ 240‚ 289; see also NSAIDs Cytochrotne c: see Apoptosis‚ mitochondrial function; and Mitochondrial energetics Cytokeratins 8 and 18; see also Fatty acid oxidation‚ regulation of CPT-I AMPK and CPT-I regulation‚ 54‚ 57-58‚ 95 caspase 3‚ 95 heat shock proteins‚ 156‚ 157 Cytokines: see Proinflammatory cytokines Dementia: see Alzheimer disease; and HIV‚ dementia Depression‚ 239‚ 320-321‚ 365 Docosahexaenoic acid (22:6‚ (DHA): see Fatty acids Diabetes mellitus‚ 48‚ 80‚ 81‚ 126‚ 154‚ 155‚ 306‚ 362‚ 364 atherosclerosis‚ 147‚ 148‚ 150‚ 152 brain‚ 208‚ 224‚ 239‚ 307-310‚ 313‚ 323‚ 324‚ 327‚ 329‚ 362 cancer cachexia‚ 62‚ 366 diffuse organ injury‚ 126‚154‚ 155‚ 362‚ 365 plasma FFA and insulin resistance‚ 6‚ 8‚ 83‚ 143-148‚ 154‚ 208‚ 239‚ 264‚ 279‚ 292‚ 310‚ 324‚ 327‚ 329‚ 362-365 Dicarboxylic fatty acids: see Fatty acids Eicosanoids‚ 14‚ 47‚ 53‚ 55‚ 57‚ 64‚ 80‚ 84‚127‚131‚179‚187‚269‚289 Eicosapentaenoic acid (EPA): see Fatty acids Electron “leakage” and ROS generation: see Mitochondrial energetics; and Reactive oxygen species Electron transport chain: see Mitochondrial energetics Endothelium: see Atherosclerosis‚ endothelial lesion Enterocyte‚ 32‚ 148‚ 150‚ 260 Epidermal growth factor (EGF)‚ 22‚ 25‚ 28‚ 30‚ 32‚ 61‚ 127‚ 177 Epithelial restitution‚ 32‚ 361 Estrogens‚ 317‚ 325‚ 330 Ethanol‚ protective effects‚156-157‚ 309‚ 325‚ 330 adverse effects‚ 94‚ 147 Excitotoxicity: see Synaptic Transmission Extramitochondrial fatty acid oxidation: see Fatty acid oxidation Familial adenomatous polyposis (FAP): see Adenomatous polyposis coli (APC) Fas‚ FasL: see Apoptosis Fatty acid amide hydrolase‚ 292; see also Cannabinoids Fatty acid binding proteins (FABP)‚ 81‚ 226‚ 231-233‚ 262‚ 292 adipocyte lipid binding protein (ALBP‚ aP2)‚ 82‚ 151‚ 185‚ 238 astrocyte expression (brain‚ heart‚ keratinocyte‚ liver isoforms)‚ 262 brain isoform (B-FABP)‚ 233‚ 262 cutaneous isoform (C-FABP)‚ 53 heart isoform (H-FABP)‚ 262 keratinocyte isoform (K-FABP)‚ 262 liver isoform (L-FABP)‚ 49‚ 78‚ 180‚ 185‚ 262 cell proliferation‚ 49‚ 180‚ 185 fatty acids‚ 49

376

Index

Fatty acid binding proteins (FABP)‚ (cont.) 78‚ 180 Fatty acid biosynthesis‚ 4‚ 20‚ 22‚ 25‚ 55‚ 78‚ 152-154‚ 232‚ 258‚ 363; see also Acetyl CoA carboxylase; and Lipogenesis fatty acid synthase (FAS)‚ 49‚ 53‚ 153‚ 237 Fatty acid oxidation; see also AMPK; and Brain substrate utilization; and Feeding behavior; and Reactive oxygen species; and adverse effects‚ various tissues‚ 143-160 cell growth‚ 5‚ 47‚ 49‚ 51‚ 53-55‚ 57-59‚ 62‚ 77‚ 361 growth inhibitor effects‚ 121‚123‚ 125-130 interaction with Bcl-2‚91 liver regeneration‚ 180-182‚ 184-189 mitochondria‚ carnitine palmitoyltransferase-I (CPT-I)-mediated‚ 7-8‚ 21-22 fatty acid override of‚ 21‚ 59‚ 83‚ 144‚ 145‚ 148‚ 181-182‚ 186‚ 212‚ 231‚ 238‚ 239‚ 362 long chain fatty acids‚ 20‚ facing p. 21‚ 22 medium‚ short chain fatty acids‚ 57; see also Fatty acids fatty acids‚ 51‚ 52‚ 57 regulation of mitochondrial fatty acid oxidation activation by AMPK‚ 57‚ 58‚ 94 cytokeratins 8/18 (malonyl CoA-independent)‚ 54‚ 57‚ 58‚ 95‚ 156‚ 157‚ 187 inhibition by fibrate peroxisome proliferators‚ 13‚ 49‚ 78‚ 80-82‚ 94‚ 151‚ 154‚ 180‚ 185 leptin‚ 58 malonyl CoA-dependent‚ 20‚ 22‚ 51‚ 54‚ 57‚ 58‚ 92‚ 154‚ 185‚ 186‚ 228‚ 232‚ 237‚ 238‚ 328‚ 363 malonyl CoA decarboxylase‚ 59 peroxisomes‚ 7‚ 8‚ 51-52‚ 54‚ 78-80‚ 83‚ 144‚ 182 ROS generation‚ 54‚ 83‚ 144‚ 148‚ 150‚ 151‚ 155‚ 182‚ 264‚ 362 smooth endoplasmic reticulum‚ 78-80‚ 144‚ 148‚ 180‚ 182‚ 232 Fatty acids adverse effects on mitochondrial function‚ 143-145 cell growth‚ 47-64 dicarboxylic fatty acids‚ peroxisome proliferation‚ 78‚ 180‚ 232 medium chain fatty acids (MCFA)‚ 49‚ 57‚ 79‚ 143‚ 230 oleic acid‚ 262 omega-3 unsaturated fatty acids brain‚ 262‚ 286‚ 327 cell growth‚ 15‚ 47‚ 52-53‚ 57‚ 160‚ 187‚ 361‚ 367 metabolic effects‚ 6‚ 51-52 antioxidant‚ uncoupling effects‚ 6‚ 15‚ 51‚ 53‚ 129‚ 131‚ 144‚ 152 fatty acid oxidation‚ 6‚ 47‚ 51-52‚ 143‚ 145 glycerolipid/lipoprotein synthesis‚ 51-52 omega-6 unsaturated fatty acids‚ 47-50‚ 55‚ 57 brain‚ 262‚ 327 cell growth‚ 47-50‚ 52‚ 53‚ 55‚ 57‚ 179‚ 180‚ 187‚ 189‚ 361 metabolism‚ 51‚ 52 ROS generation‚ 48‚ 55‚ 57 PI3K/Akt/PKB activation‚ 48 saturated fatty acids‚ 49‚ 144‚ 309

Index Fatty acids‚ (cont.) short chain fatty acids (SCFA)‚ 50‚ 57‚ 230‚ 318‚ 319‚ 361; see also Butyrate trans fatty acids‚ 8‚ 144‚ 150‚ 151‚ 157‚ 362 Feeding behavior‚ hypothalamic regulation‚ 6-7‚ 233-237‚ 365; see also Cannabinoids cannabinoids‚ 237 fatty acid oxidation‚ 233-236 fatty acid synthase and C75‚ 237 leptin and insulin effects‚ 234-236 relationship to plasma [FFA]‚ 234-235 Fibrate hypolipidemic agents: see Atherosclerosis; and Fatty acid oxidation‚ CPT-I; and Forkhead transcription factors‚ 28‚ 29‚ 33‚ 34‚ 84‚ 93‚ 158 Free (unesterified) fatty acids (FFA); see also Diabetes mellitus; and Fatty acid oxidation‚ CPT-I override; and Human immunodeficiency virus; and Insulin resistance advanced glycation products‚ 147-148‚ 151 brain: see Brain substrate utilization effects in various tissues‚ 143-160 endothelial lesion of atherosclerosis‚ 148-154 nicotine‚ 148‚ 155‚ 239‚ 328‚ 329‚ 365 visceral adipose tissue‚ 148‚ 179 Fructose-2‚6-bisphosphate (-phosphatase) [F2‚6BP(ase)]‚ 20‚ 21‚ 25; see also Gluconeogenesis; and Glycolysis Gamma-aminobutyric acid (GABA) mitosis‚ 319 neurotransmission‚ 259‚ 262‚ 282‚ 284‚ 319 Germ-free animals‚ 124‚ 185 Glia: see Astrocyte Gluconeogenesis‚ 7‚ 21‚ 22‚ 145-148‚ 313; see also Astrocyte cancer cachexia‚ 50‚ 62‚ 64 growth regulatory factor effects‚ 25‚ 126 kidney‚ 7‚ 62‚ 64‚ 179‚ 182 liver regeneration‚ 16‚ 50‚ 177-182‚ 184‚ 187‚ 189 Glucose transporters (GLUT 1‚ 3‚ 4)‚ 23‚ 30‚ 34‚ 55‚ 218‚ 309‚ 324‚ 366 ROS-induced GLUT 3 adducts‚ 309‚ 324 Glutamate; see also Synaptic transmission; and Mitochondrial energetics‚ VDAC impairment of cystine uptake and glutathione balance‚ 91‚ 283‚ 285-286‚ 288-289 neurotransmission‚ 256‚ 259‚ 282-293 postsynaptic neuronal uptake: see Synaptic transmission respiratory substrate and source of ROS‚ 54‚ 124‚ 148; see also Cell proliferation‚ substrates ROS inhibition of uptake by astrocyte‚ 265 Glutamine and intestinal epithelium‚ 124; see also Astrocyte Glutamine export‚ shuttle: see Astrocyte Glutathione and apoptosis‚ 91 Glycogen storage disease‚ type I‚ 13‚ 14‚ 189 Glycogen synthase see PI3K/Akt/PKB signaling Glycolysis aerobic‚ and cell proliferation‚ 14‚ 16‚ 20‚ facing p. 21‚ 23‚ 25‚ 26‚ 32‚ 35‚ 55-58 anaerobic‚ 20

377

Index

378 Glycolysis‚ (cont.) neuronal activation‚ 209‚ 210‚ 212-214‚ 218 (Glycogen synthase see PI3K/Akt/PKB signaling

Heart: see Cardiac myocyte Heat shock factor (HSF1)‚ 156 Heat shock proteins (Hsp27‚ Hsp70‚ Hsp90) attenuation of effect (Hsp70)‚ 157 induction by ethanol‚ thermal stress (Hsp27‚ Hsp90)‚ 156-157 inhibition of apoptosis (Hsp27‚ Hsp90)‚ 90 interaction with intermediate filaments (Hsp27‚ Hsp90)‚ 156 ischemic preconditioning (Hsp27‚ Hsp90)‚ 156 stabilization of Akt/PKB (Hsp27‚ Hsp90)‚ 34‚ 93‚ 156‚ 157 Hep G2 cells‚ 53‚ 153 Hepatectomy: see Liver regeneration Hepatic tumors‚ 127; see also Glycogen storage disease‚ Type I Hepatitis: see Apoptosis‚ cell cycle; and Nonalcoholic steatohepatitis Hepatocellular carcinoma‚ 28‚ 62‚ 79‚ 155 Hepatocyte growth factor (HGF) activation of adipocyte lipolysis‚ 55‚ 177-178 glycolysis‚ 25 liver regeneration‚ 177 PI3K/Akt/PKB‚ 28‚ 31‚ 55‚ 89 SREBP signaling‚ 55 Hexokinase glycolysis‚ 20‚ 23 hexokinase II‚ 23; see also Mitochondrial energetics‚ membrane contact sites‚ VDAC activation by PI3K/Akt/PKB‚ 27‚ 30‚ 89‚ 92‚ 157 cell proliferation‚ 23-27‚ 30‚ 366 activation‚ 26 inhibition of Bax-induced cytochrome c release‚ 24‚ 27‚ 89 neuronal activation‚ 209-211‚ 213‚ 216‚ 218-219 VDAC‚ 22-24‚ 218-219 fatty acid-induced dissociation‚ 59-60‚ 144 kinetic advantages‚ 24 Hexose monophosphate shunt (pentose phosphate pathway)‚ 20‚ 22‚ 25‚ 158‚ 184‚ 239‚ 255‚ 313 Hippocampus‚ 228‚ 306‚ 310 HMG CoA reductase: see CoA reductase Human immunodeficiency virus (HIV) infection‚ 94‚ 240 apoE4‚ Alzheimer disease‚ 306‚ 327‚ 330‚ 365; see also Alzheimer disease‚ apoE dementia‚ 208‚ 267-268 proinflammatory cytokine activation‚ 267-268; see also Liver regeneration‚ proinflammatory cytokines fatty acid metabolism‚ 6‚ 8‚ 9‚ 144‚ 145‚ 208‚ 238‚ 239‚ 264‚ 267-268‚ 279-280‚ 292‚ 365 Hydrogen peroxide see Reactive oxygen species (ROS)

see Ketone bodies CoA reductase (HMG CoA reductase)‚ 57‚ 152-153 4-Hydroxynonenal (HNE)‚ 324

Index

379

Hypercholesterolemia: see Alzheimer disease‚ cholesterol; and Atherosclerosis‚ cholesterol Hyperglycemia‚ 126‚ 147‚ 148‚ 151‚ 155‚ 310 advanced glycation endproducts (AGE)‚ 147-148‚ 151 Crabtree effect‚ 27‚ 148 Hypoxia‚ 14‚ 19‚ 26‚ 27‚ 34-35‚ 50‚ 77‚ 81‚ 91‚ 93‚ 158‚ 181‚ 264‚ 291 angiogenesis‚ 26‚ 34‚ 59‚ 84‚ 361 hypoxia-inducible hydrocarbon nuclear translocator 26-27‚ 34-35‚ 81‚ 93‚ 158‚ 362 ROS activation of PI3K/Akt/PKB‚ 34‚ 128‚ 362‚ 363 vascular endothelial growth factor (VEGF)‚ 34‚ 93‚ 362‚ 366 von Hippel-Lindau (VHL) protein‚ 34-35 Insulin‚ 6‚ 8‚ 28‚ 129‚ 362‚ 364‚ 365; see also Diabetes mellitus; and Feeding behavior; and Insulin resistance apoptosis‚ 89‚ 97 brain‚ 218‚ 222‚ 223‚ 234-236‚ 238‚ 239‚ 255‚ 264‚ 279‚ 287‚ 292 Alzheimer disease‚ 306-311‚ 313-318‚ 320-324‚ 327-330 cancer cachexia‚ 62‚ 63 growth regulation‚ 22‚ 25‚ 27‚ 28‚ 30-34‚ 48‚ 49‚ 51‚ 54‚ 55‚ 57‚ 58‚ 81‚ 83‚ 160 liver regeneration‚ 177‚ 179‚ 180‚ 182‚ 185-189 contrast with 187‚ 189 Insulin-degrading enzyme (IDE) and Alzheimer disease‚ 306‚ 322‚ 323 Insulin-like growth factor-1 (IGF-1)‚ 27‚ 28‚ 30‚ 94‚ 188‚ 218‚ 317 IGF-2‚62 Insulin receptor‚ 25‚ 185‚ 188‚ 218‚ 235‚ 238‚ 309‚ 311‚ 315‚ 321‚ 322‚ 328 Insulin receptor substrate-1 (IRS-1)‚ 23‚ 28‚ 30‚ 89‚ 315‚ 316‚ 321 expression in early hepatocellular carcinoma‚ 25 PI3K/Akt/PKB signaling‚ 28‚ 30‚ 89‚ 315‚ 316‚ 321 Insulin resistance; see also Alzheimer disease; and Diabetes mellitus; and Obesity plasma FFA‚ 6‚ 8‚ 144‚ 145‚ 147‚ 148‚ 152‚ 154‚ 208 visceral adipose tissue‚ 148‚ 179 Insulin secretion‚ 14‚ 27‚ 62‚ 154‚ 155‚ 364 Interleukin-1‚ -6 (IL-1‚ -6): see Proinflammatory cytokines Intermediate filaments: see Cytokeratins 8‚ 18 Ischemia/reperfusion injury‚ 8‚ 15‚ 86‚ 155-157‚ 362 Ischemic preconditioning‚ 156‚ 157‚ 362 Islet cell: see Pancreatic islet cell Isoprenoids (farnesyl-‚ geranyl-geranyl-pyrophosphate)‚ 22‚ 55‚ 57‚ 64‚ 152-154‚ 158‚ 232‚ 366 Ketogenesis: see Ketone bodies see Cell proliferation‚ substrates; and 2-Oxoglutarate; and Synaptic transmission Ketone bodies (acetoacetate‚ see also Alzheimer disease‚ cerebral metabolism; and Astrocytes; and Brain substrate utilization cell proliferation‚ 48-50‚ 123‚ 130‚ 145‚ 268 ketogenesis‚ 21‚ 83 decrease in regenerating liver‚ 50‚ 181‚ 185 mitochondrial fatty acid oxidation‚ 6‚ 21‚ 22‚ 58‚ 59‚ 121‚ 123‚ 126‚ 130‚ 145‚ 211‚ 230‚ 263

380

Index

Ketone bodies (cont.) mitochondrial oxidation ROS‚ oxidative stress generation‚ 123‚ 129‚ 208‚ 222-224‚ 264-268‚ 282‚ 292‚ 324 uncoupling‚ uncoupling-like effects‚ 122‚ 123‚ 129‚ 208‚ 211‚ 222-224‚ 231‚ 263‚ 282‚ 312‚ 324 Kupffer cells: see Macrophages Lactate astrocyte gluconeogenesis‚ export‚ 255-259 glycolysis‚ 7‚ 20‚ 25‚ 26‚ 209‚ 213‚ 214‚ 218 ketone body inhibition of hepatocyte uptake‚ 256 neuronal uptake‚ utilization‚ 212‚ 224‚ 256‚ 278‚ 326 Leptin‚ 7‚ 58‚ 97‚ 151‚ 234-237‚ 365; see also Feeding behavior Lipid-mobilizing factor (LMF)‚ 52‚ 61-62; see also Cancer‚ cancer cachexia Lipogenesis; see also Fatty acid biosynthesis cancer cachexia‚ 49‚ 64 cell proliferation‚ 4‚ 5‚ 20-22‚ 53-56‚ 77‚ 78‚ 81‚ 92‚ 158‚ 185-187‚ 238‚ 363 fatty acid oxidation‚ 22‚ 58‚ 232 SREBP regulation‚ 55-56‚ 92‚ 152-154‚ 232 suppression by AMPK‚ 4‚ 57‚ 58 suppression by fatty acids‚ 51 Lipotoxicity‚ 144‚ 155 Lithium 6‚ 19‚ 33‚ 92‚ 208‚ 290‚ 314‚ 320-323‚ 325‚ 330‚ 365 Liver regeneration‚ 14-16‚ 177-189‚ 364 adipocyte lipolysis‚ plasma FFA‚ 177 fatty acids‚ 180 visceral adipose tissue‚ 179 cancer cachexia‚ comparison‚ 47‚ 50‚ 63-64 counterregulatory hormones‚ plasma glucose‚ and insulin‚ 177 fatty acid oxidation extramitochondrial oxidation‚ 180‚ 182 ketogenesis‚ 181 mitochondrial‚ 180-184 180‚ 182 gamma-aminobutyric acid (GABA)‚ 319 gluconeogenesis‚ liver and kidney‚ 178-180 glycolysis‚ 180 liver fatty acid binding protein (L-FABP)‚ 49‚ 180 mitochondrial energetics and redox balance‚ 181-187 uncoupling protein-2‚ 129‚ 187 mitosis‚ acinar gradient‚ 78 DNA synthesis‚ 184 187-189 nitric oxide synthase (NOS)‚ 188 proinflammatory cytokines‚ 78‚ 184-188‚ 239 vs. insulin‚ 187‚ 189 steatosis‚ 184 60 Liver transplantation‚ 182‚ 184

Index

381

Liver X receptor (LXR)‚ 55‚ 92‚ 152‚ 154 Long-term potentiation (LTP): see Synaptic transmission Low density lipoprotein (LDL)‚ 55‚ 149-153‚ 267‚ 305 LRP: LDL receptor-related protein‚ 305 LXR: see Liver X receptor MAC-13‚ MAC-16: see Cancer‚ colon cancer Macrophages atherosclerosis‚ 149-154 brain‚ 267-268 endothelial‚ adipocyte fatty acid binding protein (aP2)‚ 82 fatty acid uncoupling effect‚ 51 ICAM‚ scavenger receptor expression‚ 51 Kupffer cell activation‚ cytokine secretion‚ 78‚ 82‚ 184‚ 185‚ 364 mitochondrial ROS and UCP expression‚ 82‚ 156 PPAR expression: see PPARs Malic enzyme‚ 78‚ 258 Malonyl CoA; see also Acetyl CoA carboxylase decarboxylase‚ 57‚ 59‚ 144 inhibition of CPT-I‚ 20‚ 22‚ 51‚ 63‚ 92‚ 144‚ 363 AMPK vs.‚ 54‚ 56-59‚ 79 atherosclerosis‚ 153-154 brain‚ 228‚ 232‚ 237-238‚ 328 liver regeneration‚ 181‚ 185-186 Medium chain fatty acids (MCFA): see Fatty acids Medium chain triacylglycerols (MCT)‚ antiproliferative effect‚ 49-50 Melanocortin‚ 236 Mesenteric adipose tissue: see Adipose tissue‚ visceral Metabolic syndrome‚ 152 Mitochondrial energetics‚ 21-22‚ facing p. 21‚ 24 calcium stimulation of mitochondrial dehydrogenases‚ 61‚ 287‚ 288 electron transport chain‚ 21-22‚ facing p. 21‚ 24 electron “leakage” and ROS generation‚ 59‚ 127-130; see also ROS inner membrane potential 21‚ facing p. 21‚ 24 membrane contact sites‚ 23‚ 24‚ 59-60‚ 89‚ 144‚ 218-220 oxidative phosphorylation‚ 21 Alzheimer disease‚ 304‚ 310‚ 311‚ 314‚ 316‚ 317 cell proliferation‚ 22-26 cytoplasmic diffusion barriers‚ 23 uncoupling‚ and uncoupling proteins‚ 6‚ 14‚ 22‚ 49‚ 51‚ 53‚ 59‚ 62‚ 82‚ 83‚ 123‚ facing p. 123‚ 127‚ 129-131‚ 144‚ 154-155‚ 157‚ 159‚ 181‚ 211‚ 222‚ 223‚ 231‚ 233‚ 240‚ 286‚ 289‚ 290‚ 362‚ 367 uncoupling-like effects‚ 122‚ 123‚ 129‚ 208‚ 211‚ 222-224‚ 231‚ 263‚ 282‚ 324 voltage-dependent anion channel (VDAC)‚ 23-27‚ 59-60‚ 89‚ 218-220 components and structure‚ 23‚ 24; see also Hexokinase II adenine nucleotide translocase (ANT)‚ 23‚ 24‚ 59‚ 219 disruption by fatty acids‚ 59-60‚ 89‚ 144 modulation by glutamate‚ 219-220‚ 284‚ 291 permeability transition pore opening by fatty acids‚ 59-60‚ 89‚ 144 Mitogen-activated protein kinase (MAPK)‚ 81‚ 317

382

Index

Monocarboxylate carrier‚ 221‚ 224 Muscarinic acetyl choline receptor‚ 305‚ 328 NAD(P)‚ NAD(P)H: see related processes NADPH oxidase and ROS‚ 34‚ 154 Neural progenitor cells‚ 268-269 Neuritic plaques‚ 304‚ 305 Neurodegenerative disease‚ 6‚ 15‚ 149‚ 239‚ 267‚ 280‚ 291‚ 292‚ 307‚ 308‚ 321‚ 322‚ 328‚ 330‚ 362‚ 365 Neuronal activation‚ 6‚ 8‚ 31‚ 32‚ 207-215; see also Alzheimer disease; and Brain substrate utilization energetics‚ 207-214 oxidative phosphorylation‚ 214‚ 218‚ 304‚ 310‚ 311‚ 314‚ 316‚ 317 glucose utilization and citric acid cycle‚ 207-214‚ 218-220 ketone bodies‚ 220-225 cerebral blood flow (CBF) and glucose uptake‚ 207‚ 208‚ 213; see also Alzheimer disease glucose-oxygen dissociation‚ 208‚ 225‚ 278-279‚ 314 substrate efficiency‚ 278‚ 314 Neuropeptide Y‚ 236‚ 237 see Nuclear factor Nicotine‚ nicotine receptor‚ 148‚ 155‚ 239‚ 322‚ 328-329‚ 365; see also Alzheimer disease N-methyl-D-aspartate (NMDA)‚ 282‚ 286-290‚ 292; see also Synaptic transmission Nitric oxide (NO)‚ 86‚ 94‚ 125‚ 158‚ 182‚ 187‚ 188‚ 213‚ 239‚ 240‚ 266‚ 313 Nitric oxide synthase (NOS)‚ 57‚ 86‚ 125‚ 129‚ 131‚ 187‚ 188‚ 238-240‚ 289 Nonalcoholic steatohepatitis‚ 15‚ 94‚ 150‚ 155 Nonsteroidal antiinflammatory drugs (NSAIDs)‚ 14‚ 85‚ 93‚ 121‚ 127-131‚ 143‚ 145‚ 160‚ 320‚ 325‚ 328‚ 330 antiproliferative effects‚ 127-131 colon tumors‚ 127 COX2‚ eicosanoid independence‚ 127‚ 129-131‚ 320‚ 325 insulin‚ 129 mitochondrial effects‚ facing p. 123‚ 127-131 electron transport chain inhibition‚ 130 uncoupling‚ antioxidant effects‚ 127-131 and AP-1 activation‚ 128-131 salicylate‚ 15‚ 121‚ 127-131‚ 145‚ 240‚ 290‚ 325‚ 367 sulindac‚ 130 Nuclear factor activation by 86‚ 187‚ 188 AP-1‚ 34‚ 48‚ 129 excitotoxicity‚ 289‚ 290 liver regeneration‚ 187-189 ROS‚ 34‚ 48‚ 51‚ 82‚ 94‚ 126‚ 129-131‚ 152‚ 157‚ 158 salicylate‚ 129-130‚ 290 survival/protective effects‚ 28‚ 34‚ 129‚ 199 neuroprotection‚ 240‚ 325‚ 362 125

Index

383

Obesity‚ 6‚ 8‚ 31‚ 83‚143-145‚ 148‚150-152‚ 154‚ 155‚ 185‚ 208‚ 235‚ 237-239‚ 264‚ 279‚ 288‚ 292‚ 327-329 365; see also Adipose tissue‚ visceral; and Diabetes mellitus; and Insulin resistance Oxaloacetate‚ 256‚ 258‚ 259 Oxidative phosphorylation: see Mitochondrial energetics Oxidative stress: see Reactive oxygen species (ROS) Oxidized LDL: see Atherosclerosis 2-Oxo-glutarate‚ 26‚ 35‚ 58‚ 124‚ 146‚ 148‚ 259‚ 278‚ 284‚ 285‚ 289‚ 291; see also Cell proliferation‚ substrates; and Mitochondrial energetics; and Synaptic transmission 2-Oxo-glutarate dehydrogenase (2-OGDH)‚ 61‚ 83‚ 305‚ 310 Oxysterols‚ 55‚ 152 P450 2E1‚ 182 P450 4A‚ 78‚ 180‚ 182‚ 232 P450 4X1‚ 232 p53‚ 26-28‚ 90-93; see also Apoptosis‚ mitochondrial function apoptosis‚ 26‚ 27‚ 90-93‚ 96‚ 97‚ 121‚ 125‚ 319‚ 320‚ 366 cell cycle‚ 90-93‚ 96‚ 97‚ 121‚ 125‚ 319‚ 366 mitochondria‚ 90-93‚ 145 telomerase‚ 91-93 Pancreatic islet cell‚ 14‚ 27‚ 154‚ 155‚ 234‚ 235‚ 364 Pentose phosphate pathway: see Hexose monophosphate shunt Peroxisome proliferator-activated receptor 78-83‚ 92‚ 122 activator ligands‚ 49‚ 78-83‚ 130‚ 151‚ 180‚ 232 antioxidant effects‚ 77‚ 81-83‚ 145 apoptosis‚ 79-80‚ 94 atherosclerosis‚ 82‚ 84‚ 151‚ 153 COX2‚ prostaglandins‚ NSAIDs‚ 130-131 dicarboxylic fatty acids‚ 78‚ 180‚ 232 fatty acid oxidation‚ 51‚ 55‚ 63‚ 78-83‚ 92‚ 122‚ 145‚ 159 peroxisomes and SER‚ 7‚ 8‚ 51‚52‚ 59‚ 63‚ 78‚ 80‚ 83‚ 122‚ 144‚ 148‚ 150‚ 151‚ 155‚ 182‚ 230-232‚ 266‚ 362 mitochondrial ROS generation‚ 145-148; see also Fatty acid oxidation‚ mitochondria; and Mitochondrial energetics fibrates and rodent hepatocarcinogenesis‚ 78‚ 79 contrasts vs. regeneration‚ 78‚ 82 hepatocyte proliferation‚ liver regeneration‚ 13‚ 54‚ 78-83‚ 180‚ 182 human liver‚ 83 hypoxia‚ 81 L-FABP‚ 180 MAPK‚ 81 ‚ 188 SREBP‚ 55 tissue and macrophage expression‚ 82‚ 84‚ 231‚ 268 absence from human astrocytes‚ 231‚ 264‚ 268‚ 327 absence from Kupffer cells‚ 82 arterial endothelium‚ 82 82‚ 186 transcriptional regulation‚ 78-83

384

Index

Peroxisome proliferator-activated receptors (cont.) 84-85 activator ligands fatty acids‚ 84 effects‚ 85‚ 291 15dPGJ2‚ thiazolidinediones‚ 85 antiproliferative‚ differentiating effects‚ 84 antitumor effects‚ 84-85 induction by butyrate‚ NSAIDs‚ 84‚ 122‚ 125‚ 126‚ 130 atherosclerosis‚ 84‚ 151-153 155 tissue expression‚ 84‚ 231 85‚ 131‚ 231 Peroxisome proliferators: see Peroxynitrite (ONOO)‚ 158‚ 239‚ 313 Phosphatase and tensin homolog deleted on chromosome ten (PTEN): see PI3K/Akt/PKB signaling‚ PTEN Phosphatidyl-inositol-3-kinase (PI3K): see PI3K/Akt/PKB signaling Phosphoenolpyruvate carboxykinase (PEPCK): see Gluconeogenesis Phosphofructokinase-2 (PFK2)‚ 25‚ 30‚ 180 Phosphoinositide-dependent kinase-1 (PDK1)‚ 27‚ 31 Phospholipase A2‚ 151‚ 289‚ 328 PI3K/Akt/PKB signaling (wg/Wnt-like signaling); see also Alzheimer disease‚ 304‚ 314-325‚ 328 APC‚ 30‚ 31‚ 33‚ 315‚ 316‚ 318‚ 366 axin‚ 30‚ 31‚ 315‚ 316‚ 318 Bcl-2 family‚ 88-93 epithelial restitution‚ 32-33 epithelial-to-mesenchymal transition‚ 125 27-33 HSF1 activation‚ 156‚ 158 inhibition of caspase 3‚ 95 butyrate‚ 121 fatty acids‚ 31‚ 83‚ 86‚ 144‚ 155‚ 188 p53‚ 90-93 PTEN‚ 27‚ 28‚ 31‚ 48‚ 90‚ 97‚ 269‚ 366 ‚ 125-126 leptin‚ 236: see also Feeding behavior lithium 6‚ 19‚ 33‚ 92‚ 208‚ 290‚ 314‚ 320-321‚ 325‚ 330‚ 365 neuronal activation and synaptic function‚ 6‚ 207‚ 285‚ 287-288‚ 290‚ 292‚ 304‚ 316‚ 317‚ 324‚ 325‚ 328; see also Cannabinoids nicotine‚ 155‚ 322‚ 328-329 proliferative and survival effects‚ 5‚ 6‚ 15‚ 27-35‚ 48‚ 54‚ 55‚ 86‚ 90-94‚ 155-158‚ 182‚ 240‚ 269‚ 313-321‚ 361-364‚ 366 signaling by CD40‚ EGF‚ HGF‚ IGF-1‚ Wnt‚ 30-31 86‚ 91‚ 187‚ 188‚ 364 ROS activation of‚ 33-35‚ 158‚ 362‚ 363 stabilization by Hsp27 and Hsp90‚ 93‚ 157

Index

385

PI3K/Akt/PKB signaling‚ (cont.) telomerase‚ 90-93 Placenta‚ 14‚ 26‚ 365 Porin: see Mitochondrial energetics‚ VDAC Portal hypertension‚ 230 Pre-eclampsia‚ 365 Presenilin 1‚ 2 (PS1‚ 2)‚ 31-32‚ 306‚ 315-319‚ 324‚ 329 Prion protein (PrP): see Alzheimer disease‚ PI3K/Akt/PKB Procaspase(s): see Apoptosis Programmed cell death: see Apoptosis Proinflammatory cytokines; see also Apoptosis; and Human immunodeficiency virus infection; and Liver regeneration brain‚ 237-240‚ 267-268 interleukin-1 (IL-1)‚ 78‚ 184‚ 185‚ 237-239‚ 340 interleukin-6 (IL-6)‚ 78‚ 184‚ 185‚ 237-239 tumor necrosis factor 28‚ 29‚ 31‚ 55‚ 61-63‚ 78‚ 82‚ 85‚ facing p. 85‚ 86‚ 89‚ 91‚ 92‚ 95‚ 129‚ 145‚ 157‚ 160‚ 184 -189‚ 237-240‚ 287‚ 291‚ 317‚ 328‚ 364 receptor (TNFR)‚ 31‚ 85‚ 86‚ 91‚ 129‚ 185‚ 187‚ 188‚ 240‚ 287 PI3K/Akt/PKB‚ 86‚ 91‚ 187‚ 188‚ 240‚ 287 Prostaglandins (PG)‚ 52‚ 53‚ 83‚ 85‚ 127‚ 130‚ 131 PGD‚ PGE‚ 52‚ 53‚ 130‚ 131 15-deoxy-delta-12‚14-PGJ2‚ 83‚ 85 Protein kinase A (PKA)‚ 15l‚ 287 Protein kinase C (PKC)‚ 31‚ 53‚ 60‚ 317 Proteolysis-inducing factor (PIF)‚ 61-62 p53-upregulated modulator of apoptosis (PUMA): see Apoptosis‚ Bcl-2 family Pyruvate carboxylase (PCB)‚ 256‚ 258‚ 259 Pyruvate dehydrogenase (PDH)‚ 20‚ facing p. 21‚ 22‚ 28-30‚ 32‚ 33‚ 62‚ 81‚ 83‚ 91‚ 120‚ 122‚ 124‚ 130‚ 144‚ 145‚ 148‚ 180‚ 182‚ 209‚ 212‚ 220‚ 222‚ 223‚ 231‚ 256‚ 261‚ 263‚ 264‚ 279‚ 281‚ 282‚ 284‚ 292‚ 310‚ 311‚ 314‚ 317‚ 318‚ 323-325 Pyruvate kinase (PK)‚ increased activity in human cancers‚ 25‚ 180‚ 366 Randle cycle‚ 211‚ 223 Reactive oxygen species (ROS)‚ hydrogen peroxide and superoxide anion see also Hypoxia; and Mitochondrial energetics beneficial/protective effects of low level transient ROS‚ 8‚ 19‚ 33-35‚ 156-159‚ 362-364 ethanol‚ ischemic preconditioning‚ heat stress‚ 156-157 ROS at peak mitochondrial energization‚ 158-159 oxidative stress‚ injurious effects‚ 362-365 apoptosis‚ 89-91‚94‚ 96‚ 124 atherosclerosis‚ 149-154 brain and Alzheimer disease‚ 208‚ 212‚ 220‚ 224‚ 231‚ 233‚ 234‚ 238‚ 240‚ 264‚ 265‚ 267‚ 268‚ 279‚ 282‚ 284-287‚ 289-291‚ 303‚ 304‚ 310-313‚ 319‚ 322-324‚ 327‚ 328 glutamate‚ glutamine‚ 148 hyperglycemia and‚ 147-148 Crabtree effect‚ 27‚ 148 ischemia/reperfusion injury‚ 8‚ 15‚ 86‚ 155-157‚ 362 mitochondria‚ 77‚ 82‚ 127-131‚ 145-148 cardiac myocyte‚ 154-155 fatty acids‚ 5‚ 7‚ 8‚ 14‚ 22‚ 48‚ 55‚ 57‚ 83‚ 144‚ 148-149

386

Index

Reactive oxygen species‚ (cont.) hepatocarcinogenesis‚ 155 liver regeneration‚ 181‚ 182‚ 187 mitigation by uncoupling‚ 51‚ 82‚ 123‚ 129-131‚ 240 nonalcoholic steatohepatitis‚ 155 pancreatic 154-155 pathological consequences‚ 149-160 plasma FFA‚ in obesity‚ insulin resistance‚ type II diabetes mellitus‚ HIV‚ 144‚ 145‚ 148; see also Diabetes mellitus; and HIV; and Insulin resistance; and Obesity peroxisomes‚ 83‚ 144 pyruvate‚ 147 125-126 Retinal neurodegeneration in diabetes mellitus‚ 308; see also Alzheimer disease Retinoblastoma protein (pRb)‚ 96‚ 122‚ 189 Retinoid X 78‚ 84 Rho/Ras/Raf‚ 153 (Raf)‚ 153 and 287 (Ras)‚ 193 (Rho) ROS: see Reactive oxygen species Saccharomyces cerevisiae (S. cerevisiae)‚ 3‚ 4‚ 14‚ 23‚ 57‚ 189 Secretase 306‚ 317‚ 320‚ 323‚ 325 Short chain fatty acids (SCFA): see Fatty acids Smads‚ 93‚ 125 Smoking: see Nicotine Smooth endoplasmic reticulum (SER); see also Atherosclerosis‚ oxidized LDL redox conditions‚ 149-150 fatty acid oxidation‚ 78‚ 144; see also Fatty acid oxidation SNF1‚4‚ 5‚ 57‚ 189 Sterol regulatory element binding protein (SREBP)‚ 28‚ 55‚ 90‚ 92‚ 152-155‚ 232‚ 328‚ 366 “Statin” hypolipidemic agents: see Acetyl CoA carboxylase; and Alzheimer disease‚ cholesterol; and Atherosclerosis‚ hypolipidemic drugs Streptozotocin‚ 23‚ 48‚ 147‚ 309‚ 324 Superoxide anion see Reactive oxygen species Superoxide dismutase (SOD)‚ 29‚ 128‚ 145‚ 151‚ 182‚ 266‚ 286‚ 325 Synaptic transmission‚ 282-293; see also Glutamate; and Mitochondrial energetics glutamate removal from synaptic cleft‚ 282-285 augmented neuronal exposure to glutamate‚ 285-293 long-term potentiation‚ 285-288 AMPAR expression‚ 282‚ 286-288 excitatory postsynaptic current (EPSC)‚ 286‚ 287 PI3K/Akt/PKB‚ 287-288 excitotoxicity‚ 288-293 neuroprotection‚ 289-293 ketone body and cannabinoid effects‚ 292-293 Tau isoform of microtubule-associated protein‚ 32‚ 304‚ 306‚ 315 Telomerase‚ 28‚ 90-93‚ 317 Tetrahydrocannabinol: see Cannabinoids Thiazolidinedione‚ 84‚ 85 Thioredoxin‚ 128

Index

387

Tobacco: see Nicotine Transforming growth 94 Transforming growth antiproliferative‚ pro-apoptotic effects‚ 60‚ 80‚ 84‚ 93‚ 121‚ 125-127‚ 157‚ 366 metabolic effects‚ 125-126 association with mitochondria‚ 125 brain‚ 313‚ 320‚ 324‚ 325 fatty acid and glucose metabolism‚ 25‚ 53‚ 126‚ 145‚ 160 ROS generation‚ 125 signaling‚ 93‚ 125 Triacylglycerol‚ triglyceride (TG)‚ 20‚ 51-53‚ 150‚ 180‚ 182‚ 186‚ 211‚ 238‚ 256 Tumor necrosis and receptor (TNFR): see Apoptosis; and Proinflammatory cytokines Tumorigenesis‚ 13‚ 14‚ 25‚ 33‚ 52‚ 53‚ 84‚ 98‚ 127‚ 130‚ 155‚ 189‚ 268‚ 269‚ 315‚ 318 Ubiquinone‚ 24‚ 128‚ 183‚ 220‚ 224‚ 290 Ultraviolet light: see Apoptosis‚ FasL-independent Fas signaling Uncoupling‚ Uncoupling proteins (UCP)‚ 49‚ 62‚ 82‚ 129‚ 157‚ 233‚ 286‚ 362; see also Mitochondrial energetics‚ oxidative phosphorylation Uncoupling-like effect: see Butyrate; and Ketone bodies; and Mitochondrial energetics‚ oxidative phosphorylation Ursodeoxycholic acid: see Apoptosis‚ FasL-independent Fas signaling Vascular endothelial growth factor (VEGF)‚ 34‚ 93‚ 362‚ 366; see also Hypoxia Very low density lipoprotein (VLDL)‚ 53‚ 149‚ 150‚ 182‚ 186‚ 238‚ 305 “Vicious cycle” in Alzheimer disease pathogenesis: 304‚ 321-326; see also Alzheimer disease Visceral adipose tissue: see Adipose tissue‚ visceral Voltage-dependent anion channel (VDAC): see Mitochondrial energetics von Hippel-Lindau protein (VHL)‚ 34; see also Hypoxia Wingless(wg)/Wnt-like signaling pathway: see PI3K/Akt/PKB signaling Yeast: see S. cerevisiae

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